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Department of Environmental Systems Engineering, Nagaoka University of Technology, Kamitomioka, Nagaoka, Niigata, Japan
Katsumasa Abe
Department of Environmental Systems Engineering, Nagaoka University of Technology, Kamitomioka, Nagaoka, Niigata, Japan
Yoshio Kera
Department of Environmental Systems Engineering, Nagaoka University of Technology, Kamitomioka, Nagaoka, Niigata, Japan
*Address all correspondence to:
1. Introduction
1.1. Flame retardants
Flame retardants (FRs) are chemicals used in polymers to protect the public from accidental fires by preventing or retarding the initial phase of a developing fire (EFRA, 2007). These chemicals are now found in numerous consumer products, including construction materials, upholstery, carpets, electronic goods, furniture and also children’s products such as car seats, strollers and baby clothing. FRs have become indispensable to modern life, and have saved numerous lives by preventing unexpected fires across the globe.
FRs are divided into two general classes based on their relation to host polymers: additive and reactive FRs (WHO, 1997). Additive FRs are simply mixed with host polymers. The lack of chemical bonding between the FRs and host polymers enables the FRs to leach out of or volatilize from host polymers over time into the ambient environment. Reactive FRs are incorporated into host polymers by covalent bonding into the polymer backbone, and are thus less likely to leach into the environment. Additive FRs are mainly used in thermoplastics, textiles and rubbers, whereas reactive FRs are usually used in thermoset plastics and resins (SFT, 2009a).
FRs are sub-divided into six groups characterized by their chemical composition: 1) aluminum hydroxide, 2) brominated, 3) organophosphorus, 4) antimony oxides, 5) chlorinated and 6) other FRs. These groups account for 40%, 23%, 11%, 8%, 7% and 11% of the annual FR global consumption in 2007, respectively (Beard & Reilly, 2009). The total market for FRs in the United States, Europe and Asia in 2007 amounted to about 1.8 million tons.
1.2. Organophosphorus flame retardants
Organophosphorus flame retardants (PFRs) are based primarily on phosphate esters, phosphonate esters and phosphite esters. The total consumption of FRs in Europe was an estimated 465,000 tons in 2006, of which 20% comprised PFRs (KLIF, 2010). Of the PFRs consumed, 55% were chlorinated. Halogenated PFRs are the preferred form of FRs because halogen inhibits flame formation in organic materials, and non-halogenated PFRs are typically used as flame-retardant plasticizers (KLIF, 2010).
1.3. Tris(1,3-dichloro-2-propyl) phosphate and tris(2-chloroethyl) phosphate
Tris(1,3-dichloro-2-propyl) phosphate (TDCPP) and tris(2-chloroethyl) phosphate (TCEP) are typical examples of additive chlorinated PFR (Fig. 1 and Table 1).
Figure 1.
Chemical structure of tris(1,3-dichloro-2-propyl) phosphate (TDCPP) and tris(2-chloroethyl) phosphate (TCEP)
TDCPP is a viscous colorless to light yellow liquid and is produced by the epoxide opening of epichlorohydrin in the presence of phosphorus oxychlorine (ATSDR, 2009). TDCPP is used primarily in flexible polyurethane foams but also in rigid polyurethane foams, resins, plastics, textile coatings and rubbers (California EPA, 2011). TDCPP was a common ingredient of sleepwear for children in the 1970s, but was voluntarily withdrawn by manufactures in 1977 because of its proven mutagenicity (California EPA, 2011). However, the PFR can still be found in many baby products (Stapleton et al., 2011). Currently, TDCPP is used mostly in flexible polyurethane foams for upholstered furniture and automotive products. TDCPP consumption has increased following the ban on common FR polybrominated diphenyl ethers (PBDEs). Consequently, total TDCCP production has increased, being an estimated 4,500-22,700 tons in the United States in 2006 and <10,000 tons in Europe in 2000 (van der Veen & de Boer, 2012).
TCEP is colorless to pale yellow liquid and is highly soluble in water (Fig. 1 and Table 1). The compound is chemically synthesized via condensation of phosphorus oxychloride and chloroalkyl alcohol at low temperatures and pressures to avoid formation of alkyl chlorides (ATSDR, 2009). Previously, the main purpose of TCEP was to reduce the brittleness of flame-resistant rigid or semirigid polyurethane foams. More recently, it has been used as a flame-retarding plasticizer and viscosity regulator in unsaturated polyester resin (accounting for around 80% of current use) (EURAR, 2009). TCEP-containing polymers are commonly used in the furniture, textile and building industries (for example, more than 80% of the TCEP consumption in the EU is invested in roofing insulation). TCEP is also used in car, railway and aircraft materials, and in professional paints. Since the 1980s, TCEP has been progressively replaced by other flame retardants, primarily tris(1-chloro-2-propyl) phosphate (TCPP). Consequently, global consumption of TCEP in the EU, which exceeded 9,000 tons in 1989, declined to below 4,000 tons by 1997. TCEP is no longer produced in the EU (EURAR, 2009).
