Biodegradation DOI 10.1007/s10532-014-9685-2
ORIGINAL ARTICLE
Microorganisms hydrolyse amide bonds; knowledge enabling read-across of biodegradability of fatty acid amides Roy Geerts • Patrick Kuijer • Cornelis G. van Ginkel • Caroline M. Plugge
Received: 2 October 2013 / Accepted: 30 January 2014 Ó Springer Science+Business Media Dordrecht 2014
Abstract To get insight in the biodegradation and potential read-across of fatty acid amides, N-[3(dimethylamino)propyl] cocoamide and N-(1-ethylpiperazine) tall oil amide were used as model compounds. Two bacteria, Pseudomonas aeruginosa PK1 and Pseudomonas putida PK2 were isolated with N-[3-(dimethylamino)propyl] cocoamide and its hydrolysis product N,N-dimethyl-1,3-propanediamine, respectively. In mixed culture, both strains accomplished complete mineralization of N-[3(dimethylamino)propyl] cocoamide. Aeromonas hydrophila PK3 was enriched with N-(1-ethylpiperazine) tall oil amide and subsequently isolated using agar plates containing dodecanoate. N-(2-Aminoethyl)piperazine, the hydrolysis product of N-(1ethylpiperazine) tall oil amide, was not degraded. The aerobic biodegradation pathway for primary and secondary fatty acid amides of P. aeruginosa and A. hydrophila involved initial hydrolysis of the amide bond producing ammonium, or amines, where the fatty acids formed were immediately metabolized. Complete mineralization of secondary fatty acid amides depended on the biodegradability of the R. Geerts (&) P. Kuijer C. G. van Ginkel AkzoNobel N.V., P.O. Box 9300, 6800 SB Arnhem, The Netherlands e-mail:
[email protected] C. M. Plugge Laboratory of Microbiology, Wageningen University, P.O. Box 8033, 6700 EJ Wageningen, The Netherlands
released amine. Tertiary fatty acid amides were not transformed by P. aeruginosa or A. hydrophila. These strains were able to utilize all tested primary and secondary fatty acid amides independent of the amine structure and fatty acid. Read-across of previous reported ready biodegradability results of primary and secondary fatty acid amides is justified based on the broad substrate specificity and the initial hydrolytic attack of the two isolates PK1 and PK3. Keywords Fatty acid amides Hydrolysis Biodegradation pathway Substrate specificity Read-across
Introduction Surfactants are amphiphilic molecules, consisting of a hydrophilic (water-friendly) and a hydrophobic (water-repellent) domain, typically a long-chain hydrocarbon. Fatty acid amides are surfactants produced by reacting a fatty acid with ammonium, an amine or polyamine (Johansson 2003). The amide bond of these surfactants can be primary, secondary or tertiary (Fig. 1). Fatty acid amides are used in asphalt applications, as corrosion inhibitors, fabric softener and in personal care products like hair conditioners (Hauthal 2009; AkzoNobel 2010). Use in industrial and domestic products leads to discharge of these surfactants into wastewater treatment systems and the aquatic environment. Release of these surfactants into
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Biodegradation
Fig. 1 General structure of primary, secondary and tertiary fatty acid amide; in which R is a saturated or unsaturated alkyl chain derived from a fatty acid; R1 and R2 may be an alkyl, aryl, alkylene oxide, amine or polyamine group
the environment warrants assessment of their biodegradability. Amidases are widely spread in nature, present in archaea, eukarya and bacteria having various biochemical functions. These amide bond-cleaving enzymes are able to catalyze four different types of reactions namely amidotransferase, acyltransferase, acid transferase and ester transferase. Using water, hydroxylamine or hydrazine as deacylating cosubstrate amidases are able to transform diverse amides, acids, esters or nitriles in their corresponding carboxylic acids, hydroxamic acids or acid hydrazides (Fournand and Arnaud 2001; Sharma et al. 2009). The initial step in the biodegradation of primary amides, such as acetamide, propionamide, butyramide, valeramide, stearamide, acrylamide, benzamide and nicotinamide is a hydrolytic reaction by an amidase releasing the fatty acid and ammonium (Clarke 1980; Nawaz et al. 1992; Nawaz et al. 1994; Brown et al. 1969; Kagayama and Ohe 1990; Sonke and Kaptein 2012). Yamane et al. (2008) proposed a biodegradation pathway for N-[3-(dimethylamino)propyl]docosanamide, a secondary fatty acid amide. N-[3-(Dimethylamino)propyl]docosanamide is probably cleaved hydrolytically at the amide bond by a mixed culture derived from river water resulting in a transient accumulation of N,N-dimethyl-1,3-propanediamine. Hydrolytic activity is also present in a Pseudomonas sp initiating the biodegradation of Nmethyltaurine oleoyl amide, a tertiary amide (Denger et al. 2008). The aim of REACH (Registration, Evaluation, Authorization of Chemicals), a European legislation is to ensure a high level of protection of human health and the environment. To this end, REACH advocates the use of read-across as a means to satisfy the information requirements for environmental risk
