2.0 literature review -...

7

2.0 LITERATURE REVIEW

An extensive review of literature was carried out prior to the definition of the

problem, as well as before the objectives were specified for the current study, so as to

understand the need for phenol removal from wastewater, the available technologies,

their advantages and disadvantages, importance of bioremediation approach and need

for isolation of new strains from phenol contaminated environments, followed by

critical analysis of each aspect. These literature analyses proved to be beneficial in

problem definition and for setting the objectives of research. Review of literature was

continued, in search of methodologies to be adopted to meet each of the specified

objectives, as well as for comparing the results obtained in the current research with

those obtained by other researchers in the field, wherever applicable. This chapter

summarizes the relevant literature review carried out during the current study.

2.1 Phenol – Characteristics

Phenols are hydroxy compounds of aromatic hydrocarbons that are also called

as carbolic acid, phenic acid, phenylic acid, phenyl hydroxide or oxybenzene (Nair et

al., 2008). It is a white crystalline solid which is soluble in most organic solvents and

has a distinctive odour (ATSDR, 2008; WHO, 1994). The crystals turn pink or red on

exposure to air and light, hastened in presence of alkalinity. Phenol has an acrid smell

and a sharp burning taste, moderately volatile at room temperature (evaporates more

slowly than water) and quite flammable. The detailed properties of phenol are given

in Table 1.

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8

Phenol has one hydroxyl group attached to the benzene ring, as shown in Fig

1, and is the basic structural unit for a variety of synthetic organic compounds.

Figure 1 Structure of phenol

2.1.1 Sources of phenol

The origin of phenol in the environment is from three different sources such as

natural, man-made and endogenous sources. Phenol is released primarily to the air

and water as a result of its manufacture and use, wood burning and auto exhaust.

Phenol mainly enters waters from industrial effluent discharges.

2.1.1.1 Natural sources

Phenol is a constituent of coal tar. During decomposition of organic materials

and forest fires increased phenol levels in environment. It has also been detected

among the volatile components from liquid manure with an average concentration of

30 µg/kg dry weight (RIVM, 1986).

2.1.1.2 Man-made sources

Man-made sources of phenol and other related aromatics are from industrial

wastes of fossil fuel extraction, phenol manufacturing plants, pharmaceutical industry,


9 wood processing industry, pesticide manufacturing plants, petroleum refinery,

petrochemicals, basic organic chemical manufacture; coal refining, tannery and pulp

& paper mills (Kumaran and Paruchuri, 1997).

Table 1.Properties of phenol

Properties Description

Molecular formula C6H5OH

Molecular weight (g/mol) 94.14

Density (g/cm3) 1.072

Water solubility (g/L at 25 ºC) 87

Melting point (ºC) 43

Boiling point (ºC) 181.8

Auto ignition temperature 715 ºC

Molecular diffusivity in water (cm/sec) 6.0 x 10-4

Relative vapour density 3.24 (air=1)

Dipole moment (debyes) 1.450007

Liquid surface tension (dynes/cm) 36.5 @ 55 °C

Excess enthalpy (kJ/mol) [S/D]* 1/8

Acidity constant, pKa(25 oC) 9.90

Molecular diffusivity in water (cm/sec) 6.0 x 10-4

Air-water partition coefficient, Kaw(25 oC) 2.5 x 10-5

Polarizability, Pi 0.89

Excess free energy (kJ/mol) 10

Excess entropy (J/mol K) [S/D]* -9/-2

Fraction in neutral form at pH 7 0.998


10 2.1.1.3 Endogenous sources

Phenol may be the formed from various xenobiotics such as benzene benzyl

alcohols, phenyl acetic acid, chloroform, etc (Pekari et al., 1992) under the influence

of light (Hoshino and Akimoto, 1978).

Cumene route method used to manufacture phenol since the 1960s has been

through the oxidation of 1- methylethylbenzene. Phenolic resins are used as a binding

material in, insulation material, chipboard and triplex, paints and casting sand

foundries. In future, many chemicals including phenol may be produced in relatively

small reactors about the size of a large desktop. One potential micro-reactor to

produce phenol involves the use of a small diameter (2 mm), porous tube of alumina

coated with a layer of palladium metal. A mixture of benzene and oxygen is fed

through the tube and hydrogen gas is passed over the tube, the tube is heated to 150 -

250°C ( Basha et al., 2010).

2.1.2 Applications of phenol

Industrial Use: Phenol is used in many industries but not restricted to

petroleum refineries, gas and coke oven industries, resin manufacturing,

tanneries, explosive manufacture, plastic and varnish industries, textile

industries, smelting and related metallurgical operations etc.

(Mahadevaswamy et al., 1997; Bandyopadhyay et al., 1998; Marrot et al.,

2006; Bodalo et al., 2008; Jayachandran and Kunhi, 2008).

In hospitals and sanitations: Phenol has anti-bacterial and anti-fungal

properties and hence used in the production of slimicides, disinfectants,


11

antiseptics and medicinal preparations such as ear and nose drops,

mouthwashes and sore throat lozenges (ATSDR, 2008).

In pharamaceuticals: Phenol is also a building block for the synthesis of

pharmaceuticals (e.g., aspirin) (Busca et al., 2008).

Cosmetics: Phenol is also used in the preparation of cosmetics including

sunscreens, hair dyes, and skin lightening products.

Agricultural aids: Phenol is used for the manufacture of herbicides and

pesticides.

2.1.3 Phenol -Health hazards

Phenol is highly toxic, corrosive, and mutagenic. It is also known as a

carcinogenic and teratogenic agent, which affects both the environment and human

beings. Phenols are toxic to human beings and effects several biochemical functions.

Phenol may be fatal by ingestion, inhalation, or skin absorption, since it quickly

penetrates the skin and may cause severe irritation to the eyes and the respiratory tract

(El-Naas et al., 2009). Acute exposure of phenol causes central nervous system

disorders, which can lead to coma. Phenol toxicity can lead to a condition known as

hypothermia whose symptoms can include muscular convulsions with significant

reduction in body temperature. Renal damage and salivation may be induced by

continuous exposure to phenol (Nair et al., 2008).


12 2.1.4 Release of phenol into the environment

Man-made phenolic compounds are found in the air, surface water, ground

water, soil and sediment in abundance, due to agricultural and industrial activities

(HazDat, 1998).

2.1.4.1 Air

Phenol is released to the atmosphere from phenol manufacturing and

automobile exhaust (Scow et al., 1981), waste incinerator plant at 0.36ppb, in

cigarette smoke and plastics (Graedel, 1978) and home fires, especially wood-

burning, may contain substantial quantities of phenol (Den Boeft et al., 1984).

2.1.4.2 Water

Phenol releases to water by the most common anthropogenic sources such as

coal tar (Thurman, 1985), waste water from manufacturing industries such as resins,

plastics, fibers, adhesives, iron, steel, aluminum, leather, rubber, and influents from

synthetic fuel manufacturing (Parkhurst et al., 1979), from paper pulp mills (Keith,

1976) and wood treatment facilities (Goerlitz et al., 1985). Other release of phenol

results from commercial use of phenol and phenol containing products, including

slimicides, general disinfectants (Hawley, 1981; Budavari et al., 1989), medicinal

preparations such as ointments, ear and nose drops, cold sore lotions, mouthwashes,

gargles, toothache drops, analgesic rubs, throat lozenges (USEPA, 1980), and

antiseptic lotions (Musto et al., 1977). It has been estimated that 3.8kg/day of phenol

release to seawater from municipal treatment facilities (Crawford et al., 1995).


13 Animal and decomposition of organic wastes are the two natural sources of phenol in

aquatic media.

2.1.4.3 Soil

Phenol is released to the soil during its manufacturing process, loading and

transport when spills occur, and when it leaches from hazardous wastes sites and

landfills (Xing et al., 1994). According to ATSDR, (1998), generally the data on

concentrations of phenol found in soil at sites other than hazardous sites are lacking.

This may be due to a rapid rate of biodegradation and leaching. Phenol can be

expected to be found in soils that receive continuous or consistent releases from a

point source. Phenol that leaches through soil to groundwater spends at least some

time in that soil as it travels to the groundwater. Phenol has been found in

groundwater, mainly at or near hazardous wastes sites.

2.1.5 Phenol Regulations

Presence of phenol at a concentration higher than the standard limit in the

water bodies may cause adverse effects to human beings, animals, plants etc. Phenols

are toxic or lethal to fish at even relatively low levels of 5–25mgL-1 (Kumar et al.,

2005). Table 2 gives an account of phenol level in industries wastewater. Owing to

this, phenol has also been listed as one among the priority organic pollutants by the

US Environmental Protection Agency (Keith and Telliard, 1979). Hence, stringent

regulations have been imposed by various organizations. United State Environmental

Protection Agency (USEPA) has set a water purification standard, i.e. surface water

must contain less than 1.0 g/L phenol (Chung et al., 2003). As per the rules of central


14 pollution control board, India the minimum permissible level for phenol in

environment is 0.1mg/l (Kumaran and Paruchuri, 1997; Nuhoglu and Yalcin, 2005

and Saravanan et al., 2008). The World Health Organization (WHO) recommends the

threshold permissible phenolic concentration of 0.001mg/L in portable waters and

threshold concentration of phenol in drinking water should be below 1.0 g/L. While

Ministry of Environment and Forests (MoEF), Government of India, have set a

maximum concentration level of 1.0 mg/L of phenol in the industrial effluents for safe

discharge into surface waters.

2.2 Conventional methods of phenol removal and its disadvantages

Many procedures have been applied in order to remove phenol from aqueous

streams. Among the most commonly used techniques are adsorption (Adak and Pal,

2009), ion exchange (Caetano et al., 2009), membrane separation (Li et al., 2010),

advanced oxidation process (Mahvi et al., 2007) and solvent extraction (Atlow et al.,

1984). These classical or conventional techniques give rise to several problems such

as unpredictable hazardous compound formation and generation of toxic sludge,

which often require extreme caution in their disposal (Xia and Liyuan, 2002).

However, the costs to set up the required equipment and to operate these processes are

prohibitively high for large-scale treatment (Mahajan, 1985).