General aspect of Tris(1,3-dichloro-2-propyl) phosphate (TDCPP) and tris(2-chloroethyl) phosphate (TCEP)
1.4. Occurrence and behavior of TDCPP and TCEP in the environment
TCEP and TDCPP have been detected in various environments worldwide, including indoor and outdoor air, surface and ground waters, and even drinking water (Tables 2 and 3). It is unlikely that these compounds are produced naturally. Their environmental presence is thus considered to be the result of human activity. Because these PFRs are physicochemically and microbiologically stable in the environment and are also reportedly toxic, they are a serious threat to human and ecosystem health.
1.4.1. TDCPP
Detected air concentrations of TDCPP have attained up to150 ng m3-1 in Sweden houses, and in Belgium office and stores, they have reached 73 ng m3-1 (Table 2). In outdoor air, TDCPP levels near a main road in Sweden ranged from <0.04-0.072 m3-1, and significant amounts have been detected globally in air borne particles over the Pacific, Indian, Arctic and Southern Oceans. TDCPP has been also found in indoor dust at relatively higher concentrations. Levels of TDCPP have tended to be higher in public buildings than in domestic buildings.
With respect to water environments, TDCPP concentrations have been detected at up to ~50 ng L-1 in German rivers and at 1,335 ng L-1 in Italian lakes. In these countries, it also occurs in rain and/or snow, as a result of volatilization from host materials. A much higher TDCPP concentration was detected in raw water at a disposal site in Japan, suggesting that the compound leaches and migrates to water sources. In the United States and Germany, TDCPP has even been detected in drinking water processed in treatment plants (DWTs). Relatively higher concentrations of TDCPP occur in landfill site sediments. Much higher concentrations still have been found in sediments near a car demolition site in Norway.
TDCPP has been also detected in the effluents of sewage treatment plants (STPs) and waste water treatment plants (WWTPs) in European countries and Japan, revealing that effluents are a source of aquatic TDCPP contamination. Comparable levels have been observed in the influents, indicating that the compound persists in the treatment plants. Degradation of TDCPP in the environment has been reported as low. Together, these observations suggest that TDCPP is likely to accumulate in the environment.
TDCPP has also been detected in biological samples, including fishes, mussels and birds. In Norway, fishes and mussels were observed to contain up to 8.1 and 30 ng g-1 of TDCPP, respectively. In bird blood/plasma and eggs respectively, TDCPP levels range from <0.11-0.16 and from <0.72-1.9 ng g-1. In Sweden, freshwater fishes close to emission sources contained 49-140 ng g-1 TDCPP. Worryingly, TDCPP has also been detected in the breast milk of Swedish women.
1.4.2. TCEP
In Sweden, the highest detected air concentration of TCEP was 730 ng m3-1 inside an office furnished with linoleum floor and a new photocopier (Table 3). In outdoor air, it can reach 6.2 ng m3-1 beside a main road, but remote areas harbor less than 0.2 ng m3-1, implicating road traffic as an important source of TCEP emission. TCEP has also been detected globally in air borne particles over the Pacific, Indian, Arctic and Southern Ocean. In Belgium, indoor dust can contain up to 260 μg g-1 TCEP. TCEP concentrations in dusts of public spaces tend to exceed those in domestic dusts.
TCEP ranges from <3.0-1,236 ng L-1 in German rivers, lakes and reservoirs. In this country and in Italy, it has also been detected in rain and/or snow, indicating that, like TDCPP, TCEP volatilizes from its host materials. Groundwater TCEP levels up to 754 ng L-1 have been reported in Germany, suggesting that TCEP primarily mobilizes into water rather than attaching to soil. TCEP also occurs in drinking water or finished water from DWTs; recorded concentrations are as high as 99, 25 and 1.7 ng L-1 in the United States, Korea and Germany, respectively. Much higher concentrations have been observed in raw water of waste disposal sites in Japan. Relatively higher concentrations of TCEP have been detected in landfill site sediments in Japan and Norway (up to 7,400 and 380 μg kg-1, respectively). Especially high concentrations were found in the sediment nearby a car demolition site.
TCEP has been also detected in STP or WWTP effluents in many countries. Comparable levels of TCEP are observed in the influents. These observations demonstrate that, like TDCPP, TCEP persists in the treatment plants.
Also similarly to TDCPP, TCEP has been detected in biological samples, including fishes, crabs, mussels and birds. In Norway, fishes and mussels respectively contain up to 26 and 23 ng g-1 TCEP. In birds and their eggs, TCEP levels can reach up to 6.1 ng g-1. In fishes residing near emission sources in Sweden, they reach up to 69 and 160 ng g-1 respectively. Furthermore, like TDCPP, TCEP has been detected in the breast milk of Swedish women.
Since the toxic effects of TCEP and TDCPP have been regarded as marginal compared to those of PBDEs, they have been extensively used. However, their non-negligible toxicities have been revealed in a number of studies (Tables 4 and 5). Together with their persistence in the environment, the environmental contamination of both compounds has become of serious concern.
1.5.1. TDCPP
Rats given oral doses of TDCPP absorb more than 90% of the compound within 24 h, with the highest concentrations being observed in kidney, liver and lung (EURAR, 2008). The acute toxicity of oral TDCPP has been reported as low, with LD50 values ranging from 2,250 mg kg-1 for female mice to 6,800 mg kg-1 for male rabbits (Table 4). In a 2-year chronic toxicity study in rats, the lowest observable adverse effect level (LOAEL) was 5 mg kg-1 day-1. In that study, statistically significant relationships between TDCPP dose and tumor incidences were observed in both male and female rats. Consequently, TDCPP is today classified as Carc. Cat. 3; R40 and Cat. 2; H351, denoting “limited evidence of a carcinogenic effect” and “suspected of causing cancer”, respectively.