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assessments. For read-across, the underlying hypothesis has to be that compounds within a ‘‘family’’ should have similar activities. The challenge in applying read-across of ready biodegradability tests results (OECD 1992) lies in deriving a way of determining the ‘‘family’’. While guidance is available to read-across toxicology data, there still remains little practical guidance on how read-across should be applied in evaluation of biodegradability. The only rational way to determine ‘‘families’’ is probably based on the biodegradation pathway and substrate specificities (van Ginkel 2007). The objective of this study was to describe the microbial degradation of fatty acid amides by mixed and pure cultures with particular focus on the biodegradation pathway and substrate specificities to enable read-across.
Materials and methods Chemicals N-[3-(Dimethylamino)propyl] cocoamide, N-[3(dimethylamino)propyl] hydrogenated tallow amide, N[3-(dimethylamino)propyl] rapeseed amide, N-[3-(bis(2hydroxyethyl)amino)propyl] cocoamide and N-(1-ethylpiperazine) tall oil amide were provided by AkzoNobel Surface Chemistry AB. SynperonicÒ PE/P105 was obtained from Croda. N-Methyl-N-ethanoate dodecanamide, N,N-dibutyldodecanamide, N,N-dimethyldodecanamide, N,N-dimethyl-1,3-propanediamine, oleate, dodecanoate, citrate, N-(2-aminoethyl)piperazine, hexanoamide, dodecanamide, oleamide, acetate and stearate were obtained from Sigma Aldrich. All other chemicals used were of reagent grade. Deionized water containing no more than 0.01 mg L-1 copper was prepared in a water purification system. Inoculum The activated sludge used as inoculum for the closed bottle tests and the enrichment cultures was collected from an aeration tank of a municipal wastewater treatment plant located in Duiven, the Netherlands. This plant consists of mechanical and biological stages for the treatment of predominantly domestic wastewater.
Biodegradation
Closed Bottle test The ready biodegradability test method described under no. 301D in the OECD Guidelines for Testing of Chemicals (OECD 1992) was used, and performed with two minor deviations. Activated sludge at a concentration of 2 mg dry weight L-1 was used as inoculum instead of an effluent/extract mixture and ammonium chloride was omitted from the medium to prevent oxygen consumption due to nitrification (van Ginkel and Stroo 1992).
Enrichment, isolation and characterization Selective enrichments, using N-[3-(dimethylamino)propyl] cocoamide or N-(1-ethylpiperazine) tall oil amide as sole source of carbon and energy, were performed at 30 °C in a 1.5 L fermentor with a working volume of 1 L (Applikon, Schiedam, the Netherlands). The impeller speed was 400 rpm and the pH was maintained at 6.8 with a solution of 50 g L-1 Na2HPO4, using a pH-electrode connected to a pH controller (ADI 1020, Applikon, Schiedam, the Netherlands). Mineral salts medium amended with N-[3-(dimethylamino)propyl] cocoamide or N(1-ethylpiperazine) tall oil amide was supplied with a dilution rate of 10 day-1 to the fermentor using a peristaltic pump (Meyvis & Co., Bergen op Zoom, the Netherlands). After 1 week of operation the dilution rate of the enrichment on N-[3-(dimethylamino)propyl] cocoamide was stepwise increased during a 30 days period to 1 day-1. Repeated batch subculturing with N-[3-(dimethylamino)propyl] cocoamide and N-(1-ethylpiperazine) tall oil amide was started after 40 days, using the fermentor enrichment cultures as inoculum. Microorganisms enriched on N-[3-(dimethylamino)propyl] cocoamide were also used as inoculum for repeated batch subculturing on N,N-dimethyl-1,3-propanediamine. After growth was obtained dilutions of the batch subcultures were streaked on agar plates and subsequently one colony was streaked repeatedly to purity. Analyses to identify all three strains were carried out by DSMZ (Braunschweig, Germany) using fatty acid methyl ester analysis, substrate profiling and 16S rRNA sequencing.