2.3 Biological treatment

Bioremediation has already proven itself to be a cost-effective and beneficial

addition to chemical and physical methods of managing wastes and environmental

pollutants. Recently, research for new and innovative technologies has centered on the


15 biological treatment methods. Bioremediation is the use of microorganisms to break

down toxic and hazardous compounds in the environment (Acquaah, 2004).

Bioremediation includes three main processes: (i) transformation or insignificant

alteration of the molecule; (ii) fragmentation or degradation of the molecule to

simpler compounds; and (iii) mineralization or conversion of the complex compound

into simpler ones ( 2 , 2, 2, NH3, CH4, etc.) (Krastanov et al., 2013). It

generally utilizes microbes (bacteria, fungi, yeast, and algae), although higher plants

are used in some applications.

New bioremediation approaches are emerging based on advances in molecular

biology and process engineering. Recently developed rapid-screening assays can

identify organisms capable of degrading specific wastes and new gene-probe methods

can ascertain their abundance at specific sites. New tools and techniques for use of

bioremediation in situ, in biofilters, and in bioreactors are contributing to the rapid

growth of this field (Bonaventura and Johnson, 1997).

2.4 Microorganisms in phenol biodegradation

Bioremediation processes mainly involve the use of microorganisms. For this

reason, the evaluation of polluted areas prior to bioremediation often includes

detection, quantification, and activity determination of the xenobiotic-degrading

microorganisms. The biodegradation activity of microorganisms has become

particularly topical over the past decades with regard to the increased presence of

resistant anthropogenic pollutants in the biosphere in extents exceeding the self-

cleaning abilities of nature (Krastanov et al., 2013). Phenols are metabolized by


16 microorganisms from a variety of different genera and species of bacteria, fungi, yeast

and algae (Table 3).

Table 2. Levels of Phenol Reported in Industrial Wastewaters (Metcalf and Eddy, 2003)

Industrial Source Phenol Concentration (mgL-1)

Petroleum refineries 40 - 185

Textile 100 - 150

Coke ovens (without dephenolization) 600 - 3900

Ferrous industry 5.6 - 9.1

Pulp and paper industry 22

Phenolic resin production 1600

Fiberglass manufacturing 40 - 2564

Petrochemical 200 - 1220

Paint manufacturing 1.1

Phenolic resin 1270 - 1345

Leather 4.4 - 5.5

Coal conversion 1700 - 7000

Rubber industry 3 - 10

Wood preserving industry 50- 953

Algae have the poor relations of the environmental microbiologist, in spite of

their ubiquitous distribution, their central role in the fixation and turnover of carbon

and other nutrient elements, their contribution to eutrophication, and recognition of

their heterotrophic abilities. The biodegradation of phenol by microalgae occurs only

under aerobic conditions, while many phenols show acute toxicity to algae (Shigeoka

et al., 1988). Both cyanobacteria and eukaryotic microalgae are capable of

biotransforming aromatic compounds, including phenols (e.g., Chlorella sp.,


17 Scenedesmus sp., Selenastrum capricornutum, Tetraselmis marina, Ochromonas

danica, Lyngbya gracilis, Nostoc punctiforme, Oscillatoria animalis, Phormidium

foveolamm) (Semple and Cain, 1995; Lika, and Papadakis, 2009), but to date, no

pathways for the oxidative cleavage of the aromatic ring and the assimilation or

mineralization of its products by algae have been elucidated, although there is

evidence that some algae must be capable of this process (Ellis, 1977).

Fungi share a significant part in the recycling of aromatic compounds in the

biosphere and several studies have shown that diverse fungi are capable of phenols

mineralization. They are capable of consuming a wide variety of carbon sources by

enzymatic mechanisms, thus providing possibilities for metabolizing phenols and

other aromatic derivates (Stoilova et al., 2007). The most abundant fungi in polluted

environments are yeasts. Some yeast like Candida tropicalis, Fusarium flocciferium,

and Trichosporon cutaneum are capable of utilizing phenol as the major carbon and

energy source (Agarry et al., 2008).

Rubilar et al., (2008) analyzed the degradation of chlorophenols by white rot

fungi, which are a group of organisms very suitable for the removal of chlorinated

phenolic compounds. They are robust organisms that are tolerant to the presence of

high concentrations of various pollutants, even with a low bioavailability and this

ability is mainly due to their very powerful extracellular oxidative enzymatic system.


18

Table 3. Microorganisms in phenol degradation

Microorganisms References

Bacteria

Acinetobacter radioresistens S13 Mazzoli et al., (2007)

Alcaligenes eutrophus Muller and Babel (1996)

Alcaligenes faecalis Bai et al., (2007)

Bacillus brevis Arutchelvan et al., (2005)

Bacillus cereus Banerjee and Ghoshal (2010)

Bacillus laterosporus Topalova et al., (1995)

Bacillus stearothermophilus Mutzel et al., (1996)

Bacillus thermoleovorans Feitkenhauer et al., (2001)

Burkholderia sp Reardon et al., (2002)

Pseudomonas aeruginosa Jayachandran and Kunhi, (2009)

Pseudomonas cepacia Arutchelvan et al., (2005)

Pseudomonas fluorescens Viggor et al., (2008)

Pseudomonas pictorium Annadurai et al., (2000)

Pseudomonas putida Onysko et al., (2000)

Pseudomonas putida Viggor et al., (2008)

Pseudomonas stutzeri SPC-2 Ahamad and Kunhi (1996)

Pseudomonas stutzeri Jayachandran and Kunhi, (2009)

Rhodococcus erythropolis Margesin et al., (2005)

Sulfolobus solfataricus 98/2 Christen et al., (2011)

S. solfataricus P2 Izzo et al., (2005)

Fungi

Aspergillus niger Garcia et al., (2000)

Aspergillus terreus Garcia et al., (1997), (2000)

Candida tropicalis Adav et al., (2007)

Coprinus cinereus Guiraud et al., (1999)

Coprinus micaceus Guiraud et al., (1999)

Fusarium flociferum Cai et al., (2007)


19

Table 3. - contd..

Fusarium strains FE11, FE16 Santos and Linardi, (2004)

Geotrichum candidum Garcia et al., (1997), (2000)

Graphium sp. Strains LE6, LE11, LA1, LE9, LA5, FIB4, AE2 4

Santos and Linardi, 2004

Penicillium sp. strains AF2, AF4, FIB9 Santos and Linardi, (2004)

Phanerochaete chrysosporium Garcia et al., (2000)

Pleurotus ostreatus Fountoulakis et al., (2002)

Trichosporon cutaneum R57 Shivarova et al., (1999)

Trichosporon cutaneum Alexieva et al., (2008)

Algae & Cyanobacterium

Chlorella vulgaris Shigeoka et al., (1988)

Ochromonas danica Semple and Cain, (1995), (1996)

Phormidium valderianum BDU30501 Shashirekha et al., (1997)

Selenastrum capricornutum Shigeoka et al., (1988)

Many studies were conducted on the basis of the potential of microorganisms

to transform toxic compounds. Through adaptation mechanisms, a number of

microbial species are capable of transforming xenobiotics into compounds that can be

included in the natural exchange of matter. The metabolism of aromatic compounds,

phenol, and its derivatives in particular, is vigorously investigated in prokaryotic

microorganisms (Watanabe et al., 1998). A lot of information is accumulated on

bacterial species from the Pseudomonas genus, which are known for their ability to

utilize diverse aromatic compounds as a single carbon source (Hinteregger et al.,

1992; Kwon and Yeom, 2009; Seker et al., 1997).


20 2.5 Phenol degrading bacteria

Bacteria that are normally used in phenol degradation include Pseudomonas sp

(Monteiro et al., 2000; Gonzalez et al., 2001; Nurdan and Azmi, 2004; Ullhyan and

Ghosh, 2012), Azotobacter sp (Hughes and Bayly, 1983), Rhodococcus sp (Prieto et

al., 2002; Suhaila et al., 2012), Alcaligenes sp (Valenzuela et al., 1997),

Acinetobacter sp (Hao et al., 2002; Hein-aru et al., 2000), Arthrobacter citreus

(Karigar et al., 2006) Alcaligenes faecalis (Jiang et al., 2007). . Many studies on

biodegradation of phenol using pure and mixed cultures have been reported (Collins

et al., 2005; Dursun and Tepe, 2005; Shen et al., 2009; Santos et al., 2009 and

Chakraborty et al., 2010).

Conventionally, the strains are isolated from a contaminated site as it is found

that the probability of phenol resistant strains appears to be higher in such sites as

these strains would have been exposed to higher concentration of phenol and thus

would have acclimatized over continuous exposure of phenol. Alternatively,

acclimatization of the microorganisms is done in the laboratory to overcome the

substrate inhibition problems that normally occurred in phenol biodegradation at high

concentration (Lob and Tar, 2000). Certain intracellular enzymes are induced during

acclimatization stage so that the microbes are available to take part in the reaction

(Kumar et al., 2005). Hao et al., (2002) studied the degradation of phenol by

Acinetobacter species at a concentration of 350 mg L-1.

Recently Corynebacterium glutamicum, an industrial soil microorganism, was

proved to be very effective for the bioremediation of phenol-contaminated soil (Lee et

al., 2010). The authors suggested that a suitable dose of C. glutamicum treatment was


21 sufficient for the large scale remediation of phenol-contaminated soil. Alva and

Peyton (2003) reported the first study of phenol and catechol biodegradation under

combined saline and alkaline conditions by the haloalkaliphilic bacterium Halomonas

campisalis. Haloalkaliphiles are bacteria that thrive in saline and alkaline

environments such as soda lakes. The haloalkaliphilic bacterium H. campisalis

degrades phenol and catechol in alkaline (pH values of 8 - 11) and saline

environments (0 –150gL-1 NaCl).

2.6 Isolation of phenol degrading bacteria from contaminated sites

Microorganisms are an important part of natural ecosystems. They are found

in industrially contaminated soils and waters as a result of their ability to survive in

restrictive conditions. In this respect, their ability to rapidly and efficiently purify the

environment of phenolic contamination is important, with a view to protecting the

living environment and human health directly. The investigations on specificity of

phenol biodegradation by different microbial strains are meaningful for the invention

of effective remediation technologies for industrial wastes where the phenolic

substrates are a common occurrence. Recently the metagenomic approaches for the

analysis of specific catabolic activity is gaining wider brand recognition in the

investigation of the enzyme systems and the capabilities of microorganisms with

pronounced degradation ability and has opened wider prospects for their direct

technological application (Krastanov et al., 2013).