A number of TDCPP genotoxicity studies have been conducted in whole mammals that have resulted in negative conclusions regarding genotoxicity (Albemarle Corp. & ICL North America Inc., 2011). However, in vitro studies using bacteria and mammalian cells have suggested that TDCPP exerts genotoxic effects, and an in vivo study showed its covalent binding to DNA (US EPA, 2005; Morales & Matthews, 1980).
Similarly, neurotoxicity studies of TDCPP involving hens and rats reveal no clear evidence that TDCPP is neurotoxic. However, a study based on undifferentiated and differentiating PC12 cells showed its potential neurotoxicity (Dishaw et al., 2011).
Whether, and to what extent, TDCPP is toxic to humans remains unknown. However, TDCPP has been shown to alter sex hormone balance in human cell lines, via alteration of steroidogenesis or estrogen metabolism (Liu et al., 2012). In addition, TDCPP concentrations in house dusts have been linked to altered hormone levels and decreased semen quality in men (Meeker & Stapleton, 2010).
TDCPP is regarded as toxic to aquatic organisms (EURAR, 2008). An acute toxicity study on fish trout yielded an LC50 value of 1.1 mg L-1. Acute and chronic toxicity studies conducted on the invertebrate Daphnia produced an EC50 value of 3.8 mg L-1. In a chronic study on the alga Pseudokirchneriella, ErC10 (10% growth-rate inhibition) was recorded as 2.3 mg L-1. Thus, TDCPP is classified as N; R51/53, denoting “Toxic to aquatic organisms, may cause long-term adverse effects in the aquatic environment”. In addition, an LC50 of 23 mg kg-1 has been reported for a terrestrial organism, the earthworm Eisenia.
Toxicity
Organism
Reference
Acute toxicity
LD50=6,800 mg kg-1 male rabbit
US EPA, 2005
LD50=3,160 mg kg-1 male rat
EURAR, 2008
LD50=2,670 mg kg-1 male mice
LD50=2,250 mg kg-1 female mice
LD50=2,236 mg kg-1 male rat
LD50=2,489 mg kg-1 female rat
Chronic toxicity
LOAEL=5 mg kg-1 day-1 rat for hyperplasia and convoluted tubule epithelium
LOEC=1.0 mg L-1 Daphnia for reproduction (21 days)
NOEC=0.5 mg L-1 Daphnia for reproduction (21 days)
ErC10=2.3 mg L-1 algae
LC50=23 mg kg-1 earthworm Eisenia
NOEC=2.9 mg kg-1 earthworm Eisenia for reproduction
NOEC=17 mg kg-1 plant Mustard
Alter hormone levels
human and zebra fish cells
Liu et al., 2012
Decreased sperm quality
human
Meeker & Stapleton, 2010
Table 4.
Toxicological information of TDCPP
1.5.2. TCEP
Rats given oral doses of TCEP absorb over 90% of the compound within 24 h, with marked accumulations in liver, kidney, fat and the gastrointestinal tract (EURAR, 2009). In animals, TCEP appears to be mainly toxic to brain, kidney and liver. Toxicity studies have implicated TCEP as moderately toxic; in rats, oral administration yields an LD50 of 430-1,230 mg kg-1 and skin contact reveals a low acute dermal toxicity (LD50 >2,150 mg kg-1) (Table 5). A 2-year chronic toxicity study of TCEP yielded LOAELs of 44 mg kg-1 day-1 in rats and 175 mg kg-1 day-1 in mice. The same study indicated that TCEP is potentially neurotoxic, with no observed adverse effect levels (NOAELs) in rats and mice being 88 mg kg-1 day-1 and 175 mg kg-1 day-1, respectively.
Toxicity
Organism
Reference
Acute toxicity
LD50=430-1,230 mg kg-1 rat
EURAR, 2009
LD50>2,150 mg kg-1 rat for dermal
EURAR, 2009
Chronic toxicity
LOAEL=44 mg kg-1 day-1 rat for kidney lesions (2 years)
EURAR, 2009
LOAEL=175 mg kg-1 mouse for kidney morphology (2 years)
EURAR, 2009
Neurotoxicity
rat and mouse
EURAR, 2009
NOAEL=88 mg kg-1 day-1 rats (16 weeks by gavage)
NOAEL=175 mg kg-1 day-1 mouse (16 weeks by gavage)
Reproductive toxicity
rat and mouse
EURAR, 2009
NOAEL=175 mg kg-1 day-1 mouse for fertility
Carcinogenicity
rat and mouse
SCHER, 2012
Toxic to aquatic organisms
killifish, trout and goldfish
EURAR, 2009
Alter sex hormone balance
human cells and Zebra fish
Liu et al., 2012
Alter cell cycle regulatory protein expression
rabbit renal proximal tubule cells
Ren et al., 2008
Table 5.
Toxicological information of TCEP
In the 2-year study, increased incidences of adenomas and carcinomas were linked to TCEP exposure, revealing TCEP as a potential carcinogen (EURAR, 2009). TCEP is thus classified as Carc. Cat. 3; R40. Because TCEP additionally exhibits reproductive toxicity in rats and mice, it is also classified as Repr. Cat. 2; R60, denoting “may impair fertility”. TCEP at environmental concentrations has been reported to affect the expression of cell cycle regulatory genes in primary cultured rabbit renal proximal tubule cells (Ren et al., 2008).
