Microbial Degradation of Phenols

Introduction

Organic pollutants are chemicals that are hazardous to human health. Most of them are extremely toxic at very low concentrations, persistent, can be transported over long range as air pollutants, bio-accumulate in human and animal tissue, and biomagnify in food chain.

Phenol and its derivatives are tox

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Organic pollutants are chemicals that are hazardous to human health. Most of them are extremely toxic at very low concentrations, persistent, can be transported over long range as air pollutants, bioaccumulate in human and animal tissue, and biomagnify in food chain.

Phenol and its derivatives are toxic aromatic hydrocarbon pollutants that have a hydroxyl group attached to their benzene ring structure. They have been used extensively for a number of industrial processes such as in the production of resins, coke, manufacturing plastics, colour, pesticides, pharmaceuticals, coal mines, steel and in aluminum industries. Phenol is also released from natural sources during the decay of lignocellulistic materials, from tannins and amino acid precursors (Abu-El-Haleem et al., 2003). These accounts for its increasing concentrations in the environment as wastewaters from these industries are not properly treated before discharge. It is classed as a priority compound (EPA, 2009) because of its toxicity at very low concentrations and possible accumlation in the environment (Shokoohi et al., 2006). Concentrations of about 1 mg are known to be toxic to some species of aquatic organisms and even lower concentrations cause problems of taste and odour in drinking water (Nair, 2008).

Organic pollutants are chemicals that are hazardous to human health. Most of them are extremely toxic at very low concentrations, persistent, can be transported over long range as air pollutants, bioaccumulate in human and animal tissue, and biomagnify in food chain.

Phenol and its derivatives are toxic aromatic hydrocarbon pollutants that have a hydroxyl group attached to their benzene ring structure. They have been used extensively for a number of industrial processes such as in the production of resins, coke, manufacturing plastics, colour, pesticides, pharmaceuticals, coal mines, steel and in aluminum industries. Phenol is also released from natural sources during the decay of lignocellulistic materials, from tannins and amino acid precursors (Abu-El-Haleem et al., 2003). These accounts for its increasing concentrations in the environment as wastewaters from these industries are not properly treated before discharge. It is classed as a priority compound (EPA, 2009) because of its toxicity at very low concentrations and possible accumlation in the environment (Shokoohi et al., 2006). Concentrations of about 1 mg are known to be toxic to some species of aquatic organisms and even lower concentrations cause problems of taste and odour in drinking water (Nair, 2008).

Toxic aromatic hydrocarbon pollutants that have a hydroxyl group attached to their benzene ring structure. They have been used extensively for a number of industrial processes such as in the production of resins, coke, manufacturing plastics, colour, pesticides, pharmaceuticals, coal mines, steel and in aluminum industries. Phenol is also released from natural sources during the decay of lignocellulistic materials, from tannins and amino acid precursors (Abu-El-Haleem et al., 2003). These accounts for its increasing concentrations in the environment as wastewaters from these industries are not properly treated before discharge. It is classed as a priority compound (EPA, 2009) because of its toxicity at very low concentrations and possible accumlation in the environment (Shokoohi et al., 2006). Concentrations of about 1 mg are known to be toxic to some species of aquatic organisms and even lower concentrations cause problems of taste and odour in drinking water (Nair, 2008).

Various methods have been employed in the treatment of phenol in industrial wastewater. They include chemical oxidation, solvent extraction, adsorption, and incineration. The high cost for treatment and the increased potential of forming even more persistent and hazardous by-products are the major problems associated with using these methods (Loh et al., 2000). Biological processes remains a safer means of removing phenol from wastewater because of the complete minieralization of the compound (Movahedyan, et al., 2009). It is also cost effective in that those organism that can degrade phenol also utilize it as a carbon and energy source. Certain microorganisms capable of degrading phenol in wastewater are inhibited by high concentrations as a result, the rate of degradation is low (Abu-El-Haleem et al., 2003).

