Diisotridecyl adipate

Endpoint summary

Additional information

The bioaccumulation of DITA via aqueous exposure is negligible. The main route of exposure will be via oral uptake. Based on the rapid metabolism of aliphatic esters, it is reasonable to conclude that the high log Kow, which indicates a potential for bioaccumulation, overestimates the true bioaccumulation potential of the substance. Therefore, there remains no uncertainty that bioaccumulation of DITA is unlikely to occur and the Weight of Evidence (WoE) approach is applicable. DITA is considered not to be bioaccumulative (B).

Bioaccumulation via aqueous exposure

DITA is poorly soluble in water (< 0.001 mg/L at 25 °C) and can be judged, in our view, as readily biodegradable but at least as ultimately biodegradable with degradation pattern very similar to that of readily biodegradable substances. According to the Guidance on information requirements and chemical safety assessment, Chapter R.7b, readily biodegradable substances can be expected to undergo rapid and ultimate degradation in most environments, including biological Sewage Treatment Plants (STPs) (ECHA, 2012a). Independent on the final conclusion of ready biodegradability the available biodegradation data indicate that this is also true for DITA. Therefore, after passing through conventional STPs, only a very low concentration of DITA is likely to be (if at all) released into the environment. The Guidance on information requirements and chemical safety assessment, Chapter R.7b (ECHA, 2012a) states that once insoluble chemicals enter a standard STP, they will be extensively removed in the primary settling tank and fat trap and thus, only limited amounts will get in contact with activated sludge organisms. Nevertheless, once this contact takes place, these substances are expected to be removed from the water column to a significant degree by adsorption to sewage sludge based on its high adsorption potential (DITA: log Koc > 7) and the rest will be extensively biodegraded (Guidance on information requirements and chemical safety assessment, Chapter R.7a, (ECHA, 2012b). Considering this one can assume that the availability of the substances in the aquatic environment will be extremely low, which reduces the probability of adsorption and uptake from the surrounding medium into organisms (e.g., see Björk, 1995, Haitzer et al., 1998).

In addition, DITA consists of main components with estimated high partition coefficients (calculated log Kow ≥ 10). Based on the high log Kow and the very low measured water solubility, one can conclude that the substance is hydrophobic and lipophilic (in nature).

If environmental concentrations facilitate exposure, the uptake of DITA from medium into organisms is expected to be very low based on the molecular weight, size and structural complexity of the substance. DITA is an adipic acid ester with two branched side chains of C11 to C13 carbon length. This large and complex structure assumes a high degree of conformational flexibility. Dimitrov et al. (2002) revealed a tendency of decreasing log BCF with an increase in conformational flexibility of molecules, which they assumed to be related to the enhancement of the entropy factor on membrane permeability of chemicals. This concludes a high probability that a substance may encounter the membrane in a conformation which does not enable the substance to permeate. A calculated mean maximum diameter of 25.75 A (lowest Dmax = 18.16 A) using Catalogic clearly supports this assumption.

This interaction between hydrophobicity, bioavailability and membrane permeability is considered to be the main reason why the relationship between the bioaccumulation potential of a substance and its hydrophobicity is commonly found to be described by a relatively steep Gaussian curve with the bioaccumulation peak approximately at log Kow of 6-7 (e.g. see Dimitrov et al., 2002; Nendza & Müller, 2007; Arno and Gobas 2003). Substances with log Kow values above 10, which have been calculated for DITA, are, however, again considered to have a low bioaccumulation potential (e.g. see Nendza & Müller, 2007; 2010). For those substances with a log Kow value > 10 it is recognized by the relevant authorities that it is unlikely that they accomplish the pass level of being bioaccumulative according to OECD criteria for the PBT assessment (log BCF = 2000; ECHA, 2011).

This assumption is also supported by QSAR calculations using BCFBAF v3.01 and Catalogic. BCF values were calculated to be 14.8 (BCFBAF v3.01) using a regression based method and even lower values of 0.982 (BCFBAF v3.01, Arnot-Gobas, upper trophic) and 7.08 (OASIS Catalogic v5.11.9) were calculated if bioavailability and biotransformation processes were taken into account.

Even if based on the high log Kow value the BCFBAF calculation may be rated as outside of the applicability domain the results are congruent with valid calculations for similar fatty acid esters registered under REACH, such as Diisopropyl adipate (CAS 6938-94-9), Dibutyl adipate (CAS 105-99-7), Dihexyl adipate (CAS 110-33-8), Diisopropyl sebacate (CAS 7491-02-3), Dibutyl sebacate (CAs 109-43-3), Decanedioic acid, bis(2-ethylhexyl) ester (CAS 122-62-3) for which BCF and BAF values ranging from <1 to 29 L/kg were calculated.

Taken all these information into account we feel confident that we provide sufficient reliable evidence that bioaccumulation via aqueous exposure is negligible.

We also think that if released into the water phase the substance will to some degree bind to particulate organic matter, and therefore, the main route of exposure for aquatic organisms such as fish will be via food ingestion or contact with suspended solids:

Bioaccumulation via oral uptake

The accumulation of a substance in an organism is determined, not only by uptake, but also by distribution, metabolism and excretion. Accumulation takes place if the uptake rate is faster than the subsequent metabolism and/or excretion.

If taken up by living organisms, aliphatic esters such as DITA will be initially metabolized via enzymatic hydrolysis to the respective dicarboxylic acid and alcohol components as would dietary fats (e.g., Linfield, 1984; Lehninger, 1970; Mattson and Volpenhein, 1972). The hydrolysis is catalyzed by carboxylesterases and esterases, with B-esterases located in hepatocytes of mammals being the most important (e.g., Heymann, 1980). However, carboxylesterase activity has also been reported from a wide variety of tissues in invertebrates and fishes (e.g., Barron et al., 1999; Wheelock et al., 2008). In fish, the high catalytic activity, low substrate specificity and wide distribution of the enzymes in conjunction with a high tissue content lead to a rapid biotransformation of aliphatic esters, which significantly reduces its bioaccumulation potential (Lech & Melancon, 1980; Lech & Bend, 1980).