Bioremediation represents an environmental friendly procedure and there is an

increasing interest in isolating and identifying microorganisms with phenol

metabolizing capacity. Phenol degrading microorganisms are usually isolated from


22 phenol polluted environments by the enrichment technique, and the genetic diversity

of the microorganisms may have been altered due to the adaptation to the target

pollutant and the enrichment procedure. Many phenol-degrading microorganisms,

including bacteria, yeast and algae have been isolated from environment, among

which the bacteria are studied extensively (van Schie and Young, 2000; Yang and

Lee, 2007).

Bacteria are omnipresent and it is estimated that only 1% of the bacteria has

been isolated and identified. Thus, there are many bacteria with excellent potential

that are unexplored which can be employed for various applications among which

phenol degrading bacteria are no exceptions. Therefore, the past few decades had led

to discovery of many bacterial strains that have great potential to degrade phenol.

Since, biodegradation has emerged as a low cost and eco-friendly technology, interest

among the researchers has fueled to search for a potent bacterial strain that can resist

high concentration of phenol as well as degrade phenol at a faster rate. According to

Wang et al., (2007) little information on bacteria with a high phenol tolerance and

high metabolizing activity is available. Therefore, there still exits the need to isolate

new phenol degrading bacteria that can grow at elevated concentration of phenol. A

new phenol-degrading bacterium Acinetobacter sp. strain PD12 was isolated from

activated sludge.

Jiang et al., (2004b) isolated 10 bacterial strains from their aerobic phenol-

degrading granules, identified their potential for degrading phenol. Heinaru et al.,

(2000) isolated 39 strains from polluted river water (38 Pseudomonas sp. and 1

Acinetobacter sp). A novel indigenous Pseudomonas aeruginosa strain (MTCC 4996)


23 isolated by Kotresha and Vidyasagar (2008) from a pulp industrial effluent-

contaminated site was capable of degrading phenol upto a concentration of 1,300 mg

L–1 within 156 hr. Paraskevi and Euripides (2005) reported a Pseudomonas sp. strain

(phDv1) isolated from a sample of petroleum-contaminated soil in Denmark capable

of growing on phenol up to concentrations of 1,200 mg L–1. Suhaila et a., (2012) had

isolated Rhodococcus sp.UKMP-5M from a petroleum contaminated soil. Karigar et

al., (2006) studied the ability of Arthrobacter citreus, isolated from a hydrocarbon

contaminated site, to consume phenol as the sole carbon source. The phenol

degradation studies in their work showed that complete degradation of the compound

occurred within 24 hr. Jiang et al., (2007) isolated a strain of Alcaligenes faecalis

from activated sludge collected from a municipal gasworks.

Shourian et al., (2009) reported that many pollution problems resulting from

releasing aromatic chemicals occur in rivers, lakes, groundwater, and process

effluents from the industry. Accordingly, environmental bacterial strains isolated from

contaminated sites are expected to play a vital role in the bioremediation of

contaminated areas. In their study, Pseudomonas sp. SA01 was isolated from

pharmaceutical wastewaters. The bacterial strain was examined for phenol

biodegradation and was suggested as an excellent and unique candidate for rapid and

efficient phenol removal, particularly for the shorter lag time at high phenol

concentrations (up to 1000mg/l) compared with those occurring in other

Pseudomonas species. Other researchers have utilized microbial biomasses for phenol

degradation which are isolated from phenol-contaminated soil (Liu et al., 2009),

wastewater plants (Khleifat, 2006; Kilic, 2009), effluents of coke processing units,

and municipal gas works (Jiang et al., 2007).


24

The microbiological analysis of oil refinery effluents indicated that the

effluents are a good habitat for bacteria and fungi. The isolation of various bacterial

genera from the effluents that are able to utilize phenol as carbon and energy sources

showed that they are potential phenol degraders present in such specific habitats

(Kopytko and Jacome, 2008). Studies have shown that lower doses of phenol are

more readily utilized than higher doses. These corroborate with the high growth

responses at low doses (0.5 and 1.0mM respectively) of phenol. Stimulation of

dehydrogenase activity at low dose of phenol was reported for Bacillus and

Pseudomonas species isolated from petroleum refinery wastewater (Nweke and

Okpokwasili, 2010a).

A number of aerobic phenol degrading bacteria have been described

previously; however, little information on bacteria with a high phenol tolerance with

high metabolizing activity is available (El-Sayed et al., 2003). Despite the significant

amount of information gathered during the years, the problem is still topical and

significant, which stimulates researchers for new developments in the biodegradation

of aromatic compounds and the characterization of new and more effective microbial

species. A lot of examples exist of the considerable interest in this area. Bacterial

strains have been isolated from phenol-containing industrial wastewater and identified

as Pseudomonas cepacia and Bacillus brevis, which show extremely high

effectiveness of phenol degradation. Adapted cultures from these strains are capable

of degrading 2.5 and 1.75g/L phenol for 144 h, respectively. The study demonstrates

that these microorganisms are able to degrade phenol even in the presence of highly

toxic compounds, such as thiocyanides, sulfides, and cyanides, which makes them


25 suitable for the technological treatment of wastewaters having similar content

(Arutchelvan et al., 2005).

A large number of phenol degrading bacteria have been isolated and

characterized at the physiological and genetic levels (El-Sayed et al., 2003; Koutny et

al., 2003; Shen et al., 2004; Arutchelvan et al., 2005; Polymenakou and Stephanou,

2005; Wu et al., 2005; Wang et al.,2007). A total of 39 phenol- and p-cresol-

degraders isolated from the river water continuously polluted with phenolic

compounds of oil shale leachate were studied. Species identification by BIOLOG GN

analysis revealed 21 strains of Pseudomonas fluorescens (Heinaru et al., 2000).

Wael et al., (2003) isolated new phenol degrading bacteria with high tolerance

and high biodegradation activity. The isolates were Burkholderia capacia PW3 and

Pseudomonas aeroginosa AT2. Both the isolates could grow aerobically on phenol as

sole carbon source and tolerated up to 3000ppm of phenol. The metabolic pathway for

phenol biodegradation in both the strains was assigned to the meta-cleavage activity

of catechol 2,3-dioxygenase.

Arutchelvan et al., (2006) have isolated a novel strain of Bacillus brevis from

the phenol–formaldehyde resin manufacturing industrial wastewater and utilized for

biological degradation of phenol. The authors optimized various environmental

conditions for phenol biodegradation in a batch reactor with pure culture of B. brevis

and the maximum biodegradation of phenol was at pH 8.0, 5% (v/v) of inoculum size

and without any co-substrate.


26

Wenjing et al., (2008) isolated 46 phenol/benzoate degrading and iron

reducing bacteria from long term irrigated tropical paddy soils by enrichment

procedures. Pure cultures and some prepared mixed cultures were examined for ferric

oxide reduction and phenol/benzoate degradation. All the isolates were iron reducers,

but only 56.5% could couple iron reduction to phenol and/or benzoate degradation, as

evidenced by depletion of phenol and benzoate after one week incubation. Analysis of

degradative capability using Biolog MT plates revealed that most of them could

degrade other aromatic compounds such as ferulic acid, vanillic acid, and

hydroxybenzoate. Mixed cultures and soil samples displayed greater capacity for

aromatic degradation and iron reduction than pure bacterial isolates. Bacteria capable

of coupling these reactions may be major contributors to the microbial cycling of

large molecule carbon substrates and are thus promising tools for bioremediation and

elimination of organic pollutants in Fe mediated anaerobic simulators.

Kilic et al., (2007; 2009) reported the isolation and characterization of the

bacterium Ochrobactrum sp. It was metabolized phenol through catechol, followed by

ortho- or meta- pathways (El-Sayed et al., 2003). There is no report investigating

phenol degradation capacity and the conditions affecting phenol degradation by

Ochrobactrum sp. Furthermore, some other phenolic compounds like 4-nitrocatechol

(Zhong et al., 2007), p-nitrophenol (Qiu et al., 2007), and 2,4,6- tribromophenol

(Yamada et al., 2008) were metabolized by Ochrobactrum species.

Jame et al., (2010) isolated four different Pseudomonas species viz FA, SA,

TK and KA. All these isolates could completely degrade phenol up to 600ppm. Isolate

Pseudomonas FA degraded 800ppm phenol completely in 72 h, but the isolates


27 Pseudomonas SA, TK and KA degraded only 39.33, 43.83 and 33.16% of 800ppm

phenol respectively in 96 h. From mixed cultures of Pseudomonas putida A (a) and

Pseudomonas sp. SA, it was found that P. putida A (a) degraded 600ppm phenol in 24

hour and Pseudomonas sp. SA degraded the same concentration in 72 h when they

were cultured individually. The mix culture of this two Pseudomonas sp. degraded the

same concentration in 20 h.

Hong-xia (2011) isolated 15 different bacterial strains from marine sources on

the beef extract peptone agar plates with 1500mgL-1 phenol. Among them, the strain

SM5 could tolerate 4500 mg/L phenol on solid beef extract peptone plates and its

phenol biodegradation rate was 96.4% in basal salt (BS) medium under the optimum

conditions when the concentration of phenol was 1000 mg/L. These conditions were;

initial pH 7.0, 37°C, 3 days, 20 ml medium/50 ml flask and inoculums biomass 12.5%

(v/v). Rate of phenol biodegradation of the strain was up to 92.0% under the optimum

conditions even when the phenol concentration was increased to 2500 mgL-1.

Staphylococcus aureus was isolated from effluent sample by enrichment of the

effluent. This isolated bacterium was tested for its potential of phenol remove from

effluent by added 1000ppm of phenol to Bushnell Haas (BH) medium as a sole source

of carbon and nitrogen. The result indicated that the strain has potential to remove

phenol up to 800ppm within 7 days (Naresh et al., 2012).