TCEP is toxic to aquatic organisms, being classified as N; R51/53 (EURAR, 2009). Short term exposure to TCEP is mildly-moderately adverse to the aquatic invertebrate organisms Daphnia and Planaria, and TCEP presents low acute toxicity to killifish, trout and goldfishes.
The toxic effects of TCEP in humans are largely unknown. However, neurotoxic signs have been reported in a 5-year old child who slept in a room with wood paneling containing 3% TCEP (Ingerowski & Ingerowski, 1997). In addition, an epidemiological study of children in school environments found a potential association between the TCEP content in air-bone dusts and impaired cognitive ability (UBA, 2008). TCEP has been further reported to alter the sex hormone balance in human cells, as well as in fish cells.
1.6. Removal technique for TDCPP and TCEP
The persistence of chlorinated FRs TCEP and TDCPP in current waste water and drinking water treatment processes has accelerated the investigation of alternative water treatment techniques that will dispel these compounds.
Echigo et al. showed that TDCPP in distilled water and an effluent from a solid waste landfill site is effectively degraded by O3/vacuum UV or O3/H2O2 process, although degradation products were not determined in this study (Echigo et al., 1996). Westerhoff et al. reported that >20% of approximately 30 ng L-1 of TCEP in surface water samples can be removed with powdered activated carbon, but that other adsorptive processes, metal salt coagulation and lime softening, and oxidative processes (chlorination and ozonation) are ineffective (Westerhoff et al., 2005). Lee et al. showed that > 90% removal efficiency of 100 µg L-1 of TCEP in river and sea waters is possible using tight nanofiltration membranes with a low molecular weight cutoff of approximately 200 (Lee et al., 2008). Watts et al. demonstrated that the higher removing efficacy (> 95%) of 5 mg L-1 of TCEP in a water is achieved by a UV/H2O2 advanced oxidation process with the highest UV fluence at 6,000 mJ cm-2 (Watts & Linden, 2008). In this study, the generation of stoichiometric amount of chloride ion was observed. In addition, Benotti et al. reported that UV/TiO2 supplemented with H2O2 can decrease the concentration of TCEP in a river water, although the degradation was not so effective and not completed (Benotti et al., 2009).
2. Microbial degradation and detoxification of TDCPP and TCEP
FRs have been widely distributed commercially and are necessary to prevent or reduce mortality from accidental fires. However, the leaching of additive FRs has led to global contamination of the environment. The chlorinated PFRs TCEP and TDCPP persist in the environment and exhibit varying toxic effects, raising concerns about their effects on human and ecological health. Although several physicochemical methods for removing TCEP and TDCPP have been reported (as described above), biotechnological techniques offer an attractive alternative, being potentially cost-effective, eco-friendly and enabling in situ remediation of contaminants. However, prior to recent isolation of TCEP- and TDCPP-degrading bacteria by our group, no biological degrading agent for such compounds was known.
2.1. Isolation and characterization of TDCPP- and TCEP-degrading bacteria
2.1.1. Enrichment of TCEP and TDCPP-degrading bacteria
2.1.1.1. Enrichment cultivation of TCEP and TDCPP-degrading bacteria
To obtain microorganisms that can degrade TDCPP and TCEP, we used an enrichment culture technique in which one of TDCPP or TCEP served as the sole phosphorus source (Takahashi et al., 2008). Forty six environmental samples (soils and sediments) in Japan were cultivated at 30°C in minimal medium containing approximately 20 µM of each compound. Significant degradation of TCEP and TDCPP was seen in ten and three of the samples, respectively. In the first cultivation round, each compound had disappeared within 2 to 5 days; successive sub-cultivations reduced the degradation time to within one day. The enrichment cultures displaying the highest degradation efficacy against TCEP and TDCPP were designated 67E and 45D, respectively. Culture 67E completely degraded 20 µM of TDCPP in 3 h and TCEP in 6 h (Fig. 2A and B), while culture 45D completely degraded the same concentration of TDCPP in 3 h and TCEP in 24 h. During the degradations, 2-CE was liberated from TCEP and 1,3-DCP from TDCPP, indicating that the degradation pathway involved hydrolysis of phosphoester bonds.
Figure 2.
Degradation of TCEP (A) and TDCPP (B) by enrichment cultures. The enrichment cultures, 67E (circles) and 45D (triangles), were cultivated on 20 µM of TCEP or TDCPP as the sole phosphorus source.
2.1.1.2. 2-CE and 1,3-DCP degradation ability of enrichment cultures
The metabolites 2-CE and 1,3-DCP are also persistent and toxic: 1,3-DCP is a known genotoxin and carcinogen (NTP & NIEHS, 2005), while 2-CE exhibits genotoxicity, fetotoxicity and cardiotoxicity (National Toxicology, 1985). We analyzed whether the cultures can degrade the metabolites by measuring chloride ion formation. Cultures 67E and 45D liberated chloride ions from 2-CE and 1,3-DCP, respectively. After 120 h reaction, the proportion of chloride ion was approximately 100% and 68.5% of the total chlorine contained in the supplied 2-CE and 1,3-DCP, respectively. This shows that both cultures can dehalogenate their respective chloroalcohols and can therefore potentially detoxify chlorinated PFRs in the environment.