The role of bacterial communities in the success of most bioremediation activity has led to the increased phenotypic and genetic analysis of communities to find common or wide-spread degraders. In other to optimize suitable conditions necessary for biodegradation of phenols, identification of microorganisms from natural environment endowed with this capacity is required. Conventional culture based methods are time consuming, very low in sensitivity and can only be used in isolating 1% of the total environmental samples (Movahedyan, et al., 2009). Some researchers have suggested that enrichment methods be applied to mixed microbial populations in order to increase the activity of the cells to degrade phenol (Yang and Lee, 2007). Although the effectiveness of culture dependent methods have been debated, they are still indispensible when detailed analysis of bacterial group is required (Watanabe et al., 1998).

Molecular tools such as polymerase chain reaction (PCR), denaturing gradient gel electrophoresis (DGGE), amplified ribosomal DNA restriction analysis ARDRA, Terminal-Restriction fragment length ploymorphism (T-RFLP) have been applied recently to study changes observed in microbial communities as a result of pressures within an enrichment process. (Watanabe, 1998; Yand and Lee, 2007). The advantage of these molecular methods is that they have higher specificity and sensitivity, less time consuming, and larger population of organisms are observed at once (Movahedyan, et al., 2009).

Aim and Objectives

The aim of this research is to identify and compare the phenol degrading capacity of bacterial communities from different activated sludge plants.

Objectives

1.  To determine the common bacterial communities (and their relative abundance) capable of degrading phenol from the different activated sludge.
2. To evaluate the effect of temperature on the variability of the bacterial communities and phenol degradation.
3. To determine the effect of immigration on the structure of bacterial communities.
4. To ascertain the effect of inocula enrichment on variability within the bacterial community.

Regulation of chemicals

Regulation of chemicals became more prominent in the 1960’s with the global realization that chemicals can cause irreversible harm to human health and the environment. In determining the effect a chemical has on the environment, it is necessary to assess the chemical’s fate in the environment, in particular, its persistence and bioaccumulation potential. These assessments are done by laboratory testing (Goodhead, 2009). A number of laboratory standard methods, such as the International Organisation for Standardisation (ISO standards), United States Environmental Protection Agency (USEPA) testing guidelines, and the Organisation for Economic Cooperation and Development (OECD) have been developed to assess the biodegradability of chemicals. These tests are used to predict the effect of biodegradation on the fate and transport of these chemicals in the environment (Paixao et al, 2006).

The OECD guideline is the most widely accepted guidance document for the testing of chemicals. It provides guidelines for determining the fate and effect of a chemical in the environment and the probability of the chemical to undergo degradation in the environment. These testing procedures consist of a series of standardized tests starting with the screening of chemicals to determine their degradability. The first part of these tests is the screening test also called the ready biodegradability test. In RBTs, the test substance serves as the sole carbon source, which is diluted in a medium containing a relatively low concentration of biomass, incubated at 300C for 28 days. This is to allow for sufficient time for the organisms to adapt to the test chemical (OECD, 1993).

In ready biodegradability tests, the basic factors which affect the reliability of the results are the inoculum, the source of the microorganisms for the test, and its state of acclimatization and adaptation (Paixao et al., 2006). Although issues such as inoculum pre-treatment have been addressed in a number of studies carried out by Vazquez-Rodriquez et al. (2007), Goodhead (2009) and Paixao, et al (2006), an important factor that has not been studied so much is the incubation temperature. Currently, RBTs are carried out at 30oC; this also might have its own effect on the variability of the bacterial community involved in the biodegradation process. It is common knowledge that temperature slows down the rates of chemical and biological reactions in living organisms. Changes in temperature affect the composition of the bacterial population (Erdal and Randall, 2002) and may also affect the efficiency and kinetics of the degradation process.