Nwanyanwu et al., (2012) studied about the fungal and bacterial population of

petroleum refinery effluent samples by microbial enumeration and determination of

growth responses of bacterial isolates in increasing doses (0 - 15mM) of phenol in

mineral salt-phenol agar. From the effluent samples Bacillus sp. RWW, Aeromonas


28 sp. RBD, Escherichia coli OPWW and Staphylococcus sp. DP were isolated. Growth

responses of the isolates on increasing doses of phenol in mineral salt-phenol agar

showed that Bacillus sp. RWW and Escherichia coli OPWW had highest growth on

0.5mM ( 47.06mgL-1) while Aeromonas sp. RBD and Staphylococcus sp. DP had

their highest growth on 1.0mM ( 94.11mg L-1). Bacillus sp. RWW and Escherichia

coli OPWW showed least growth on 15.0mM ( 1,412mg L-1) of phenol. Escherichia

coli OPWW exhibited highest growth in mineral salt broth containing 11.0mM of

phenol with OD540nm of 0.324 in 144 hr resulting in the fastest utilization of phenol

for growth. The highest specific growth rate of 0.013 h-1 at 11 mM ( 1,035mg L-1) of

phenol was obtained for Bacillus sp. RWW and Escherichia coli. Staphylococcus sp.

DP had the lowest specific growth rate of 0.011 h-1 at 11 mM of phenol. These

bacterial strains could be considered phenol-resistant and are potentially applicable in

the removal of phenolic compounds from contaminated environmental media.

Six phenol degrading bacteria designated as PND-1 to PND-6 were isolated

from natural soil by the direct spreading plate method for avoiding the biodiversity

alteration by enrichment method. On the basis of morphology, physiological and

biochemical characteristics and 16S rDNA sequence analysis, PND-1, PND-2 were

identified as Pseudomonas sp., PND-4 and PND-5 were from the genus

Acinetobacter, and strain PND-3 and PND-6 belonged to Comamonas sp. and

Cupriavidus sp., respectively. All these strains were able to utilize phenol as the sole

carbon source to support their growth. Five of them could tolerate the phenol

concentration up to 6mM or more, while strain PND-3 could tolerate only 1mM of

phenol. The sequences of the partial largest subunit of multicomponent phenol

hydroxylase (LmPH) gene were compared among these strains. It was found that the


29 physiological groupings in the phylogenetic tree formed by their 16S rDNA

sequences were correlated with that of based on their partial amino acid sequences of

LmPH, which indicated the potential application of LmPH as a molecular marker for

the phylogenetic analysis of phenol-degrading strains. Four of the six strains degraded

phenol through catechol ortho fission pathway, whereas strain PND-3 and PND-6

harbored the both ortho and meta fission pathways simultaneously (Dong et al.,

2008).

Eight morphologically different bacterial strains capable of biodegrading

phenol were isolated from activated sludge of a pharmaceutical industry using

enrichment culture technique. Two of the bacterial strains capable of utilizing phenol

as a sole source of carbon were isolated from the wastewater of a pharmaceutical

industry. On the basis of morphological and biochemical characteristics these strains

were identified as Pseudomonas aeruginosa and Pseudomonas pseudomallei. Both of

these strains were very efficient for phenol degradation. P. pseudomallei degraded

phenol at a maximum concentration of 1500 mg L 1 within seven days with a specific

growth rate of 0.013 h 1 and phenol degradation rate of 13.85 mg L 1 h 1. Maximum

initial concentration of phenol utilized by P. aeruginosa was 2600 mg L 1 with 0.016

h 1 specific growth rate and 26.16 mg L 1 h 1 phenol degradation rate (Afzal et al.,

2007).

A bacterium able to degrade bisphenol A (BPA) as sole carbon and energy

source was isolated from a planted fixed bed reactor continuously running with BPA.

The Gram-negative aerobic bacterium was identified as Cupriavidus (formerly

Wautersia, Ralstonia) basilensis JF1 by using physiological characterization,


30 BIOLOG and 16S rRNA-analysis. Furthermore, C. basilensis JF1 is able to utilise

phenol, 4-isopropylphenol, ethanol, methanol, acetone and several other aromatic and

aliphatic compounds as sole carbon sources. By using phenol as biostimulant either in

shake cultures or in continuously running sand columns, the BPA-degradation rate of

the bacterium could be significantly enhanced (Fischer et al., 2010).

A sulfate- reducing bacterium isolated from swine manure used phenol as its

sole source of carbon and energy. Sulfate was used as the electron acceptor. The

major end product of phenol metabolism was acetic acid. For every mole of phenol

degraded, almost two moles of acetic acid were produced. Acetic acid was not

degraded further to CO2, indicating that this sulfate-reducing bacterium (SRB) is an

incomplete oxidizer unable to carry out the terminal oxidation of organic compounds.

The SRB isolate also used P- chlorophenol as the sole source of carbon and energy.

However; it did not use the chlorophenolic compounds containing two or more

chlorine atoms, dichlorophenol and pentachlorophenol (Boopathy, 1995).

2.7. Mode of biodegradation of phenol

Degradation of phenol using microorganisms has been extensively studied and

their mechanism is classified based on their mode of respiration into aerobic and

anerobic biodegradation. The two types are adapted from Basha et al., (2010) and

discussed in the following sessions.

2.7.1 Aerobic biodegradation of phenol

Aerobic biodegradation has been studied extensively. In the first step of the

aerobic pathway (Figure 2) for the biodegradation of phenol, molecular oxygen is


31 used by the enzyme phenol hydroxylase to add a second hydroxyl group in ortho-

position to the one already present in which reaction requires a reduced pyridine

nucleotide (NADH2). The resulting catechol (1, 2-dihydroxybenzene) molecule can

then be degraded via two alternative pathways depending on the responsible

microorganism. In the ortho or -ketoadipate pathway, the aromatic ring is cleaved

between the catechol hydroxyls by a catechol 1, 2-dioxygenase (intradiol fission)

(Harwood and Parales, 1996; Stanier and Ornston, 1973). Preliminary evidence for

the production of -ketoadipate during the degradation of phenol by strain 'Vibrio

01'was first presented by Evans and Kilby (Evans, 1947; Kilby, 1948). The resulting

cis, cis muconate is further metabolized via -ketoadipate to Krebs cycle

intermediates. In the meta-pathway, ring fission occurs adjacent to the two hydroxyl

groups of catechol (extradiol fission). The enzyme catechol 2, 3-dioxygenase

transforms catechol to 2-hydroxymuconic semialdehyde. This compound is

metabolized further to intermediates of the Krebs cycle. The organisms which utilize

phenol by aerobic pathway are Acientobacter calcoceticus, Pseudomonas species and

Candida tropicalis and most of the eukaryotes typically employ ortho pathway. The

aerobic genus Pseudomonas species have been subject to various studies and its

versatility to utilize a wide spread of aromatic substrates makes it an attractive

organism for use in waste water treatment applications.

The aerobic and anaerobic degradation of phenol has been studied extensively

using various microorganisms (Ruiz-Ordaz et al., 2001; Mendonça et al., 2004; Yan

et al., 2005). Phenol may be converted by bacteria under aerobic conditions to carbon

dioxide (Aquino et al., 1988) and under anaerobic conditions to carbon dioxide

(Tschech and Fuchs, 1987) or methane (Fedorak et al., 1986). Under aerobic


32 condition, oxygen is used as electron acceptor for transfer of electrons between the

electron-donor and electron-acceptor. For these process substrates is essential for

creating and maintaining biomass. In the phenol biodegradation process, the primary

substrate (phenol) must be made available in order to have biomass active. According

to Rittmann and Saez (1993) once active biomass is present, any biotransformation

reaction can occur, provided the microorganisms possess enzymes for catalyzing the

reaction. These enzymes that are involved in the aerobic metabolism of aromatic

compounds usually define the range of substrates that can be transformed by certain

metabolic pathway (Pieper and Reineke, 2000).

First step of aerobic phenol metabolism is catechol production by a NADPH-

dependant flavoprotein phenol hydroxylase (EC 1.14.13.7) (Enroth et al., 1998). The

second step is catalyzed by catechol 1,2-dioxygenase (EC 1.13.11.1; ortho fission) or

catechol 2,3-dioxygenase ( EC 1.13.11.2; meta fission). After several subsequent

steps, the products are incorporated into the tricarboxylic acid cycle (TCA) or Krebs

cycle (Shingler, 1996). It has been established that the aerobic degradation of phenolic

compounds is metabolized by different strains through either the ortho-or the meta-

cleavage pathway (Bayly and Barbour, 1984; Ahamad and Kunhi, 1996; Shingler,

1996).


33

PHENOL

Hydroxylase

META PATHWAY CATECHOL ORTHO PATHWAY

Catechnol 2, 3 - dioxygenase Catechnol 1, 2 - dioxygenase

2- HYDROXYMUCONIC SEMIALDEHYDE CIS, CIS MUCONATE

2-HMSA dehydrogenase Lactonizing enzyme 4 – OXALOCROTONIC ACID MUCONOLACTONE

Decarboxylase, CO2 I Isomerase

2 –OXYOPENT- 4- ENOATE 3-OXOADIPATE-ENOL-LACTONE

2-keto-4-pentenoate hydratase 3-Oxoadipate enol - lactonase

4- HYDROXYL -2- OXOVALERATE 3-OXOADEPATE

Aldolase Transferase

ACETALDEHYDE + PYRUVATE 3-OXOADIPYL COA

Thiolase Acyltransferase

SUCCINYL CO-A ACETYL CO-A

Figure 2 Flow chart of aerobic degradation pathway for phenol (Basha et al., 2010)


34 2.7.2 Anaerobic biodegradation of phenol

Phenol can also be degraded in the absence of oxygen and it is less advanced

than the aerobic process. It is based on the analogy with the anaerobic benzoate

pathway proposed for Paracoccus denitrificans in 1970 (Williams and Evans, 1975).