Figure 3.
Effect of exogenous phosphate on the degradation of TCEP (A) and TDCPP (B) and the chloride ion formation from TCEP (C) and TDCPP (D). The enrichment cultures, 67E (A and C) and 45D (B and D), were cultivated on 20 µM of TCEP or TDCPP as the sole phosphorus source, respectively, with various concentrations of inorganic phosphate (NaH2PO4): 0 mM (closed circles), 0.02 mM (closed triangle), 0.2 mM (closed squares) and 2 mM (closed diamonds). Control culture without cell inoculation is indicated by open circles. Each data point represents the mean of at least two independent determinations.
2.1.1.3. Effect of exogenous phosphate on the degradation ability of enrichment cultures
Phosphate-sufficient conditions are well known to repress the expression of genes involved in phosphorus utilization. We thus examined the effect of exogenous inorganic phosphate on TDCPP and TCEP degradations and chloride ion formation (Fig. 3). At concentrations of 0.02, 0.2 and 2 mM, exogenous inorganic phosphate did not significantly inhibit TCEP and TDCPP degradation by the respective cultures (Fig. 3A and B), but chloride ion formation was enhanced at concentrations up to 0.2 mM (Fig. 3C and D). From these results, we concluded that efficient PFR detoxification could be achieved by optimizing the inorganic phosphate concentration.
2.1.1.4. Bacterial communities of enrichment cultures
To profile the bacterial communities in the cultures, we performed denaturing gradient gel electrophoresis (DGGE) analysis (Fig. 4). In the absence of inorganic phosphate, two bands (C1 and C2) were observed in the fingerprint of TCEP-supplemented 67E, which persisted throughout cultivation (Fig. 4A). With inorganic phosphate added, the intensity of C2 markedly decreased at later incubation stages (Fig.4A). In 45D supplemented with TDCPP, a single band (D3) was observed at the beginning of cultivation, but at later times two additional bands (D1 and D2) appeared, regardless of the presence or absence of inorganic phosphate (Fig. 4B). However, with inorganic phosphate added, the intensity of D2 and D3 decreased while that of D1 increased at the late stage of cultivation (Fig. 4B). The nucleotide sequence of C1 and D1 was affiliated with the genus Acidovorax, that of D2 with the genus Aquabacterium, and C2 and D3 were assigned to the genus Sphingomonas (Table 6). Together with the effect of exogenous inorganic phosphate on chlorinated PFRs degradation with liberation of chloride ions, these results imply that the Sphingomonas-related bacteria hydrolyze the PFRs, and that the Acidovorax-related bacteria dehalogenate the chloroalcohols. Among these bacterial genera, a strain of Sphingomonas sp. has been reported to hydrolyze some organophosphate pesticides, such as chlorpyrifos (Li et al., 2007). However, bacteria that are known to dehalogenate the chloroalcohols were not identified in the enrichment cultures, suggesting that a new member, possibly Acidovorax sp., is responsible for dehalogenating the chloroalcohols in the cultures.
Culture
Band
Phylogenetic affiliation
Species
67E
C1
Acidovorax sp.
C2
Sphingomonas sp.
45D
D1
Acidovorax sp.
D2
Aquabacterium sp.
D3
Sphingomonas sp.
Table 6.
Phylogenetic affiliation of microorganisms represented by bands in DGGE profiles of the enrichment cultures 67E and 45D.
Figure 4.
DGGE profile of the enrichment cultures 67E (A) and 45D (B) during cultivation in the presence of absence of inorganic phosphate. The arrowheads indicated the DNA fragments sequenced.
2.1.2. Isolation and characterization of TDCPP- and TCEP-degrading bacteria
2.1.2.1. Isolation of TDCPP- and TCEP-degrading bacteria
We attempted to isolate the bacteria responsible for degrading TDCPP and TCEP in the cultures 67E and 45D. (Takahashi et al., 2010). In the case of 45D, isolation was achieved by limiting dilution method. The culture was repeatedly serially diluted in a minimal medium containing 20 µM of TDCPP and cultivated at 30°C. Finally, the culture was spread onto a minimal agar plate containing 232 µM of TDCPP as the sole phosphorus source. A single colony grown on the plate was named strain TDK1 (Fig. 5A). In the case of 67E, the culture was spread onto a minimal agar plate containing 232 µM of TCEP as the sole phosphorus source and incubated at 30°C. Single colonies were then cultivated in a minimal medium containing 20 µM of TCEP as the sole phosphorus source. This isolation procedure was repeated three times, and a single colony was named strain TCM1 (Fig. 5B).