Biodegradation of phenols

Microbial degradation is the most dominant elimination mechanism of organics from the environment. Persistence of xenobiotics represents a real risk for ecosystems as well as for human beings. Therefore, the knowledge of the biodegradability of these xenobiotics is one of the most important aspects of understanding their behaviour in the natural environment and during the biological treatment of wastewater

To date, a number of phenol-degrading bacteria have been isolated, and their phenol degradative pathways have been studied. Aerobic degradation of a phenolic compound is known to be initiated by its hydroxylation to form corresponding catechols (Harayama et al. 1992). This step is catalyzed by phenol hydroxylase (phenol 2- monooxygenase, EC 1.14.13.7), which is considered to be the rate-limiting step in the degradative pathway (Hino et al. 1998). Two types of bacterial phenol hydroxylases, the single-component type and multicomponent type, are known; among them, multicomponent phenol hydroxylase (mPH) is considered to be the major enzyme in the natural environment (Peters et al. 1997; Watanabe et al. 1998; Futamata et al. 2001). Several genes coding for mPHs have been cloned and sequenced from phenol-degrading bacteria . All these mPHs are similar in their enzyme structure; they comprise six subunits, among which the catabolic site exists within the largest subunit (approx. 60 kD). Some of these enzymes have been found to exhibit different substrate specificity for substituted phenols (Teramoto et al. 1999). To date, a number of phenol-degrading bacteria have been isolated, and their phenol degradative pathways have been studied.

Aerobic degradation of a phenolic compound is known to be initiated by its hydroxylation to form corresponding catechols (Harayama et al. 1992). This step is catalyzed by phenol hydroxylase (phenol 2- monooxygenase, EC 1.14.13.7), which is considered to be the rate-limiting step in the degradative pathway (Hino et al. 1998). Two types of bacterial phenol hydroxylases, the single-component type and multicomponent type, are known; among them, multicomponent phenol hydroxylase (mPH) is considered to be the major enzyme in the natural environment (Peters et al. 1997; Watanabe et al. 1998; Futamata et al. 2001). Several genes coding for mPHs have been cloned and sequenced from phenol-degrading bacteria.

All these mPHs are similar in their enzyme structure; they comprise six subunits, among which the catabolic site exists within the largest subunit (approx. 60 kD). Some of these enzymes have been found to exhibit different substrate specificity for substituted phenols (Teramoto et al. 1999). (Watanabe, 2002)

Microbial degradation is the most dominant elimination mechanism of organics from the environment. Persistence of xenobiotics represents a real risk for ecosystems as well as for human beings. Therefore, the knowledge of the biodegradability of these xenobiotics is one of the most important aspects of understanding their behavior in the natural environment and during the biological treatment of wastewater.

References

1.     EPA, (2009). National Recommended Water Quality Criteria. Office of Water (4304T) United States Environmental Protection Agency. Washington, DC. pp 4-17. <http://water.epa.gov/scitech/swguidance/waterquality/standards/current/upload/nrwqc-2009.pdf> Accessed on: 14th March 2011.
2.      Erdal, U. G. and Randall, C. W. (2002). The effects of temperature on system performance and bacterial community structure in EBPR systems. Enviro 2002/IWA 2nd World Water Congress, Melbourne, Australia.
3.      Goodhead A. K. (2009). Towards Rational Risk Assessment: Improving biodegradation Tests through an Understanding of Microbial Diversity
4.       Loh, K. C., Chung, T. S., and Ang, W. F. (2000). Immobilized-cell membrane bioreactor for high-strength phenol wastewater. J. Environ. Eng.-ASCE, 126 (1),75-79.
5.       Nair, C. I., Jayachandran, K., and Shashidhar, S. (2008). Biodegradation of Phenol. African Journal of Biotechnology Vol. 7 (25): 4951-4958. <http://www.academicjournals.org/AJB> Accessed on: 11th March, 2011.
6.       Paixao, S. M., Saagua, M. C., Tenreiro, R., Anselmo, A. M. (2006). Biodegradability Testing Using Standardized Microbial Communities as Inoculum. Environmental toxicology. 21(2): 131-140.
7.       Sloan WT, Lunn, M., Woodcock, S., Head, I., Nee, S. and Curtis, T. P. 2006) Quantifying the roles of immigration and chance in shaping prokaryote community structure. Environ Microbiol 8:732–740.
8.       Watanabe, K., Teramoto, M., Futamata, H., and Harayama, S. (1998). Molecular detection, isolation, and physiological characterization of functionally dominant phenol-degrading bacteria in activated sludge. Appl. Environ. Microbiol. Vol. 64, 4396-4402

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