In this pathway phenol is carboxylated in the para position to 4 hydroxybenzoate

which is the first step in the anaerobic pathway. Here the enzyme involved is the 4-

hydroxy benezoate carboxylase. The anaerobic degradation of several other aromatic

compounds has been shown to include a carboxylation reaction. Carboxylation of the

aromatic ring in para position to the hydroxy group of o-cresol resulting in 3-methyl

4-hydroxybenzoate has been reported for a denitrifying Paracoccus like organisms, as

well as methogenic consortium was later shown to travel a varity of phenolic

compounds including o-cresol, catechol and ortho halogenated phenols via para

carboxylation followed by dehydroxylation. The organisms capable of degrading

phenol under anaerobic conditions were Thauera aromatica and Desulphobacterium

phenolicum.

2.8 Key Enzymes in the biodegradation of phenol and its derivatives

The ability of microorganisms to transform xenobiotics into compounds that

can enter the normal cycle of matter is due to specific microbial enzymes. Thus, the

investigation of enzyme reactions including degradation and detoxification of phenol

pollutants is the focus of attention for many specialists. The metabolism of aromatic

compounds and its regulation is extensively studied in prokaryotes. Phenol is

widespread, microorganisms capable of utilizing this compound as a carbon and

energy source can be found in many different habitats. There are both aerobic and

anaerobic microorganisms that are able to complete the phenol degradation process

(Agarry et al., 2008).


35

PHENOL

Decarboxylase

4-HYDROXYBENZOATE

p-hydroxy benzoate 3- monooxygenase

PROTOCATECHUATE

Protocatechuate 3, 4 dioxygenase

ß – CARBOXYMUCONATE

Cycloisomerase

– CARBOXYMUCONATE

Decarboxylase

3-OXOADEPATE ENOL-LACTONE

Enol - lactonase

3-OXOADIPATE

Transferase

3-OXOADIPYL CO A Thiolase Acyltransferase

Succinyl Co-A Acetyl co-A

Figure 3 Flow chart of anaerobic degradation pathway for phenol (Basha et al., 2010).


36

The phenol degradation under aerobic condition in eukaryotes such as yeasts

and filamentous fungi is catalyzed mainly by two key enzymes namely phenol

hydroxylase (PH) and catechol 1,2-dioxygenase (C1,2D) via the ortho ring cleavage

pathway of catechol to cis,cis-muconic acid (ccMA). The end products of phenol

degradation in eukaryotic cells were succinic acid and acetyl CoA. On the other hand,

phenol degradation in prokaryotes such as Pseudomonas sp. is catalyzed by phenol

hydroxylase (PH) and catechol 2,3-dioxygenase (C2,3D) of catechol to

2-hydroxymuconic semialdehyde (2-HMSA) via the meta ring cleavage pathway. The

activity of these enzymes could be affected by temperature, pH and concentration of

initial phenol concentration. The end products of phenol degradation are non-toxic

intermediate compounds that enter into the Tricarboxylic acid cycle (TCA) or Krebs

cycle through ortho- or meta-pathways of degradation (Schie and Young, 2000).

There are reports on many microorganisms capable of degrading phenol

through the action of verity of enzymes. Enzymes involves in phenol degradation are

located in the cytoplasm. Table 4 shows the enzymes involves in the phenolic

compound degradation. There are four key enzymes in phenol degradation: phenol

hydroxylase (EC 1.14.13.7), catechol 1,2-dioxygenase (EC 1.13.11.1), cis,cis-

muconate lactonizing enzyme (EC 5.5.1.1), and 3-oxoadipate enol-lactone hydrolase

(EC 3.1.1.24) (Neujahr and Kjellen, 1978).

2.8.1 Phenol hydroxylase

Phenol hydroxylase catalyzes the attachment of a hydroxyl group at the ortho-

position of the aromatic ring, thus hydroxylating phenol to catechol. This reaction is

realized by an enzyme characterized as an NADP-dependent flavin monooxygenase


37 and is the first step in the degradation of aromatic compounds in microorganisms. The

phenol hydroxylase isolated and described in yeast was characterized as a mixed

function oxydase containing FAD and strictly dependent on cofactor NADPH (Basha

et al., 2010).

Table 4. Enzymes involved in the phenolic compounds biodegradation

Type of Phenol Enzyme Reference

Phenol Phenol hydroxylase Gurujeyalakshmi and Oriel (1988)

Phenol Polyphenol Oxidase Burton et al. (1993); Cano et al., (1997); Schneider et al.,

(1999)

Monophenol Polyphenol Oxidase Edwards et al., (1999)

Chlorogenic acid (Natural phenol) Polyphenol Oxidase Leonardes et al., (2005)

Phenol Catechol 2,3 dioxygenase Ali et al., (1998)

Phenol Catechol 1,2 dioxygenase An et al., (2001)

Phenol Laccase Bollag et al., (1998)

Phenol Peroxidase Ghioureliotis and Ncell, (1999)

Bisphenol Peroxidase Sakurar et al., (2001)

Phenol Horse radish peroxidase Wu et al., (1998)

Phenol Tyrosinase Xiangchun et al.,

(2003), Siegbahn, (2003)

Phenyl phosphate Phenyl phosphate carboxylase

Lack and Fuchs, (1992)


38

An interesting fact is that phenol hydroxylase could also hydroxylate catechol,

and the product of the reaction was pyrogallol. When phenol was the only substrate

for the enzyme in the reaction mixture, the formation of pyrogallol could only be

observed at very high substrate concentrations (Basha et al., 2010). This could

probably be explained by the inhibition of phenol hydroxylase in the case of substrate

excess (phenol) and the change in enzyme specificity in such conditions (Krastanov et

al., 2013).

There are many reports on phenol hydroxylase involved in the biodegradation

of phenol (Leonard and Lindey, 1999). Phenol hydroxylase from Trichosporon

cutaneum showed different activity toward the so-called “substituted phenols”, which

was not observed for the enzyme isolated from Pseudomonas picketti PK01. The

enzyme characteristic for P. picketti PK01 was induced by only two phenolic

substrates (phenol and 3-methylphenol), while the enzyme from T. cutaneum was

induced by all three isomers of methylphenol and the three fluorophenol isomers. All

these specific differences could be explained by the different linking positions of the

substrates, as well as by the different structure of the enzymes isolated from different

microorganisms. The detailed study on the protein structure of the enzyme established

that there was homology in the section at the N-end of phenol hydroxylase in yeasts

and p-hydroxybenzoate hydroxylase in Pseudomonas sp. (Nurk et al., 1991).

In genus Pseudomonas strains, the structural gene for phenol hydroxylase was

plasmid determined, and the gene was sequenced and cloned (Nurk et al., 1991). It

was found that this gene had 46% homology with 2,4-dichlorophenol hydroxylase

from A. eutrophus (Perkins et al., 1990). The cloning, sequencing, and expression of


39 the T. cutaneum phenol hydroxylase gene in Escherichia coli was first reported by

Kalin et al., (1992). Scientists also observed some differences during the investigation

of the enzyme structure of phenol hydroxylase isolated from different yeast species.

For example, the partially purified enzyme from C. tropicalis ATCC 46491 showed

higher substrate specificity than the enzyme isolated from a T. cutaneum strain.

2.8.2 Catechol dioxygenase

The second enzyme from the ortho- mechanisms of the 3-oxoadipate pathway

for phenol degradation is catechol 1,2-dioxygenase (EC 1.13.11.1.). In the meta-

mechanisms, the catechol 2,3-dioxygenase enzyme (EC 1.13.1.2.) hydrolyzes the

bond at the meta- position in the aromatic ring. The product of this reaction is

2-hydroxymuconic semiadlehyde, which is later broken down to acetaldehyde and

pyruvate (Krastanov et al., 2013). This enzyme is often used for evaluation of the

potential of different microbial associations with defined or undefined content for

aerobic degradation of aromatic compounds (Grekova-Vasileva and Topalova, 2008).

The first intermediate product of phenol degradation is catechol. The

dioxygenase enzyme that catalyzes the aromatic ring cleavage of catechol and its

derivatives realizes the critical step in the aerobic degradation of aromatic compounds

in microorganisms. Two classes of such enzymes are identified on the basis of

aromatic ring cleavage mechanisms: intradiol-dioxygenases and

extradioldioxygenases (Krastanov et al., 2013). Contemporary genomic, structural,

spectroscopic, and kinetic studies broaden the knowledge of the distribution,

evolution, and action mechanisms of these enzymes. Extradiol dioxygenases are


40 generally believed to have more activity than intradiol dioxygenases (Vaillancourt et

al., 2006).

Initially, the enzyme was isolated and purified from bacteria of the

Pseudomonas genus. It was established that the enzyme dioxygenase incorporated

molecular oxygen directly in the aromatic ring of catechol, which resulted in the

formation of cis,cis-muconic acid. The enzyme catechol 1,2-dioxygenase described in

Pseudomonas is highly dependent on ferro- and ferri-ions and has high substrate

specificity (Nakai et al., 1990).

Recently, a new catechol 1, 2-dioxygenase was isolated form a Pseudomonas

aeruginosa TKU002 strain capable of assimilating benzoic acid as a single carbon

source. The enzyme has unique characteristics, such as very low molecular mass

(22 kD), highest activity against pyrogallol, high medium acidity for enzyme

production, etc., which distinguishes it from other microbial catechol dioxygenases

(Wang et al., 2006). One of the best characterized eukaryotic catechol 1,

2-dioxygenases was isolated from the phenol-assimilating C. albicans TL3 strain. An

ortho-mechanism for phenol degradation was determined through the application of

enzyme, chromatographic, and mass spectrophotometric analysis. The strain was also

capable of degrading formaldehyde, which is one of the major pollutants in

wastewaters from phenolic product manufacture (Tsai and Li, 2006).

During the investigation of a gene-coding catechol 2,3-dioxyenase in the

aniline degrading bacterium Acinetobacter sp. YAA, it was found to exhibit different

activity. This enzyme has a tetramer structure of identical subunits with molecular

mass of 35 kD (Takeo et al., 2007). In bacteria of which meta- cleavage of phenolic


41 compounds is characteristic, the meta-pathway coding genes are often large plasmids

like TOL or NAH plasmids (van der Meer et al., 1992).Unlike them, the genes

responsible for the ortho-cleavage pathway are generally situated on the chromosome

(Wagner et al., 1999).