2.1.2.2. Identification of TDCPP- and TCEP-degrading bacteria
Both strains were short-rod-shaped bacteria (0.8-1.0 × 1.0-2.5 µm) and produced yellow, circular, convex colonies with smooth, glistening surfaces on a nutrient agar plate. As carbon sources, both strains assimilated glucose, maltose and L-arabinose; in addition, strain TCM1 assimilated potassium gluconate, while strain TDK1 assimilated D-mannose, N-acetyl-D-glucosamine, and D, L-malate. Both strains tested negative for indole, urease, arginine dihydrolase, nitrate reduction, gelatine hydrolysis, and glucose fermentation, and were positive for esculin hydrolysis. TCM1 and TDK1 tested negative and positive for cytochrome oxidase, respectively. The morphological and physiological characteristics of the strains were similar to those of Sphingomonas spp. Furthermore, the 16S rRNA gene sequence of the strains is closely related to those of sphingomonads, comprising the genera Sphingomonas, Sphingobium, Novosphingobium and Sphingopyxis (Takeuchi et al., 2001). The phylogenetic tree constructed from the sequences of these genera showed that strains TCM1 and TDK1 belong to Sphingobium and Sphingomonas, respectively
Figure 5.
SEM micrographs of TCEP- and TDCPP-degrading bacteria Sphingobium sp. strain TCM1 (A) and Sphingomonas sp. stain TDK1 (B).
2.1.2.3. Degradation ability of TCEP and TDCPP-degrading bacteria
Both strains completely degraded 20 µM of TDCPP within 6 h (Fig. 6A and B). Strain TDK1, however, was 48 times less effective in degrading TCEP than TCM1 (TCEP degradation time was 144 h for TDK1, versus 3 h for TCM1) (Fig. 6A and B). During the degradations, 1,3-DCP and 2-CE were detected in the cultures of both strains and were not further degraded (Fig. 6C and D). These results showed that the strains degrade the compounds by hydrolyzing their phosphotriester bonds. To date, TCM1 and TDK1 are the only isolated microorganisms reported to degrade the persistent PFRs.
We then analyzed whether the strains can degrade other PFRs by utilizing them as sole phosphorus source. Both strains grew on tris(2,3-dibromopropyl) phosphate, tricresyl and triphenyl phosphates. Stain TDK1 did not grow on all trialkyl phosphates tested, whereas strain TCM1 grew moderately on tributyl phosphate and slightly on tris(2-butoxyethyl) phosphate, triethyl phosphate and trimethyl phosphate. These results demonstrate that the strains can degrade not only TDCPP and TCEP but also other PFRs, and that the strains have different substrate specificity for trialkyl phosphates.
Figure 6.
Degradation of TDCPP and TCEP by strains TCM1 (A) and TDK1 (B) and generation of 2-CE and 1,3-DCP (C and D). The cultivations were performed aerobically at 30°C in a minimal medium containing 20 μM of TCEP or TDCPP as the sole phosphorus source. (A and B) Open circles and triangles represent the concentrations of TCEP and TDCPP, respectively, and their filled forms represent concentrations for autoclaved control cells. (C and D) Open circles and triangles represent the concentrations of 2-CE and 1,3-DCP, respectively. Each data point represents the mean of at least two independent determinations.
2.2. Microbial detoxification of TDCPP and TCEP by two bacterial strains
We have successfully isolated TCEP- and TDCPP-degrading bacteria. However, neither strain can degrade the resulting toxic and persistent metabolites 2-CE and 1,3-DCP. Elimination of the metabolites is required before the strains can be used to degrade TDCPP and TCEP in practice. Fortunately, bacteria with chloroalcohol-degrading ability have been well-documented. We thus attempted to completely detoxify the PFRs by combining strain TCM1 with bacteria capable of degrading the chloroalcohols (Takahashi et al., 2012a; Takahashi, et al., 2012b).
2.2.1. Microbial detoxification of TDCPP using Sphingobium sp. strain TCM1 and Arthrobacter sp. strain PY1
Several 1,3-DCP-degrading bacteria have been reported, including Arthrobacter sp. strains PY1 (Yonetani et al., 2004) and AD2 (van den Wijngaard et al., 1991), A. erithii H10a (Assis et al., 1998), Agrobacterium radiobacter strain AD1 (van den Wijngaard et al., 1989), and Corynebacterium sp. strain N-1074 (Nakamura et al., 1991). Of these, Arthrobacter sp. strain PY1 exhibits high 1,3-DCP degradation ability. Therefore, we attempted to detoxify TDCPP by co-habitation of strain TCM1 and Arthrobacter sp. PY1 in a resting cell reaction (Fig. 7) (Takahashi et al., 2012a).
Figure 7.
Complete detoxification of TDCPP by Sphingobium sp. strain TCM1 and 1,3-DCP-degrading bacterium Arthrobacter sp. strain PY1.
2.2.1.1. Freezing and lyophilization of strains TCM1 and PY1 cells
For resting cell preparation, we first examined the effect of freezing and lyophilization on the activity of strains TCM1 and PY1. The TDCPP-hydrolyzing activity of strain TCM1 intact cells was 1.07 μmol h-1 OD660-1, whereas respective activities of frozen and lyophilized cells were 0.90 and 0.84 μmol h-1 OD660-1. On the other hand, the 1,3-DCP-dehalogenating activity of strain PY1 intact cells was 0.22 μmol h-1 OD660-1, with respective frozen and lyophilized cell activities of 0.23 and 0.26 μmol h-1 OD660-1. These results reveal that freezing and lyophilization treatments cause no significant decline in degradation activities of the strains.
