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Environmental fate & pathways

Bioaccumulation: aquatic / sediment

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Description of key information

If aquatic exposure occurs, polyol esters category members will be mainly taken up by ingestion and digested through common metabolic pathways providing a valuable energy source for the organisms as dietary fats. The category members are not expected to bioaccumulate in aquatic or sediment organisms and secondary poisoning does not pose a risk.

Key value for chemical safety assessment

Additional information

Aquatic bioaccumulation

Experimental bioaccumulation data are not available for the members of the polyol ester category. The high log Kow as an intrinsic property of the category members indicates a potential for bioaccumulation. But it does not reflect the behavior of the substance in the environment and the metabolism in living organisms.

 

Environmental fate

Due to ready biodegradability and high potential of adsorption, the category members can be effectively removed in conventional STPs either by biodegradation or by sorption to biomass. The low water solubility and high estimated log Kow indicate the substance is highly lipophilic. If released into the aquatic environment, the substance undergoes extensive biodegradation and sorption on organic matter, as well as sedimentation. The bioavailability of the substance in the water column is reduced rapidly. The relevant route of uptake of polyol esters in organisms is considered predominately by ingestion of particle bounded substance.

 

Metabolism of polyol esters

Should the substance be taken up by fish during the process of digestion and absorption in the intestinal tissue, polyol esters are expected to be initially metabolized via enzymatic hydrolysis in the corresponding free fatty acids and the free alcohols such as neopentylglycol (NPG), trimethylolpropane (TMP), pentaerythritol (PE) and dipentaerythritol (DiPE). The hydrolysis is catalyzed by classes of enzymes known as carboxylesterases or esterases (Heymann, 1980). The most important of which are the B-esterases in the hepatocytes of mammals (Heymann, 1980; Anders, 1989). Carboxylesterase activity has been noted in a wide variety of tissues in invertebrates as well as in fish (Leinweber, 1987; Suldano et al, 1992; Barron et al., 1999, Wheelock et al., 2008). The catalytic activity of this enzyme family leads to a rapid biotransformation/metabolism of xenobiotics which reduces the bioaccumulation or bioconcentration potential (Lech & Bend, 1980). It is known for esters that they are readily susceptible to metabolism in fish (Barron et al., 1999) and literature data have clearly shown that esters do not readily bioaccumulate in fish (Rodger & Stalling, 1972; Murphy & Lutenske, 1990; Barron et al., 1990). In fish species, this might be caused by the wide CaE distribution, high tissue content, rapid substrate turnover and limited substrate specificity (Lech & Melancon, 1980; Heymann, 1980).

 

Metabolism of enzymatic hydrolysis products

Neopentylglycol (NPG), trimethylolpropane (TMP), pentaerythritol (PE) and dipentaerythritol (DiPE) are the expected possible corresponding alcohol metabolites from the enzymatic reaction of the polyol ester category members. In general, the hydrolysis rate of fatty acid esters and polyol ester in particular varies depending on the fatty acid chain length, and grade of esterification (Mattson and Volpenhein, 1969; Mattson and Volpenhein, 1972a,b).

In the gastrointestinal GI tract(GIT), metabolism prior to absorption via gut microflora or enzymes in the GI mucosa may occur. In fact, after oral ingestion, fatty acid esters with glycerol (glycerides) are rapidly hydrolized by ubiquitously expressed esterases and almost completely absorbed (Mattson and Volpenhein, 1972a).The result of the pancreatic digestion of one NPG ester shows a degradation of the ester of almost 90% within 4 hours (Oßberger, 2012). In contrast with regard to the Polyol esters it was shown that lower rate of enzymatic hydrolysis in the GIT were showed for compounds with more than 3 ester groups (Mattson and Volpenhein, 1972a,b). In vitro hydrolysis rate of pentaerythritol ester was about 2000 times slower in comparison to glycerol esters (Mattson and Volpenhein, 1972a,b).

When hydrolysis occurs the potential hydrolysis products are absorbed and subsequently enter the bloodstream. Potential cleavage products are stepwise degraded via beta–oxidation in the mitochondria. Even numbered fatty acids are degraded via beta-oxidation to carbon dioxide and acetyl-CoA, with release of biochemical energy. In contrast, the metabolism of the uneven fatty acids results in carbon dioxide and an activated C3-unit, which undergoes a conversion into succinyl-CoA before entering the citric acid cycle (Stryer, 1994). Alternative oxidation pathways (alpha- and omega-oxidation) are available and are relevant for degradation of branched fatty acids.

The other cleavage products Polyols (NPG, TMP and PE) are easily absorbed and can either remain unchanged (PE) or may further be metabolized or conjugated (e.g. glucuronides, sulfates, etc.) to polar products that are excreted in the urine (Gessner et al. 1960, Di Carlo et al., 1964).

Lipids and their key constituent fatty acids are, along with protein, the major organic constitute of fish and they play a major role as sources of metabolic energy in fish for growth, reproduction and movement, including migration (Tocher, 2003). In fishes, the fatty acids metabolism in cell covers the two processes anabolism and catabolism. The anabolism of fatty acids occurs in the cytosol, where fatty acids esterified into cellular lipids that is the most important storage form of fatty acids. The catabolism of fatty acids occurs in the cellular organelles, mitochondria and peroxisomes via a completely different set of enzymes. The process is termed beta-oxidation and involves the sequential cleavage of two-carbon units, released as acetyl-CoA through a cyclic series of reaction catalyzed by several distinct enzyme activities rather than a multienzyme complex (Tocher, 2003).

As fatty acids are naturally stored in fat tissue and re-mobilized for energy production is can be concluded that even if they bioaccumulate, bioaccumulation will not pose a risk to living organisms. Fatty acids (typically C14 to C24 chain lengths) are also a major component of biological membranes as part of the phospholipid bilayer and therefore part of an essential biological component for the integrity of cells in every living organism (Stryer, 1994).