Growth medium, cultivation, and growth conditions The mineral salts medium used for isolation and growth of the bacteria contained per litre 1.55 g K2HPO4, 0.85 g NaH2PO4, 0.5 g NH4Cl, 0.1 g MgSO4.7H2O, 0.01 g Na2H2EDTA.H2O, 0.01 g FeSO4.7H2O and 0.1 mL of a trace solution described by Vishniac and Santer (1957). Agar plates used for isolation were incubated at 30 °C and contained beside the mineral salts medium 15 g L-1 bacteriological agar and 1 g L-1 of the substrates: N-[3-(dimethylamino)propyl] cocoamide, N,N-dimethyl-1,3-propanediamine or dodecanoate. For the isolation of a microorganism degrading N-(1ethylpiperazine) tall oil amide agar plates with 1 g L-1 dodecanoate were used because it was not possible to make agar plates containing N-(1-ethylpiperazine) tall oil amide. All three isolated strains were grown and maintained in batch cultures using 1 L flasks with 200 mL of the mineral salts medium spiked with 1 g L-1N-[3(dimethylamino)propyl] cocoamide, 1 g L-1N,Ndimethyl-1,3-propanediamine or 1 g L-1N-(1-ethylpiperazine) tall oil amide, respectively. Growth experiments with the isolates were performed in 100 mL flasks using 20 mL of mineral salts medium supplemented with different substrates (1 g L-1). In the batch cultures containing N-[3-(dimethylamino)propyl] cocoamide, N-(1-ethylpiperazine) tall oil amide and several of the other growth substrates the initial substrate concentration was toxic to the isolates. To reduce the substrate concentration in the water phase and herewith the toxicity, 16 g L-1 silica gel was added (van Ginkel et al. 1992). Growth was observed by measuring the increase in turbidity over time with a Hach Ratio XR turbidimeter (Hach Lange GmbH, Du¨sseldorf, Germany). Moreover, cell number increase was routineously determined using phase contrast microscopy. All batch cultures were shaken at 100 rpm in an orbital incubator at 30 °C. Continuous growth experiments with the isolates were performed in the fermentor systems used for the selective enrichments. At steady state conditions the mineral salts medium amended with 1 g L-1N-[3(dimethylamino)propyl] cocoamide or 1 g L-1N-(1ethylpiperazine) tall oil amide was dosed at a dilution rate of 1 day-1, the temperature was 30 °C and the pH was 6.8.
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Biodegradation
Washed cell suspensions and cell-free extracts Cell suspensions of the continuous cultures were harvested by centrifugation at 10,000 g for 10 min at 4 °C, washed three times with 15 mM phosphate buffer, pH 7.0 or for the preparation of cell-free extract with 50 mM Tris/HCl buffer, pH 9.0. The washed cell suspensions were stored at 4 °C. Cell breakage of washed cell suspensions was achieved by sonication. Debris and whole cells were removed by centrifugation (30,000 g for 10 min at 4 °C) and the supernatant containing the cell free extract was used immediately in the enzyme assay. For the preparation of cell free extracts of acetate and dodecanamide grown cells, cells were harvested from a 2 L batch culture at the end of the exponential growth phase. Oxidation rate by washed cell suspensions Oxygen uptake was measured with a Biological Oxygen Monitor (Yellow Springs Instruments, Yellow Springs, Ohio), which consisted of an electrode and a water-jacketed vessel (5 mL). Washed cell suspensions were incubated in the vessel at 30 °C for at least five minutes to allow determination of the endogenous respiration rate. Subsequently, 0.1 mL of a 1 g L-1 substrate solution or suspension was injected, and the substrate-dependent respiration rate was determined. As dodecanamide has a very low water solubility, the water solubility was increased by preparing a suspension of 1 g L-1 dodecanamide with ethylene oxide/propylene oxide block copolymer, synperonicÒ PE/P105 (1:1 w/w). Enzyme assay The amidase reaction was assayed (modified from Denger et al. 2008) discontinuously at 30 °C in 50 mM Tris/HCl buffer, pH 9.0, including 2.1 mM N-[3-(dimethylamino)propyl] cocoamide or 2.1 mM dodecanamide. The reaction was started with the addition of crude cell extract (34 and 239 lg protein mL-1 for cells incubated with N-[3-(dimethylamino)propyl] cocoamide and grown on N-[3(dimethylamino)propyl] cocoamide and acetate, respectively; 89 lg protein mL-1 was used for cells grown on N-[3-(dimethylamino)propyl] cocoamide and incubated with dodecanamide) and stopped again by addition of 0.1 mL of 3 M HCl. Samples (1 mL)
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were taken at intervals and the formation of N,Ndimethyl-1,3-propanediamine or ammonium was measured by cat-ion chromatography.