2.2.1.2. Optimum TDCPP and 1,3-DCP degradation conditions of strains TCM1 and PY1
We then determined the optimum temperature and pH for lyophilized cell activity (Fig. 8). At pH 9.0 for strain TCM1 and pH 8.5 for strain PY1, the highest activity of TCM1 and PY1 cells occurred at 30°C (2.53 μmol h-1 OD660-1) and 35°C (1.31 μmol h-1 OD660-1), respectively (Fig. 8A). At 30°C, the highest activity of TCM1 and PY1 cells occurred at pH 8.5 (2.48 μmol h-1 OD660-1) and pH 9.5 with 50 mM Tris-H2SO4 (0.95 μmol h-1 OD660-1), respectively (Fig. 8B). We thus established the optimum temperature as 30°C and 35°C and the optimum pH as 8.5 and 9.5 for strains TCM1 and PY1, respectively.
Figure 8.
Effect of temperature and pH on the degradation activity of strains TCM1 and PY1. (A) effect of temperature: TDCPP hydrolyzation activity of strain TCM1 cells (closed circle) and 1,3-DCP dehalogenation activity of strain PY1 cells (open circle) were, respectively, assayed in 50 mM Tris-H2SO4 buffer (pH 9.0) and 50 mM Tris-H2SO4 buffer (pH 8.5). (B) effect of pH: TDCPP hydrolyzation activity of strain TCM1 cells (closed symbols) and 1,3-DCP dehalogenation activity of strain PY1 cells (open symbols) was assayed at 30°C in 50 mM MOPS-NaOH buffer (circle, pH 6.0-7.5), Tris-H2SO4 buffer (triangle, pH 7.5-9.5), and glycine-NaOH buffer (square, pH 9.0-12.0). Each datum represents means of two independent determinations.
2.2.1.3. Complete detoxification of TDCPP by mixed bacteria cells
Based on the optimum conditions, we set the reaction temperature to 30°C and pH to 9.0 (50 mM Tris-H2SO4) for TDCPP detoxification by mixed bacteria (Fig. 9). Under these conditions, the respective activities of strains TCM1 and PY1 were 2.21 and 0.92 μmol h-1 OD660-1. In the detoxification reaction using a mixture of TCM1 and PY1 cells (OD660 0.05 and 0.2, respectively), approximately 50 μM of TDCPP disappeared within 1 h, and 1,3-DCP and chloride ions were formed to levels of approximately 100 and 120 μM, respectively, after 2 h (Fig. 9A). This result suggests incomplete detoxification of TDCPP due to low 1,3-DCP dehalogenation activity. Increasing the strain PY1 population to an OD660 of 4.0 decreased the TDCPP hydrolyzation rate of TCM1 cells, but completely eliminated the resulting 1,3-DCP after 10 h (Fig. 9B). At the same time, chloride ion concentration had reached its theoretical value expected from the initial TDCPP concentration, demonstrating that complete detoxification of TDCPP is achievable using strains TCM1 and PY1.
Figure 9.
Complete detoxification of TDCPP by the mixed resting cells of strains TCM1 and PY1. The reactions were performed at 30°C with 50 μM TDCPP in 50 mM Tris - H2SO4 buffer (pH 9.0), and TDCPP (circles), 1,3-DCP (triangles) and chloride ion (squares) were determined. Cell concentrations of strains TCM1 and PY1 for each reaction were, respectively, OD660 of 0.05 and 0.2 (A) and 0.04 and 4.0 (B). Each datum represents means of two independent determinations.
2.2.2. Microbial detoxification of TCEP using Sphingobium sp. strain TCM1 and Xanthobacter autotrophicus strain GJ10
Several 2-CE-degrading bacteria have been reported, including Xanthobacter autotrophicus strain GJ10 (Janssen et al., 1985), Pseudomonas putida strain US2 (Strotmann et al., 1990) and P. atutzeri strain JJ (Dijk et al., 2003). Among these, the degradation of 2-CE by X. autotrophicus strain GJ10 has been well characterized. Therefore, we attempted to detoxify TCEP by co-habitation of strain TCM1 and X. autotrophicus strain GJ10 (Fig. 10) (Takahashi et al., 2012b).
Figure 10.
Complete detoxification of TCEP by Sphingobium sp. strain TCM1 and 2-CE-degrading bacterium Xanthobacter autotrophicus strain GJ10.
2.2.2.1. Optimum TCEP degradation condition of strain TCM1
We first determined the optimum temperature and pH for TCEP degradation by strain TCM1 in a resting reaction using lyophilized cells. At pH 7.4, the highest activity was obtained at 30°C (14.1 nmol min-1 OD660-1). Maintaining this temperature and varying the pH, the highest activity was recorded at pH 8.5 (14.6 nmol min-1 OD660-1). These optimum conditions were identical to those for TDCPP, suggesting that the same enzyme(s) might be involved in the degradation of both compounds.
Under the optimum conditions, TCM1 cells completely eliminated 10, 20 and 50 μM of TCEP within 3 h, but the generated 2-CE was approximately 50% of its theoretical value based on the initial TCEP concentrations (Fig. 11). Phosphotriesterase that can hydrolyze organophosphorus pesticides structurally similar to TCEP, such as chlorpyrifos, require two zinc ions for catalysis, and enzyme activity can be maximized by replacing Zn2+ with Co2+ (Omburo et al., 1992). A bacterial phosphodiesterase that can hydrolyze alkyl phosphodiesters similarly requires divalent metals (Gerlt & Wan, 1979). We therefore examined the effect of Co2+ as well as cell amount on TCEP hydrolysis (Fig. 11). In the reaction using approximately 10 μM of TCEP without Co2+, 2-CE reached 21.2 μM (OD660 of 0.8) after 3 h. Addition of 50 μM Co2+ resulted in an increase of 2-CE to 32.3 μM, equivalent to the theoretical value of 30 μM (Fig. 11B). These results showed that complete hydrolysis can be achieved at an OD660 of 0.8 with 50 μM of Co2+.