2.8.3 Cis, cis-muconate cyclase

The third enzyme from the ortho-mechanism of the 3-oxoadipate pathway for

phenol degradation is cis, cis - muconate lactonizing enzyme (EC 5.5.1.1.). It can also

be found under the name cis, cis - muconate cycloisomerase and cis,cis-muconate

lactonase. It catalyzes the transformation of cis,cis-muconate into muconolactone.

One of the most thorough investigations on the characteristics of this enzyme, which

can be found in different variations in various microorganisms, was dedicated to the

evolution of enzyme activity in the so-called “superfamily” of enolases. The authors

divided this group of enzymes into two families of cis,cis-muconate lactonizing

enzymes: syn- and anti-, according to their different stereochemical substrate

preferences. Representative of these groups are the enzymes from Pseudomonas

fluorescens (syn), and Mycobacterium smegmatis (anti) (Sakai et al., 2009).

2.9 Factors influencing and affecting biodegradation of phenol by bacteria

Biodegradation is a multifaceted process in which many biotic and abiotic

factors are involved. There are many factors that can control degradation ability or

metabolism of microorganisms by either preventing or stimulating growth of the

organisms; hence it is very important to identify those factors to obtain maximum

biodegradation of phenol. These factors may include temperature, pH, oxygen content


42 and availability (aeration and agitation), bioavailability (availability of the

contaminants to microbes), substrate concentration, the presence of other nutrients,

and physical properties of contaminants. Each of these factors should be optimized for

the selected organism to achieve the maximum degradation of the organic compound

of choice (Agarry et al., 2008; Nair et al., 2008 and Trigo et al., 2009).

2.9.1 Effect of pH on phenol degradation by bacteria

The internal environment of all living cell is believed to be approximately

neutral (Basha et al., 2010). Hence, majority of organisms could not survive at a pH

range below 4.0 or above 9.0 (Kim and Armstrong, 1981). At a low (4.0) or high (9.0)

pH values, acids or bases can penetrate into cells more easily as they exist in

undissociated form under these conditions and electrostatic force cannot prevent them

from entering cells (Robertson and Alexander, 1992; Annadurai et al., 1999).

Many authors had reported that biodegradation occurs near neutral pH. For

instance, the optimum pH for phenol degradation is 7.0 for Pseudomonas putida

NICM 2174 (Annadurai et al., 2000). Also, the ability of bacterium isolated by Awan

et al., (2013) to degrade phenol at different pH was also observed and maximum

degradation was recorded at pH 7. At high or low pH values acid or base could burst

during into cells further simply, because they have affinity to survive in undissociated

structure underneath these circumstances and electrostatic force cannot shun them

from incoming cells (Alaxander and Robertson, 1992).

The optimum pH for phenol degradation is about 7.0 for majority of bacteria.

Naresh et al., (2012) suggested that at a pH of 7 was optimum for maximum phenol


43 removal by Staphylococcus aureus. Kotresha and Vidyasagar (2008) had observed

that the biodegradation of phenol using Pseudomonas aeruginosa strain (MTCC

4996) was optimum at pH 7. Arutchelvan et al., (2006) have reported maximum

phenol degradation using Bacillus brevis at pH 8.0. Kumar et al., (2005) also reported

the maximum degradation of 1000ppm phenol in 162 hours using P. putida MTCC

1194 under the optimum process variables of pH 7. Similarly, Bandyopadhyay et al.,

(2001) reported 1000ppm of phenol degradation using P. putida MTCC1194 under

optimum pH of 7. Suhaila et al., (2012) had found that the maximum phenol

degradation by Rhodococcus sp. was at a pH of 7.5. Table 3 lists the optimum pH for

biodegradation of phenol by bacteria that has been isolated by various researchers.

Based on the previous literature reported, the importance of pH on

biodegradation of phenol mediated by bacteria is well understood and hence in this

study, the effect of pH has been tested.

2.9.2 Effect of temperature on phenol degradation by bacteria

The cell growth is also significantly affected beyond the optimum temperature

because the variations in temperature affect the viability of the cells and are lethal for

them. At low temperatures, the fluidity of the membrane decreases sufficiently which

prevent the functioning of the transport systems, so the substrates cannot enter into

cell rapidly to support even low rate of growth. Increase in temperature affects

proteins by causing thermal denaturation, which is usually irreversible. Thus,

temperature is another important factor that determines the rate of phenol degradation.

Additionally, temperature might play an equivalent or larger role than nutrient

availability in the degradation of organic pollutants (Margesin and Schinner 1997).


44

Awan et al., (2013) had reported that 35oC was found as the optimum

temperature for the degradation of phenol. Naresh et al., (2012) suggested that at a

temperature of 37ºC was optimum for maximum phenol removal by Staphylococcus

aureus. Hank et al., (2010) had reported that the best temperature for degradation of

phenol by Pseudomonas aeruginosa (ATTC 27853) at a concentration of 100mgL-1

was found to be 30°C. Similarly Agarry et al., (2008) had reported that the optimum

temperature for the degradation of phenol by Pseudomonas aeruginosa was found to

be 30.1°C.Whereas Kotresha and Vidyasagar (2008) had observed that the

biodegradation of phenol using Pseudomonas aeruginosa strain (MTCC 4996) was

optimum at 37 ºC.

Annadurai et al., (1999) described that when the temperature increased to

beyond 30°C or 34°C, no phenol degradation was observed due to cell decay, which

is a temperature-dependent parameter. Pakula et al., (1999) had reported that phenol

biodegradation was significantly inhibited by strains isolated from a petroleum-

refining wastewater purification plant at 30 0C. At 30°C Pseudomonas SA01 had

significant degradation potential for the rapid utilization of phenol (Shourian et al.,

2009). Suhaila et al., (2012) had found that the phenol degradation by Rhodococcus

sp. was highest at a temperature of 30°C.

A study performed by Prieto et al., (2002) reported that the highest

degradation of phenol by Rhodococcus erythropolis was obtained at 300C. Kumar et

al., (2005) also reported the maximum degradation of 1000ppm phenol in 162 hr

using P. putida MTCC 1194 under the optimum temperature at 30°C. Similarly,

Bandyopadhyay et al., (2001) reported 1000ppm of phenol degradation using P.


45 putida MTCC1194 under optimum temperature of 30°C. The optimal conditions for

phenol degradation by Micrococcus sp. and Alcaligenes faecalis JH 1013 were

reported at temperature 32°C (Zeng et al., 2010). Bajaj et al., (2008) reported

maximum phenol degradation at 25°C by mixed bacterial consortium.

Table 5 lists the optimum temperature for biodegradation of phenol by

bacteria that has been isolated by various researchers.

2.9.3 Effect of agitation speed on phenol degradation by bacteria

Agitation is yet another parameter that determines the rate of biodegradation

of phenol. Insufficient agitation may lead to limitations in the transfer operations and

the appearance of regions of insufficient nutrient content or inadequate temperature or

pH (Gonzalez et al., 2003).Therefore, an intense agitation must be provided, but too

high agitation rates should be avoided to prevent attrition and metabolic stress in the

bacterial population (Toma et al., 1991; Enfors et al., 2001; Gonzalez et al., 2003).

Thus, the increase in biodegradation rate may be due to adequate high mass transfer

thus allowing more oxygen to be dissolved and made available for the metabolism of

the organism. While, the decrease may be due to higher shear stress effect thus

leading to cell loss or lower biomass concentration (Hoq et al., 1995). The speed of

120 rpm was found to be the optimum for the degradation of phenol (Awan et al.,

2013).


46

Table 5. Optimum pH and temperature for biodegradation of phenol by bacteria

Phenol degrading bacteria

Optimum pH

Optimum temperature References

P. putida DSM 548 6.8 26 ± 0.5 Monteiro et al., (2000)

P. putida ATCC 17484 6.6 30 Gonzalez et al., (2001)

Cupriavidus metallidurans 6.6 23.5 ± 1.4 Stehlickova et al., (2009)

Halomonas campisalis 8–11 30 Alva and Peyton (2003)

P. putida MTCC 1194 7.1 29.9 ± 0.3 Kumar et al., (2005)

P. putida CCRC 14365 7.0 30 Tsai and Juang (2006)

Alcaligenes faecalis 7.2 30 Jiang et al., (2007)

Actinobacillus species 7.0 37 Khleifat and Khaled, (2007)

Pseudomonas sp. SA01 6.5 30 Shourian et al., (2009)

Ochrobactrum sp. 8.0 30 Kilic (2009)

Ewingella americana 7.5 37 Khleifat (2006)

P.putida 7.0 34 Ravikumar et al., (2011)

High-efficiency bacterial strain SM5

7.0 37 Hong-xia (2011)

Pseudomonas aeruginosa MTCC 1034, Pseudomonas fluorescens MTCC 2421 and Bacillus cereus ATCC 9634.

7.0 30 Bhattacharya et al., (2012)

P. putida 7.0 30 Ullhyan and Ghosh (2012)


47

Agitation speed of 301 rpm was found to be the optimum for the degradation

of phenol by Pseudomonas aeruginosa (Agarry et al., 2008). But, a speed of 100–125

rpm was found to be the optimum for the degradation of phenol by Pseudomonas

aeruginosa (MTCC 4996) (Kotresha and Vidyasagar 2008). Khleifat (2006) had

reported that phenol degradation by Ewingella americana was effective at a rate of

200 rpm.

2.9.4 Effect of carbon sources on phenol degradation by bacteria

Phenol acts as a carbon source for most of the phenol degrading bacteria and

hence consumption of phenol by the bacteria positively influences its growth. In the

study by Rozich and Colvin (1985) it was found that the presence of glucose

attenuated the rate of phenol removal by phenol consuming cells. This shows that

supplementation of an additional carbon source interrupts the consumption of phenol

as the bacteria tend to preferentially select the carbon source for its growth. This may

be due to catabolic repression by glucose as reported by Papanastasious (1982), i.e.

the presence of glucose could inhibit utilization of the target substrate.