Analytical methods Before the determination of the non-purgeable organic carbon and total nitrogen content, the effluents were filtered using cellulose nitrate filters with pores of 0.45 lm to remove sludge particles. Filtered samples were acidified prior to injection in a TOC apparatus (Shimadzu Corporation, ‘s-Hertogenbosch, The Netherlands). N,N-Dimethyl-1,3-propanediamine, ammonium and N-(2-aminoethyl)piperazine were determined by cat-ion chromatography (Dionex-120), using (suppressed) conductivity detection (Dionex, Amsterdam, The Netherlands). N,N-Dimethyl-1,3-propanediamine and ammonium were analyzed using an CG14 (guard) and CS14 (analytical) column. The eluent used for the analysis of N,N-dimethyl-1,3-propanediamine consisted of 25 mM methylsulphonic acid and 10 % acetonitrile, and was diluted two times with deionized water when ammonium was analyzed. N-(2-Aminoethyl)piperazine was determined by using only the CG14 (guard) column and an eluent consisting of 35 mM methylsulphonic acid and 8.8 % acetonitrile. The chromatographic systems consisted of a CMMS300 4 mm suppressor operated in an external chemical mode using a pump for dispensing the regenerant (100 mM tetrabutylammonium hydroxide). Conductivity was detected with a CDM-3 flow through conductivity cell and a DS4 detection stabilizer. The eluent flow rate was 1.0 mL min-1, the sample loop was 25 lL, and analyses were performed at ambient temperature. Dissolved oxygen concentrations were determined electrochemically using an oxygen electrode (WTW Trioxmatic EO 200) and meter (WTW OXI 530) (Retsch, Ochten, The Netherlands). The pH was measured using a Knick 765 calimatic pH meter (Elektronische Messgerate GmbH, Berlin, Germany) and the temperature was measured and recorded with a thermo couple connected to a data logger. Protein concentrations of the cell extracts were determined using the bicinchoninic acid assay (SigmaAldrich) which uses bovine serum albumin as protein standard.
Biodegradation
Fig. 2 Biodegradability of N-[3-(dimethylamino)propyl] cocoamide(unfilled square), N,N-dimethyl-1,3-propanediamine (unfilled circle), N-(1-ethylpiperazine) tall oil amide (filled square) and N-(2-aminoethyl)piperazine (filled circle) in Closed Bottle tests (OECD 301D)
Results The Closed Bottle test (OECD 1992) is a screening test designed to assess rapid and complete biodegradation of chemicals in the environment. The biodegradation of N-[3-(dimethylamino)propyl] cocoamide, N-(1ethylpiperazine) tall oil amide and their hydrolysis products, N,N-dimethyl-1,3-propanediamine and N-(2aminoethyl)piperazine was assessed in Closed Bottle tests inoculated with activated sludge (Fig. 2). N-[3(Dimethylamino)propyl] cocoamide reached 70 % biodegradation thereby fulfilling the pass criterion of C60 % within 28 days. N-[3-(Dimethylamino)propyl] cocoamide is therefore regarded as readily biodegradable. The ready biodegradability of N,N-dimethyl-1,3propanediamine underpins the ready biodegradability N-[3-(dimethylamino)propyl] cocoamide. N-(1-Ethylpiperazine) tall oil amide is degraded partially in the closed bottle test, as indicated by only 34 % biodegradation achieved (Fig. 2). Inability of microorganisms to degrade N-(2-aminoethyl)piperazine is most likely the cause of this incomplete degradation. Biodegradation of both N-[3-(dimethylamino)propyl] cocoamide and N-(1-ethylpiperazine) tall oil amide started without a lag-phase indicating fast hydrolysis of the amide bonds and subsequent degradation of the fatty acids formed (Fig. 2).