 

Data from QSAR calculation

Additional information about this endpoint could be gathered through BCF/BAF calculation using BCFBAF v3.01. The estimated BCF value indicates a low bioaccumulation in organisms (BCF: 3.16 - 550 L/kg, regression based). When including biotransformation rate constants a BCF of 0.89 – 24.7 and a BAF of 0.89 – 24.7 L/kg resulted (Arnot-Gobas estimate, including biotransformation, upper trophic). Even though the members of the polyol ester category are outside the applicability domain of the model they might be used as supporting indication that the potential of bioaccumulation is low. The model training set is only consisting of substances with log Kow values of 0.31 - 8.70. But it supports the tendency that substances with high log Kow values (> 10) have a lower potential for bioconcentration as summarized in the ECHA Guidance R.11 and they are not expected to meet the B/vB criterion (ECHA, 2012).

 

Conclusion

Polyol esters are biotransformed to fatty acids and the corresponding alcohol component by the ubiquitous carboxylesterase enzymes in aquatic species. Based on the rapid metabolism it can be concluded that the high log Kow, which indicates a potential for bioaccumulation, overestimates the bioaccumulation potential of the polyol ester category members. Taking all these information into account, it can be concluded that the bioaccumulation potential of the polyol ester category members is assumed to be low.

 

References:

Anders, M.W. (1989): Biotransformation and bioactivation of xenobiotics by the kidney. In: Hutson, D.H., Caldwell, J. & Paulson, G.D., eds, Intermediary Xenobiotic Metabolism in Animals, New York: Taylor & Francis, pp. 81-97.

Barron, M.G., Charron, K.A., Stott, W.T., Duvall, S.E. (1999): Tissue carboxylesterase activity of rainbow trout. Environmental Toxicology and Chemistry. 18(11), 2506-2511.

Barron, M.G., Mayes, M.A., Murphy, P.G., Nolan, R.J. (1990): Pharmacokinetics and metabolism of triclopyr butoxyethyl ester in coho salmon. Aquatic Tox., 16, 19-32.

Di Carlo F.J., Hartigan J.M. Jr., Couthino, C.B. and Phillips, G.E. (1965): Absorption, distribution and excretion of Pentaerythritol and Pentaerythritol Tetranitrate by mice. Proceedings of the Society for Experimental Biology and Medicine. 118: 311-314.

ECHA (2012): Guidance on information requirements and chemical safety assessment. Chapter R.11: PBT Assessment.

Heymann, E. (1980): Carboxylesterases and amidases. In: Jakoby, W.B., Bend, J.R. & Caldwell, J., eds., Enzymatic Basis of Detoxication, 2nd Ed., New York: Academic Press, pp. 291-323.

Gessner PK, Parke DV, Williams RT (1960) Studies in detoxication. 80. The metabolism of glycols Biochem J 74: 1-5.

Lech, J.J. & Bend, J.R. (1980): Relationship between biotransformation and the toxicity and fate of xenobiotix chemicals in fish. Environ. Health Perspec. 34, 115-131.

Lech, J., Melancon, M. (1980): Uptake, metabolism, and deposition of xenobiotic chemicals in fish. EPA-600 3-80-082. U.S. Environmental Protection Agency, Duluth, MN.

Leinweber, F.J. (1987): Possible physiological roles of carboxylic ester hydrolases. Drug. Metab. Rev. 18: 379-439.

Mattson, F.H. and Volpenhein, R.A. (1969): Relative rates of hydrolysis by rat pancreatic lipase of esters of C2 - C18 fatty acids with C1 – C18 primary n-alcohols. J Lipid Res Vol(10): 271 – 276.

Mattson F.H. and Volpenhein R.A., 1972a: Hydrolysis of fully esterified alcohols containing from one to eight hydroxyl groups by the lipolytic enzymes of rat pancreatic juice. J Lip Res 13, 325-328.

Mattson F.H. and Volpenhein R.A., 1972b: Digestion in vitro of erythritol esters by rat pancreatic juice enzymes. J Lip Res 13, 777-782.

Murphy, P.G., Lutenske, N.E. (1990): Bioconcentration of haloxyfop-methyl in bluegill (Lepomis macrochirus Rafinesque). Environ. Intern. 16, 219-230.

Oßberger, M., 2012:Investigation of the hydrolysis behaviour of Propane-1,2,3-triyl-3,5,5-trimethylhexanoate and 2,2-dimethyl-1,3-propandiolheptanoate. Report No. 3635-12. Croda Europe Limited, Goole, UK.

Rodger, C.A., Stalling, D.L. (1972): Dynamics of an ester of 2,3-D in organs of three fish species.Weed Sci. 20, 101-105.

Stryer, L. (1994): Biochemie. 2nd revised reprint, Heidelberg; Berlin; Oxford: Spektrum Akad.Verlag.

Suldano, S., Gramenzi, F., Cirianni, M., Vittozzi, L. (1992): Xenobiotic-metabolizing enzyme systems in test fish - IV. Comparative studies of liver microsomal and cytosolic hydrolases. Comparative Biochemistry and Physiology Part C: Comparative Pharmacology. 101(1), 117-123.

Tocher, D.R. (2003): Metabolism and Function of Lipid and Fatty Acids in Teleost Fish. Review in Fisheries Science. 11(2), 107-184.

Wheelock, C.E., Phillips, B.M., Anderson, B.S., Miller, J.L., Hammock, B.D. (2008): Applications of carboxylesterase activity in environmental monitoring and toxicity identification evaluations (TIEs). Reviews in Environmental Contamination and Toxicology 195:117-178.