Naresh et al., (2012) had reported that the best phenol removal by

Staphylococcus aureus was observed when 0.5% of glucose was added. However, the

authors also concluded that there was no increase in rate of removal of phenol when

lactose and sucrose were added as compared to glucose. Kotresha and Vidyasagar

(2008) had reported that 0.25 g L–1 glucose was found to be the optimum

concentration for phenol degradation and higher concentration inhibited the phenol

degradation. The growth of Ralstonia eutropha, in which the fructose-grown cells in


48 the presence of phenol minimized the respiration rate, compared with that of only

phenol-grown cells (Leonard et al., 1999b).

Hence, knowing the importance of carbon source on phenol degradation, three

carbon sources, i.e, glucose, sucrose and starch were selected in this study and their

concentration was optimized for an effective biodegradation of phenol.

2.9.5 Effect of nitrogen sources on phenol degradation by bacteria

Naresh et al., (2012) had reported that Staphylococcus aureus has a potential

to remove maximum phenol when 0.2% of urea and ammonium chloride were used as

a nitrogen source. Kotresha and Vidyasagar (2008) had reported that peptone at low

concentrations influences the rate of phenol degradation; however, above 1.0 g L–1

peptone was inhibitory. It was also noted that the presence of yeast extract enhanced

the affinity of Pseudomonas putida for phenol (Armenante et al., 1995). Suhaila et al.,

(2012) had tested a wide range of nitrogen source such as (NH4)2SO4, phenylalanine,

glycine, ammonium chloride, histidine, alanine, leucine, sodium nitrate, proline and

cystein on the growth of Rhodococcus sp. and phenol degradation and found that

(NH4)2SO4 resulted in highest phenol degradation. Khleifat (2006) had reported that

yeast extract, casein and glutamine caused a repression in phenol degradation by 3.3,

1.6 and 0.06 fold, respectively.

Thus, in this study an organic and inorganic nitrogen sources was selected and

their concentration were optimized to enhance the biodegradation of phenol.


49 2.9.6 Effect of trace elements on phenol degradation by bacteria

Forty percent of hazardous wastes on the environmental are co-contaminated

with organic and heavy metals pollutants that pose health hazards to human and

wildlife. Common organic pollutants at these sites include phenol, polycyclic

aromatic hydrocarbons (PAHs), chlorinated solvents, cyanide, herbicide and

pesticides, while common heavy metal contaminants include arsenic, cadmium,

chromium, copper, lead, selenium, mercury, nickel, and zinc (Norena- Barrosa et al.,

2004). Isolation of bacterial strains from the co-contaminated sites are able to

degrade more than one organic pollutants and becoming increasingly important for

decontaminating polluted soil, sledges, and ground water (Jain and Sayler, 1987;

Chen et al., 2005). The use of these microorganisms may face various problems,

including poor survival, substrate accessibility or the presence of inhibitory

compounds (Lin et al., 2006).

Heavy metals are known to be powerful inhibitors of biodegradation activities

thus their presence may impair the biodegradation of aromatic compounds in polluted

sites (Said and Lewis, 1991; Roane et al., 2001; Amor et al., 2001; Lin et al., 2006;

Silva et al., 2007). Al-Saleh and Obuekwe (2005) reported that the simultaneous

contamination by heavy metals and organic compounds may also occur at industrial

areas. So, there is an increasing interest in bacterial strains that are able to degrade

aromatic compounds and tolerant to toxic metals (Wasi et al., 2008). It has previously

been shown that strains of Alcaligenes eutrophus bearing plasmids of metal resistance

and plasmids of biodegradation of polychlorinated biphenyls and 2,4-

dichlorophenoxyacetic acid degrade these xenobiotics more effectively in the


50 presence of nickel or zinc as compared to sensitive strain (Springael et al., 1993;

Collard et al., 1994). However, there have been no data on the interaction of genetic

systems of aromatic compounds degradation, cyanide assimilation, and metal

resistance and their effect on physiology, biodegradation efficiency, and the activity

of the key enzyme in multifunctional strains (El-Deeb, 2009).

Hughes and Poole (1989) and Sterritt and Lester (1980) had found that the

addition of certain metal ions at low concentration enhances the degradation rate.

Also, in some cases the metal traces had inhibited the degradation of phenol. For

instance, Kuo and Sharak Genther (1996) found that phenol biodegradation was most

sensitive when Hg (II) was added at a concentration of 0.1 and 0.7 ppm. Kotresha and

Vidyasagar (2008) had reported that metals, such as Fe, Cu, Pb, Zn and Mn,

stimulated and enhanced the rate of phenol degradation. The degradation of phenol in

the presence of metals may be due to the fact that microbes display a large range of

tolerance and resistance to heavy metals (Trevors et al., 1985). Kuo and Sharak

Genther (1996) also reported that the presence of Cd (II), Cu (II) and Cr (VI) at 0.01

ppm increased degradation of phenol.

Copper (Cu) an essential micronutrient, but above the certain threshold level

they become toxic to human and microorganism (Aelion et al., 2009). Copper is an

essential mineral found in almost all living systems. In animals copper is necessary

for hemoglobin (Hb) production and required for the uptake and utilization of iron. It

is one constituent of the metalloenzyme superoxide dismutase, which eliminates

harmful oxygen radicals from the body. Copper is also a component of cytochrome

oxidases, enzymes that play many key roles in organisms, including oxidative


51 phosphorylation. In bacteria Cu, Zn superoxide dismutase are located in the periplasm

or anchored to outer membrane (Steinman, 1987; D’orazio et al., 2001; Battistoni et

al., 2000). Cu can be extremely toxic to microbes and thus necessitates homeostatic

mechanisms for cell survival (Munson et al.,2000; Rensing and Grass,2003). Copper

induces transcription of both copA and cueO, which encode for the copper

transporting ATPase and periplasmic multicopper oxidase, respectively, leading to the

detoxification of copper (Petersen and Moller, 2000; Stoyanov et al., 2001).

Iron (Fe) is an essential element for bacteria due to its participation in the

tricarboxylic acid cycle, electron transport, amino acid and pyrimidine biosynthesis,

DNA synthesis, and other critical functions (Earhart, 1996). Iron is a micronutrient for

in vitro cultures and the typical concentration needed for optimal growth of

Pseudomonas aeruginosa is 0.3 to 1.8 µM (Shuler and Kargi, 2002; Vasil and

Ochsner, 1999). The effect of iron limitation on bacterial growth has been

documented for Escherichia coli cultures (Hartmann and Braun, 1981). Two studies

have shown that production of the phytotoxins, syringomycin, and syringotoxin from

P. syringae responds in batch culture to iron supplementation (Gross, 1985; Morgan

and Chatterjee, 1988). Iron is known to alter the physiology of other pseudomonads in

both batch and chemostat cultures (Kim et al., 2005; Ongena et al., 2008). Although

iron is the fourth most abundant element in the earth’s crust, its availability is very

low due to its low solubility in aqueous solution ([Fe3+] at pH 7, 10-18 µM) (Vasil

and Ochsner, 1999).

The biochemical importance of Fe has been demonstrated by laboratory

experiments (Greenberd et al., 1992; Geider et al., 1993 and Sudan and Huntsman,


52 1997) and theoretical predictions (Raven, 1990). Fe is required for many proteins and

enzymes and the reactions associated with them. It is also essential for cytochromes,

ferredoxin and iron-sulphur proteins. Biochemical components of nitrogen

metabolism also directly require iron. Iron occurs in nitrate reductase (NR) (Cardenas

et al., 1974) and is required for the nitrite reductase (NiR) enzyme system (Zumfr,

1974). Fe starvation has been reported to decrease the level of both these enzymes

(Verstreate et al., 1980). Iron supported the uptake of nutrients of medium by

microorganism. Boyd et al., (1996; 98) reported that the Fe enrichment enhanced

nitrate uptake.

Nickel (Ni) is an essential element in the nutrition of plants and animals. It is

essential for the activity of four known enzymes (Ankel-Fuchs and Thaner, 1988).

Currently, researches have been conducted about the effect of nickel on removal of

organic pollutants by activated sludge (Li et al., 2011; Gikas, 2007; Bryers, 1984).

Usually shock doses of nickel were added in continuous or batch reactors with various

concentrations. For continuous reactors, the shock nickel concentrations were 0.5–30

mgL-1 (Gikas, 2008; Ong et al., 2004; McDermott et al., 1965; Yetis and Gokcay,

1989; Lombrana et al., 1993). For batch reactors, corresponding concentrations

ranged from 1 to 320 mgL-1 (Gikas, 2007; Mowat, 1976; McDermott et al., 1965).

Among those studies conducted in continuous reactors, only a few researchers

examined the effect of continuous dosing Ni(II) on organic pollutants removal

(McDermott et al., 1965; Ong et al., 2004), recovery ability after termination of

dosing nickel (Ong et al., 2004), or the adaptation to nickel after acclimatization

(Yetis and Gokcay, 1989). However, the tested nickel concentration was relatively

low, with less than 10mgL-1. Therefore, it is necessary to study the effect of nickel on


53 organic removal efficiency and recovery ability when dosed continuously with high

concentration over a long time (Li et al., 2011). A few reported the effect of

cadmium, zinc, copper, lead and cobalt on microbial community of biological

wastewater treatment systems (Kelly et al., 1999; Principi et al., 2006; Foucher et al.,

2003).

Selenium (Se) is widely recognized as an important nutritional element

microorganism growth. It is involved in the active centre of the glutathione

peroxidase enzyme where it acts as an antioxidant by reducing hydrogen peroxide

(Tappel, 1974). Se is available in two form such as inorganic form and organic form.

It has been demonstrated that organic Se has higher bioavailability and greater

accumulation in tissues than the inorganic form (Mahan and Parrett 1996; Taylor et

al., 2005). Printer and Provasoli (1968) first demonstrated the stimulatory effect of Se

on the growth of three axenic marine Chrysochromulina Spp. Doblin et al., (1999)

reported that the low (nM) levels of Se are limiting the growth and biomass

production of microalgae. Concentration of Se from10-7 to 10-9M stimulated

Gymnodinium catenatum growth and biomass.

Zinc (Zn) is an essential trace element for bacterial growth and enzyme

activities. However, a high concentration of Zn shows toxicity and inhibition to

microbial processes. Community diversity is severely reduced by high levels of Zn

and only a very limited number of resistant bacteria can survive (Goulder et al., 1980;

Kelly et al., 2003; Bong et al., 2010). It is involved in a wide variety of cellular

processes. It is required for maintaining the structural stability of macromolecules and

it serves as a cofactor for more than 300 enzymes (Vallee, 1986; McCall et al., 2000).