Strains PK1, PK2 and PK3, were isolated from activated sludge using continuous cultures and subsequent batch sub-culturing. Strain PK1 was enriched and isolated using N-[3-(dimethylamino)propyl] cocoamide as the sole source of carbon and energy. Strain PK2 was enriched in a fermentor fed with N-[3(dimethylamino)propyl] cocoamide followed by repeated batch sub-culturing using N,N-dimethyl1,3-propanediamine as growth substrate. Agar plates with N,N-dimethyl-1,3-propanediamine were used to isolate this strain. N-(1-Ethylpiperazine) tall oil amide was used as growth substrate for the enrichment of strain PK3, which was subsequently streaked to purity using agar plates containing dodecanoate. The profiles of the cellular fatty acids for PK1 and PK2 were both typical for the genus Pseudomonas. The results of the phenotypic tests confirmed these identifications. Partial sequences of the 16S rRNA of PK1 and PK2 showed 100 % similarity to Pseudomonas aeruginosa and Pseudomonas putida, respectively. The cellular fatty acid profile for strain PK3 resembled that of the species Aeromonas hydrophila. This identification was supported by 99.8 % similarity with A. hydrophila using the partial sequence of the 16S rRNA and the phenotypic tests. Growth of P. aeruginosa strain PK1 and A. hydrophila strain PK3 was determined in batch cultures containing different substrates. Both strains were able to grow on the amidoamines (secondary amides), N[3-(dimethylamino)propyl] cocoamide, N-[3-(dimethylamino)propyl] rapeseed amide, N-[3-(dimethylamino)propyl] hydrogenated tallow amide, N-(1ethylpiperazine) tall oil amide and N-[3-(bis(2hydroxyethyl)amino)propyl] cocoamide. P. aeruginosa strain PK1 and A. hydrophila strain PK3 also supported growth on primary fatty acid amides dodecanamide and oleamide. Growth was also observed on citrate, acetate, dodecanoate, oleate and stearate. Strain PK1 and PK3 were not able to utilize the tertiary fatty acid amides N-methyl-N-ethanoate-dodecanamide, N,N-dibutyldodecanamide and, N,N-dimethyldodecanamide as sole source of carbon and energy. N,N-Dimethyl-1,3-propanediamine and N-(2-aminoethyl)piperazine did not support growth of both PK1 and PK3. In batch cultures containing a nitrogen free mineral salts medium amended with 1 g L-1 of N-[3-(dimethylamino)propyl] cocoamide, growth of P. aeruginosa strain PK1 and P. putida strain PK2 was only
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Biodegradation Table 1 Oxygen uptake rates for washed cell suspensions of P. aeruginosa strain PK1 grown on N-[3-(dimethylamino)propyl] cocoamide at 30 °C Substrate
Oxygen consumption (nmol of O2 min-1 mg [dry wt] of cells-1)
N-[3-(Dimethylamino)propyl] cocoamidea
79
N-[3-(Dimethylamino)propyl] rapeseed amideb
54
N-[3-(Dimethylamino)propyl] hydrogenated tallow amidec
54
N-(1-Ethylpiperazine) tall oil amided
50
N-[3-(Bis(2-hydroxyethyl) amino)propyl] cocoamidea
27
N-methyl-N-ethanoatedodecanamidee
0
N,N-dibutyldodecanamidee
0
N,N-dimethyldodecanamidee Dodecanamidef
0 6
Hexanoamide
5
Acetate
17
Dodecanoate
65
Citrate
0
Oleate
50
N-(2-Aminoethyl)piperazine
0
N,N-Dimethyl-1,3propanediamine
0
Rates of oxygen uptake have been corrected for the endogenous oxygen uptake (9 nmol of O2 min-1 mg [dry wt] of cells-1) alkyl chain length distributions specified by Karleskind (1996) amainly C12 and C14 bmainly C18 and C22 cmainly C16 and C18 dmainly C18 econtains tertiary amide bond fSynperonicÒ PE/P105 used for increasing the water solubility of dodecanamide did not influence the oxygen uptake rate
observed when the medium was inoculated with both strains. Organic carbon and nitrogen removal was determined in continuous cultures with strain PK1 growing on N-[3-(dimethylamino)propyl] cocoamide and strain PK3 utilizing N-(1-ethylpiperazine) tall oil amide. Hydrolysis of the amide bonds, followed by biodegradation of the released fatty acids would result in organic carbon removal percentages of 71 and 75 for N-[3-(dimethylamino)propyl] cocoamide and N-(1-ethylpiperazine) tall oil amide, respectively.
