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Long-term toxicity to fish

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Description of key information

No long-term toxicity expected.

Key value for chemical safety assessment

Additional information

There are no long-term fish studies available for Fatty acids, soya, 2-ethylhexyl esters (CAS 93572-14-6). However, read across data on short-term toxicity are available for all three trophic levels, fish, daphnia and algae, indicating a low potential for aquatic toxicity. Also NOECs obtained from read across studies on algal growth and daphnia reproduction are clearly above 1 mg/L (nominal), i.e. the limit of water solubility.

Additionally, the aquatic concentrations of the substance is expected to be very low. Since the substance is considered to be readily biodegradable and has a high adsorption potential (log Koc 6.47, MCI method, KOCWIN v2.00), it is expected to be eliminated in sewage treatment plants to a high extent. In the aquatic environment, the concentration in the water phase will be reduced by biodegradation and adsorption to solid particles and to sediment.

Food ingestion is likely to be the main uptake route, since the substance may adsorb to solid particles, which could be potentially ingested by fish. Also for sediment-dwelling organisms the main uptake route will be ingestion of contaminated sediment. In the case of ingestion, the substance is predicted to undergo metabolism. Esters of primary alcohols, containing from 1 to 18 carbon atoms, with fatty acids, containing from 2 to 18 carbon atoms, have been shown to be hydrolysed by pancreatic lipases in a study by Mattson and Volpenhein (Mattson and Volpenhein, 1972). Measured rates of enzyme catalysed hydrolysis varied between 2 and 5 µeq/min/mg enzyme for the different chain lengths (IUCLID section 7.1.1, Mattson and Volpenhein, 1972; and references therein). Only moderate differences in the rate of hydrolysis were observed for different long chain saturated and unsaturated fatty-acid esters, in studies investigating the fatty acid specificity of pancreatic lipases (Macrae and Hammond, 1985; and references therein). The resulting free fatty acids and alcohols are absorbed from the intestine into the blood stream. The alcohols are metabolised primarily in the liver through a series of oxidative steps, finally yielding carbon dioxide (Berg, 2001; HSDB).
Fatty acids are either metabolised via the beta-oxidation pathway in order to generate energy for the cell or reconstituted into glyceride esters and stored in the fat depots in the body (Berg et al., 2001). For fatty acids up to C22, beta-oxidation generally takes place in the mitochondria, resulting in the final product acetyl-CoA, which directly enters the citric acids cycle (Berg, 2002). Beta-oxidation of longer fatty acids takes place in the peroxisomes and is incomplete (Reddy and Hashimoto, 2001; Singh et al., 1987; Le Borgne and Demarquoy, 2012; and references therein). It gives rise to medium chain acyl-CoA, which are then taken in charge by the carnitine octanoyl transferase and converted into acyl-carnitine that can leave the peroxisome and, at least for some of them, may be fully oxidized in the mitochondria (Le Borgne and Demarquoy, 2012; and references therein). Peroxisomalβ-oxidation has also been shown to take place in fish, mussels and algae (Rocha et al., 2003; and references therein; Frøyland et al., 2000; Bilbao et al., 2009; Winkler et al., 1988). Metabolic pathways in fish are generally similar to those in mammals. Lipids and their constituents, fatty acids, are in particularly a major organic constituent of fish and play major roles as sources of metabolic energy (Tocher, 2003).

In conclusion, the substance will be mainly taken up by ingestion and digested through common metabolic pathways, providing a valuable energy source for the organism, as dietary fats. Long-term toxic effects on fish are therefore not to be expected.

Based on this information and for reasons of animal welfare, long-term testing on fish is not proposed.



Berg, J.M., Tymoczko, J.L. and Stryer, L., 2002, Biochemistry, 5th edition, W.H. Freeman and Company

Bilbao, E., Cajaraville, M.P., Cancio, I. (2009), Cloning and expression pattern of peroxisomal β-oxidation genes palmitoyl-CoA oxidase, multifunctional protein and 3-ketoacyl-CoA thiolase in mussel Mytilus galloprovincialis and thicklip grey mullet Chelon labrosus, Gene, 443(1-2): 132-42

Le Borgne, F., Demarquoy, J. (2012): Interaction between peroxisomes and mitochondria in fatty acid metabolism, Open Journal of Molecular and Integrative Physiology, 2012, 2, 27-33

Frøyland, Lie, Berge (2000), Mitochondrial and peroxisomal β-oxidation capacities in various tissues from Atlantic salmon Salmo salar, Aquaculture Nutrition, 6 (2): 85-89

HSDB – Hazardous Substances Data Bank, Toxnet Home, National Library of Medicinehttp: //toxnet. nlm. nih. gov/cgi-bin/sis/htmlgen?HSDB

Macrae, A.R., Hammond, R.C. (1985) Present and future applications of lipases, Biotechnology and Genetic Engineering Reviews, 3: 193-217

Mattson, F.H. and Volpenheim, R.A. (1972): Relative rates of hydrolysis by rat pancreatic lipase of esters of C2-C18 fatty acids with C1-C18 primary n-alcohols, Journal of Lipid Research, 10, 1969

Reddy and Hashimoto (2001) Peroxisomal beta-oxidation and peroxisome proliferator-activated receptor alpha: An adaptive metabolic System, Annual Review of Nutrition, 21, 193-230

Rocha, M.J., Rocha, E., Resende, A.D., Lobo-da-Cunha (2003) Measurement of peroxisomal enzyme activities in the liver of brown trout (Salmo trutta), using spectrophotometric methods, BMC Biochemistry, 4:2, doi:10.1186/1471-2091-4-2

Singh, H., Derwas, N. and Puolos, A. (1987) Beta-oxidation of very-long-chain fatty acids and their coenzyme A derivatives by human skin fibroblasts, Arch Biochem Biophys, 254(2): 526-33

Winkler, U., Säftel, W., Stabenau, H. (1988), beta-Oxidation of fatty acids in algae: Localization of thiolase and acyl-CoA oxidizing enzymes in three different organisms, Planta, 175(1): 91-98