54 Zn also plays a prominent role in gene expression as a structural component in a large

number of Zn -dependent transcription factors. Zn also required for the normal

healthy growth and reproduction of crop plants -is required in relatively small

concentrations in plant tissues (5–100 mg/kg). Root cell membrane permeability is

increased under Zn deficiency, which might be related to the function of Zn in cell

membranes (Parker et al., 1992).

Hence, in the current study, the effect of trace elements such as Fe, Ni, Se, Cu

and Zn has been tested.

2.10 Immobilization techniques in phenol biodegradation

Many biodegradation studies have been focused on the use of the cell free

systems however such systems become impractical for the effluent treatment. Thus,

immobilization of bacterial biomass for the degradation of phenol can be employed

for effluent treatment as they can be used for longer periods that also protect the

bacteria from high phenol concentrations as well as enables ease of separation and

reutilization of the biomass (Naas et al., 2009).

Immobilization generates continuous economic operations, automation, high

investment/capacity ratio and recovery of product with greater purity (D’Souza,

1998). Inert polymers and inorganic materials are usually used as carrier matrices.

Apart from being affordable, an ideal matrix must encompass characteristics like

inertness, physical strength, stability, regenerability, ability to increase biocatalyst

specificity/activity and reduce product inhibition, nonspecific adsorption and


55 microbial contamination. Immobilized biocatalysts can either be enzymes or whole

cells (Kawaguti et al., 2006; Singh, 2009).

Several methods are used for enzyme/cells immobilization.

Adsorption/carrier-binding method uses water-insoluble carriers such as

polysaccharide derivatives, synthetic polymers and glass (Al-Adhami et al., 2002;

Rosa et al., 2002; Wu and Lia, 2008; Cordeiro et al., 2011). In cross-linking/covalent

method, bi/multifunctional reagents such as glutaraldehyde, bisdiazobenzidine and

hexamethylene diisocyanate are used (Lee et al., 2006; Singh, 2009). Polymers like

collagen, cellulose and k-carrageenan are employed by entrapment method, while the

membrane confinement method includes formulation of liposomes and microcapsules

(Wang and Hettwer, 1982; Mislovicova et al., 2004; Hilal et al., 2006; Tumturk et al.,

2007; Rochefort et al., 2008; Jegannathan et al., 2010; Chen et al., 2011a, b; Klein et

al., 2011).

An immobilized cell is one of the approaches for incorporating bacterial

biomass into an engineering process. The advantages of the process based on

immobilized biomass include enhancing microbial cell stability, allowing continuous

process operation and avoiding the biomass-liquid separation requirement. Varieties

of microorganisms have been immobilized by entrapment methods using matrix

systems like agar, sodium alginate, activated carbon. Thus, the potential of

immobilizing cells for industrial and biotreatment applications is of great value.

Physical entrapment of organisms inside a polymeric matrix is one of the most widely

used techniques for whole-cell immobilization (Klein and Schara, 1981).


56

Immobilization of bacterial biomass for the degradation of phenol is an

important and effective technique that is usually employed to serve several purposes,

including protection of the bacteria from high phenol concentrations as well as ease of

separation and reutilization of the biomass. It has been reported that the use of free

bacterial cells for wastewater treatment in activated sludge processes creates problems

such as solid waste disposal, while immobilized microorganisms are capable of

effective treatment with little sludge formation (Ying et al., 2007; Liu et al., 2009).

The biodegradation rate of phenol can be improved by immobilizing the cells

and entrapping them on solid support particles such as alginate, polyacrylamide,

chitosan (a natural nontoxic biopolymer), diatomaceous earth, activated carbon,

sintered glass, polyvinyl alcohol (PVA), and polymeric membrane to obtain the

maximum degradation capability (Mordocco et al., 1999; Chung et al., 2005; Liu et

al., 2009 and El-Naas et al., 2009).

Immobilization of bacterial cells enables to achieve faster degradation of

phenol compared to free cells. The immobilization method is not toxic to the cells and

is inert and practical (Aksu and Bulbul, 1998). The immobilized Acinetobacter sp.

strain W-17 on porous sintered glass was completely degraded 500 mg phenol l 1 in

40 hours, but free cells took 120 hours for the degradation of phenol at a similar

concentration (Beshay et al., 2002 and Abd-El-Haleem et al., 2003). Additionally,

phenol toxicity can be overcome by the immobilized bacterial cells. For example,

Bettmann and Rehm (1984) have reported that on immobilization of Pseudomonas sp.

on polyacrylamide hydrazide was able to degrade phenol at an initial concentration of


57 up to 2gL-1 in less than two days although the free cells did not grow at this

concentration.

Ying et al., (2007) reported that the immobilized cells of Acinetobacter sp.

strain PD12 have higher phenol degradation rate with wider pH (7.2 to 10) and

temperature (20 to 450C) range than that of free cells. The tolerance ability to acid

conditions of immobilized cells was much better than that of free cells. In the range of

20 to 350C, the immobilized cells showed a higher value of degradation rate than that

of free cells. Effect of temperature was less on immobilized cells than that of free

cells, because immobilization increased the thermal stability of the cells under the

protection of PVA carrier. Storage stability and reusability tests revealed that the

phenol degradation functions of immobilized cells were stable after reuse for 50 times

or storing at 4°C for 50 days. Thus the immobilized Acinetobncter sp. strain PD12

possesses a good application potential in the treatment of phenol-containing

wastewater.

Sodium alginate, a polysaccharide extracted from seaweed, performed

excellently in the removal of organic compounds from water. The fungus

Phanerochaete chrysosporium was immobilized in several polymer matrices such as

Ca-alginate, Ca-alginate–polyvinyl alcohol, and pectin, and was then used as a

biosorbent for removing 2,4-dichlorophenol (2,4-DCP) in wastewater (Juan and Han-

Qing, 2007). Calcium alginate-activated carbon composites were used for phenol

adsorption from aqueous solutions (Jodra and Mijangos, 2003).

Polyacrylamide and silica gels have been the most extensively used

immobilization materials for laboratory research studies. The latter are very suitable


58 for immobilization of microbial cells and such immobilized system integrates two

processes in one structure-effective biocatalysts and separation. P. Pictorrm was

immobilized with chitin because it can effectively biodegrade phenol and pH 6.8–7.0

was reported to be optimal for the biodegradation of the substrates (Farrell and Quilty,

2002; Zimmerman et al., 2005).

Immobilized cell particles are spherical. Each particle has an inner

homogeneous distribution of cells initially; most bacteria are present as micro

colonies in the porous surface area; cells are able to grow by consuming phenol.

Interfacial mass transfer resistance can be ignored when the external solution is

mixed. Intraparticle mass transfer is assumed to occur only through liquid in the

pores, and the effective diffusion coefficient of phenol is independent of concentration

(Trulleyove and Rulik, 2004).

Continuous degradation of phenol at an influent concentration of 100gL-1 with

immobilized P. putida was investigated by Mordocco et al., (1999) who pointed out

the significance of this low range of concentrations in light of the potential toxicity of

phenol at concentrations as low as 5 mg L-1. Comparing the performance of the

immobilized cells in calcium alginate beads to that of free cells, the superiority of the

immobilized cell system was more pronounced. A bead diameter between 1 and 2 mm

was found to be optimal for phenol degradation at low levels.

Physical entrapment of organisms inside a polymeric matrix has been

extensively used for whole-cell immobilization (Aksu and Bulbul, 1998; Annadurai et

al., 2008). The effectiveness of this method has also been investigated by El-Naas et


59 al., (2009) in a study to assess the biodegradation of phenol by P. putida immobilized

in PVA gel matrix at different conditions.

Jianlong et al., (2002) reported the development of a novel immobilization

carrier, that is, polyvinyl alcohol (PVA)-gauze hybrid carrier. It was found that

biodegradation rate of quinoline by the microorganisms immobilized on PVA-gauze

hybrid carrier was faster than that by the microorganisms immobilized in PVA gel

beads. There are various technologies for bacteria immobilization, including bead

entrapment, carrier binding, adsorption techniques, encapsulation, cell coating, and

film attachment (Chen et al., 2007).

The merit of immobilization is due to the high surface area available for

biofilm formation, which results in high biomass concentration of 30–40 g VSS L-1,

compared with 1.5–2.5 g VSS/l for activated sludge systems (Bajaj et al., 2008). In

addition, the systems with immobilized cultures are more stable to shock loadings

than the suspended cultures with free cells and immobilization can provide high

degradation capacity compared with free cells (Sheeja and Murugesan, 2002). Kim et

al., (2006) showed that calcium alginate immobilization of microbial cells effectively

increased the tolerance of P. putida MK1 to phenol and improved the degradation of

pyridine in a binary mixture of the two compounds.

2.10.1 Reusability of immobilized cells

Immobilization also provides better stability to the cells and hence can be

reused several times. El-Naas et al., (2009) had used PVA gel to immobilize

Pseudomonas putida cells for the biodegradation of phenol that has better mechanical


60 properties, and it is more durable than Ca-alginate which is biodegradable and can be

subject to abrasion. Karigar et al., (2006) had reported that Arthrobacter citreus cells

immobilized on an agar or agarose matrix could be used continuously for a week

without the losing their degradative ability.

Entrapment in insoluble Ca-alginate gel is a rapid, nontoxic, inexpensive,

versalite and the most often used method for immobilization of cells. More than 80%

of cell immobilization processes are still carried out using alginate (Thu et al., 1996).

The calcium alginate immobilized Nocardioides sp. NSP41 for the degradation of

PNP and phenol in industrial wastewaters is feasible because of the simultaneous

degradation of PNP and phenol. The immobilized cell culture showed higher

degradation rates than that the freely suspended cell culture. Furthermore, the

immobilized cells could be reused 12 times without losing their simultaneous

degradation of PNP and phenol industrial wastewaters (Cho et al., 2000).

Hence, with the aim to use the bacterial cells at a large scale level,

immobilization of the cells were also carried out in the current study.


2.0 literature review -...

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