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These removal percentages are close to the measured organic carbon removals of 67 % for N-[3-(dimethylamino)propyl] cocoamide and 71 % for N-(1-ethylpiperazine) tall oil amide. Organic nitrogen removal was negligible in the continuous cultures. This finding is in line with the observed accumulation of N,N-dimethyl1,3-propanediamine and N-(2-aminoethyl)piperazine during biodegradation of N-[3-(dimethylamino)propyl] cocoamide and N-(1-ethylpiperazine) tall oil amide, respectively. N,N-Dimethyl-1,3-propanediamine moiety of N-[3-(dimethylamino)propyl] cocoamide was almost completely recovered (94 %) from the effluent of the continuous culture. A recovery of 84 % of N-(2aminoethyl)piperazine from the effluent strongly indicates stoichiometric formation of this diamine during degradation of N-(1-ethylpiperazine) tall oil amide. Oxidation of different substrates was studied with washed cell suspensions of N-[3-(dimethylamino)propyl] cocoamide grown P. aeruginosa strain PK1 (Table 1). Increased oxygen uptake rates were found with N-[3-(dimethylamino)propyl] alkylamides with coco-, rapeseed-, and hydrogenated tallow-alkyl chains. Strain PK1 was also capable of oxidizing N-(1-ethylpiperazine) tall oil amide, N-[3-(bis(2-hydroxyethyl)amino)propyl] cocoamide, hexanoamide, dodecanoamide, acetate, dodecanoate and oleate. No increase in the oxygen uptake rate was observed with citrate, N,N-dimethyl-1,3-propanediamine, N-(2-aminoethyl)piperazine, and the tertiary amides N-methylN-ethanoate-dodecanamide, N,N-dibutyldodecanamide and N,N-dimethyldodecanamide. Amidase activity has been demonstrated in cellfree extracts P. aeruginosa strain PK1 grown on N-[3(dimethylamino)propyl] cocoamide. Incubation of cell-free extract with N-[3-(dimethylamino)propyl] cocoamide resulted in the formation of N,N-dimethyl1,3-propanediamine (Fig. 3). Ammonium was liberated by this cell-free extract when incubated with dodecanamide. The specific enzyme activities with N[3-(dimethylamino)propyl] cocoamide as substrate were 15.5 mkat (kg protein)-1 in extracts of N-[3(dimethylamino)propyl] cocoamide grown cells and 0.1 mkat (kg protein)-1 in extracts of acetate grown cells (1 kat = 1 mol s-1). The specific activity with dodecanamide as substrate was 0.6 mkat (kg protein)-1 in cell-free extracts of PK1 cells grown on N-[3(dimethylamino)propyl] cocoamide.