Figure 11.
Effect of Co2+ and cell amount on TCEP hydrolysis by strain TCM1-resting cells. The reactions were performed at 30°C using the resting cells at OD660 of 0.4 (circles) or 0.8 (triangles) with (open symbols) or without (closed symbols) 50 μM Co2+ in 50 mM Tris-H2SO4 buffer (pH 8.5) containing 10 μM TCEP, and TCEP (A) and 2-CE (B) were determined. Each datum represents the mean of two independent determinations. The inconsistency of the initial concentrations of TCEP at zero time with the set-up ones was mainly attributed to reaction progress in several minutes to stop the reaction.
2.2.2.2. Optimum 2-CE degradation condition of strain GJ10
We prepared resting cells of intact, frozen and lyophilized cells of X. autotrophicus strain GJ10 and examined their 2-CE degradation activity. Activity was detected only in frozen cells at 4.93 pmol min-1 OD450-1, four orders lower than the TCEP degradation activity of strain TCM1. This low 2-CE degradation activity might be attributable to the lack of coenzyme regeneration of enzymes involved in the degradation process. We next examined 2-CE degradation in a growing cell reaction. The growing cells completely degraded approximately 180 μM of 2-CE within 24 h. The degradation ability was estimated to be a minimum of 7.5 μM h-1, comparable to the TCEP degradation ability of strain TCM1-resting cells (approximately 10 μM h-1). This result shows that growing cells of strain GJ10 can degrade 2-CE effectively.
2.2.2.3. Complete detoxification of TCEP by two bacterial strains
Based on the results described above, we examined whether combining TCEP hydrolysis by TCM1 resting cells and 2-CE degradation by GJ10 growing cells would completely detoxify TCEP (Fig. 12). TCM1 resting cells abolished 9.6 μM of TCEP within 4 h, releasing 2-CE at 29.0 μM, equivalent to that estimated from the initial TCEP concentration, and consistent with complete TCEP hydrolysis (Fig. 12A and B). The generated 2-CE was abolished by GJ10 growing cells within 48 h, and chloride ion concentration reached 30.2 μM after 144 h, equivalent to that estimated from the generated 2-CE (Fig. 12C and D). Taken together, these results demonstrate that complete detoxification of TDCPP can be achieved using strains TCM1 and GJ10.
We have successfully isolated two novel bacterial strains capable of degrading the persistent and potential toxic PFRs, TCEP and TDCPP, which have become worldwide environmental contaminants. The two strains TCM1 and TDK1 belong to Sphingobium sp. and Sphingomonas sp. respectively. The strains are the first microorganisms reported to degrade the persistent PFRs. They degrade the compounds by hydrolyzing their phosphotriester bonds to produce metabolites 1,3-DCP from TDCPP and 2-CE from TCEP, which are themselves toxic and non-self-biodegradable. In a successful attempt to completely detoxify the FPRs, we combined TCM1 with the 1.3-DCP-degrading bacterium Arthrobacter sp. strain PY1 (for TDCPP degradation), and with the 2-CE-degrading bacterium X. autotrophicus strain GJ10 (for TCEP degradation). This is the first description of microbial FPR detoxification. The bacteria and the microbial detoxification techniques may prove useful for the bioremediation of sites contaminated with intractable compounds. Further studies on the PFRs-degrading bacteria as well as the chloroalcohols-degrading bacteria, and on the detoxification techniques, could help to establish more efficient detoxifications, and could also provide novel insights into microbial degradation of organophosphorus compounds. We are now working towards elucidating the enzymes and the genes involved in the degradation processes.
Figure 12.
Complete detoxification of TCEP by Sphingobium sp. strain TCM1-resting cell reaction (A and B) and the following X. autotrophicus GJ10-growing cell reaction (C and D). The resting cell reaction was performed at 30°C with (+) or without (-) strain TCM1 cells at OD660 of 0.8 in 50 mM Tris-H2SO4 buffer (pH 8.5) containing 10 μM TCEP and 50 μM Co2+, and TCEP (A) and 2-CE (B) were determined. The growing cell reaction was performed at 30°C with (closed symbols) or without (open symbols) strain GJ10 cells in a medium containing the generated 2-CE as the sole carbon source, and 2-CE (C) and chloride ion (D) was determined. ND means not detected. Each datum represents the mean of two independent determinations.
This research was supported in part by a Grant-in-Aid for Scientific Research (B) (to Y. K) from the Ministry of Education, Science, Sports, and Culture of Japan, by the River Environment Fund (REF) in charge of the Foundation of River and Watershed Environment Management (FOREM) (to Y. K.), by a grant from the Uchida Energy Science Promotion Foundation (to S. T.) and by the Kurita Water and Environment Foundation (to S. T).
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Written By
Shouji Takahashi, Katsumasa Abe and Yoshio Kera
Submitted: September 12th, 2012Published: February 7th, 2013
Open access