Biodegradation
Fig. 3 Formation of N,N-dimethyl-1,3-propanediamine from N-[3-(dimethylamino)propyl] cocoamide in extracts of N-[3(dimethylamino)propyl] cocoamide grown cells (filled circle) or acetate grown cells (unfilled triangle) of P. aeruginosa strain PK1
Fig. 4 Biodegradation pathways of N-(1-ethylpiperazine) tall oil amide and N-[3-(dimethylamino)propyl] cocoamide. Length alkyl chain R is C17 for N-(1-ethylpiperazine) tall oil amide and C11 for N-[3-(dimethylamino)propyl] cocoamide. The fatty acids formed are metabolized through the beta oxidation process. Complete degradation is accomplished by a consortium of two microorganisms
Discussion The hydrophilic and hydrophobic moieties of surfactants may be linked with a primary, secondary or tertiary amide bond. Preferences of the initial points of attack for amide containing compounds by activated sludge were found to be different (Helbling et al. 2010). Primary and secondary amides hydrolyze, whereas tertiary amides are N-dealkylated, hydroxylated or initially transformed by another unknown reaction. Hydrolysis of a tertiary amide bond, however, cannot be excluded. The biodegradation of Nmethyltaurine oleoyl amide, a surfactant with a tertiary amide was initially hydrolyzed by a Pseudomonas alcaligenes (Denger et al. 2008). N-Methyltaurine was excreted quantitatively during growth, while the fatty acid was used as sole source of carbon and energy. The pathway of complete mineralization of N-[3(dimethylamino)propyl] cocoamide (Fig. 4) is demonstrated upon found ability of P. aeruginosa strain PK1 to hydrolyze N-[3-(dimethylamino)propyl] cocoamide and to degrade fatty acids. Strain PK2 converts the hydrolysis product N,N-dimethyl-1,3-propanediamine. The pathway used by strain PK1 was previously proposed based on the transiently formed N,Ndimethyl-1,3-propanediamine during biodegradation
of N-[3-(dimethylamino)propyl]docosanamide in river water (Yamane et al. 2008). Hydrolase activity in cellfree extracts of P. aeruginosa strain PK1 has been demonstrated because cofactors were not required in the reaction mixture. Specific activities measured in cell-free extracts of acetate grown cells indicate a constitutive amidase activity for strain PK1. Hydrolysis of the amide bond in dodecanamide by cell-free extracts of strain PK1 resulted in the formation of a fatty acid and ammonium. The hydrolytic activity with dodecanamide was low compared to the activity with N-[3-(dimethylamino)propyl] cocoamide, most likely due to the low bioavailability of water insoluble dodecanamide. Most amidases currently known and described in literature were found in bacteria from which the majority shows optimal growth at neutral pH. Substrate specificity of amidases varies from broad spectrum to very specific and the enzymes are either induced or constitutively present (Sharma et al. 2009). A broad spectrum amidase was described by Nawaz et al. (1989) who measured growth and hydrolytic activity of P. putida on a wide range of substrates (acetamide, propionamide, butyramide, methacrylamide, isobutyramide and succinamide). Pseudomonas alcaligenes is able to grow on the N-oleoyl-N-
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Biodegradation
methyltaurine and possesses a substrate specific amidase which is responsible for the initial hydrolytic degradation step (Denger et al. 2008). Substrate specificity of the amidase determined by Kelly and Clarke (1962) indicated that the N-substituted amides could not be attacked by this enzyme. Later it was found that a single amino acid change in the enzyme allowed a small group substituted on the amide nitrogen as growth substrate and therefore a slight change in the conformation of the enzyme protein might be already enough to allow hydrolysis of amides with larger substituted groups on the nitrogen (Clarke 1980). Brevibacterium sp. R312 is able to hydrolyse primary amides and hydrolysis of the secondary amide N-methylacetamide is expected based on the found acyl transfer of N-methylacetamide to hydroxylamine (Thie´ry et al. 1986). Villarreal et al. (1994) tested the biodegradability of a series of N-phenyl acetamide analogs with alkyl substitution on the nitrogen atom. By selective enrichment four bacteria were isolated from soil that could grow on N-phenyl acetamide, containing a secondary amide. The four isolates showed induced enzyme activity with secondary amide analogs but not with tertiary amides. The fatty acid amide degrading isolates PK1 and PK3 have a broad substrate specificity, shown by growth of both microorganisms on primary and secondary fatty acid amides and the oxidation of these substrates by washed cell suspensions of strain PK1 (Table 1). Fatty acid amides containing a tertiary amide bond, such as N-methyl-N-ethanoate-dodecanamide, N,N-dibutyldodecanamide and N,N-dimethyldodecanamide did not support growth of the isolates. Fatty acids formed by the amidase activity are converted into fatty acyl-CoA followed by b-oxidation yielding a succession of acetyl-CoA units, which enter the central metabolism of microorganisms. The b-oxidation cycle also handles unsaturated fatty acids. Isomerization of the double bonds occurs to ensure the correct positioning of the double bonds in substrate for the b-oxidation sequence (Ratledge 1994). The biodegradation pathway of fatty acids demonstrates that a single microorganism can degrade both saturated and unsaturated fatty acid amides with different alkyl chain lengths. It is unlikely that the biodegradability of fatty acid amides differs significantly with varying alkyl chain lengths due to the broad substrate specificity of microorganisms degrading fatty acids (van Ginkel 2007).
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REACH allows assessment of biodegradability (non persistence) through read-across of ‘‘families’’ of chemicals. Definition of ‘‘families’’ for biodegradability purposes has already been based on biodegradation pathways (van Ginkel 2007). The ability of secondary fatty acid amides grown microorganisms to oxidize fatty acid amides is restricted to substances with primary and secondary amide bonds. Tertiary fatty acid amides should therefore not be included in the ‘‘family’’ used to read-across the biodegradability of primary and secondary fatty acid amides. Biodegradation of primary fatty acid amides immediately results in the formation of ammonium. A single microorganism with amidase activity and the ability to degrade fatty acids does result in complete degradation and possibly a ready result. Biodegradability of tallowamide, oleoylamide and N-[3-(N0 ,N0 -dimethyl-N0 -carboxymethyl)propyl] cocoamide was 73, 80 and 93 %, respectively, after 28 days in the Closed Bottle test demonstrating the ready biodegradability of these amides (EPA 2001). Biodegradation percentages in excess of 60, measured in other ready biodegradability tests (OECD 301B and 301C), demonstrated ready biodegradability of N-[3-(dimethylamino)propyl]docosanamide and N-[3(dimethylamino)propyl]octadecanamide, respectively (Yamane et al. 2008). Complete biodegradation of fatty acid amides (except the primary amides) requires at least two microorganisms. Strain PK1 only degraded the alkyl chain upon the hydrolysis of the amide bond. The released N,N-dimethyl-1,3-propanediamine was metabolized by strain PK2 liberating ammonium. The ammonium was used by both strains as nitrogen source when grown under nitrogen limiting conditions. A co-operation between two microorganisms enables a complete breakdown of these surfactants into carbon dioxide, water and inorganic salts (Fig. 4). Fatty acids are regarded as readily biodegradable (van Ginkel 2007). Ready biodegradability of N,Ndimethyl-1,3-propanediamine has been demonstrated (this study; ECB 2000a). These results are in line with the complete degradation of N-[3-(dimethylamino)propyl] cocoamide. N-(1-Ethylpiperazine) tall oil amide was degraded 34 % in the Closed Bottle tests indicating partial degradation (Fig. 2). The partial degradation of this surfactant can be attributed to the oxidation of the fatty acid formed through hydrolysis of the amide bond. The N-(2-aminoethyl)piperazine formed is however not biodegradable (this study; ECB
Biodegradation
2000b). Secondary fatty acid amides can only be classified readily biodegradable through read-across if the liberated amine is known to be readily biodegradable. The biodegradability of the amine or polyamine formed should therefore be included in a proper readacross.
Conclusions The aerobic biodegradation pathway for primary and secondary fatty acid amides involves an initial hydrolysis of the amide bond resulting in the formation of ammonium or (poly)amine and fatty acids. This hydrolytic reaction was found independent of the amine structure and fatty acid chain. The initial point of attack and the broad substrate specificity of microorganisms degrading these fatty acid amides, allows read-across of ready biodegradability test results. Ready biodegradability of a number of primary and secondary fatty acid amides has been demonstrated. Ready (ultimate) biodegradation of secondary fatty acid amides however depends on the biodegradability of the released amine. The inability of microorganisms to support growth of N-(2-aminoethyl)piperazine explains partial degradation of N-(1-ethylpiperazine) tall oil amide. Acknowledgments Financial support for the research described in this paper has been provided by AkzoNobel Surface Chemistry
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