Alcohols ranging from C11 (Iso-undecanol) to C13 (iso-tridecanol) are the expected hydrolysis products from the enzymatic reaction catalyzed by carboxylesterase. These metabolites exhibit no potential for bioaccumulation (e.g., see published REACH dossiers for isotridecanol [experimental BCF = 2.27/1.41] or adipic acid [valid calculation, BCF = 3.16]). The metabolism of alcohols has been extensively reviewed in the literature (e.g., see Rizzo et al., 1987; Hargrove et al., 2004). The free alcohols can either be esterified to form wax esters (which are similar to triglycerides) or they can be transformed to fatty acids in a two-step enzymatic process catalyzed by alcohol dehydrogenase (ADH) and aldehyde dehydrogenase (ALDH). The responsible enzymes ADH and ALDH are present in a large number of animals including plants, microorganisms and fish (e.g., Sund & Theorell, 1963; Nilsson, 1990; Watabiki et al., 1999; Reimers et al., 2004; Lassen et al., 2005).

The metabolism of alcohols in fish was extensively studied by Reimers et al. (2004). They isolated and characterized two cDNAs from the zebra fish, Danio rerio, encoding ADHs, which showed specific metabolic activity in in-vitro assays with various alcohol components ranging from C4 to C8. The emerging aldehydes were shown to be further oxidized to the corresponding fatty acid by ALDH enzymes. The most effective ALDH2, which is mainly located in the mitochondria of liver cells showed a sequence similarity of 75% to mammalian ALDH2 enzymes and a similar catalytic activity (also see Nilsson, 1988). The same metabolic pathway was shown for longer chain alcohols, such as stearyl and oleyl alcohol in fish (e.g., Sand et al., 1973). Furthermore, cleavage products with high water solubility like adipic acid do not have the potential to accumulate in adipose tissue due to their low log Pow and are thus widely distributed within the body and rapidly eliminated via renal excretion. To a smaller extent the dicarboxylic acids are also metabolised via peroxisomal beta-oxidation.

This assumption about the fate of aliphatic esters such as DITA is confirmed by studies performed with Bis(2-ethylhexyl) adipate (DEHA) (CAS 103-23-1). The potential for accumulation of the poorly soluble, highly lipophilic substance in aquatic organisms was examined in a bioconcentration test with bluegill sunfish (Lepomis macrochirus) using 14C-labelled DEHA (Felder et al., 1986). The test was carried out for 42 days. Concentrations of DEHA in water, whole fish, viscera, and fillet were analyzed at intervals during the test. After the first 35 days of exposure, the remaining fish were exposed to clean water for an additional 14 days and concentrations of DEHA were measured in the fish at intervals. A whole fish bioconcentration factor (BCF) of 27 was reported at day 35. Following exposure to clean water, a depuration rate for DEHA of 0.26/day (t 1/2 = 2.7 days) was determined. The results imply that the accumulation of DEHA is low despite a high log Pow (log Pow = 8.94), most likely due to rapid metabolization. Furthermore, when transferred to freshwater, the substance is apparently rapidly and extensively excreted from the fish. Similar results were observed in monkeys, rats and mice.

Since this experimental results can be easily explained by the general enzymatic processes as mentioned above, which are numerously published in the scientific literature, we can see absolutely no indication that the ADME pattern confirmed for DEHA should not be the same for DITA, a substance which differs to DEHA structurally only slightly in the chain length of the fatty alcohol.

Hence, based on the rapid metabolism we think it is reasonable to conclude that the high log Kow, which indicates a potential for bioaccumulation, overestimates the true bioaccumulation potential of the substance. Therefore, we think that there remains no uncertainty that bioaccumulation of DITA is unlikely to occur.

References

Arnot JA and Gobas FAPC (2003) A generic QSAR for assessing the bioaccumulation potential of organic chemicals in aquatic food webs. QSAR Comb. Sci 22: 337-345

Barron MG et al (1999) Tissue carboxylesterase activity of rainbow trout. Environ Toxicol Chem 18(11): 2506-2511

Björk M (1995) Bioavailability and uptake of hydrophobic organic contaminants in bivalve filter-feeders. Ann Zool Fenn 32(2): 237-245

Dimitrov SD et al (2002) Predicting bioconcentration factors of highly hydrophobic chemicals. Effects of molecular size. Pure Appl Chem 74 (10): 1823-1830

ECHA (2012a) Guidance on information requirements and chemical safety assessment, Chapter R.7b: Endpoint specific guidance, version 2.2 (August 2013), Helsinki, Finland

ECHA (2012b) Guidance on information requirements and chemical safety assessment, Chapter R.7a: Endpoint specific guidance, version 1.2 (November 2012), Helsinki, Finland#

ECHA. (2011) Guidance on information requirements and chemical safety assessment – Part C: PBT assessment, Helsinki, Finland

Heymann E (1980) Carboxylesterases and amidases. Pp 291-316. In: Jakoby WB (ed) Enzymatic basis of detoxification Vol 2. Biochem Pharmacol Toxicol: A series of monographs, Academic Press

Haitzer M et al (1998) Effects of dissolved organic matter (DOM) on the bioconcentration of organic chemicals in aquatic organisms: a review. Chemosphere37(7): 1335-1362

Hargrove JL (2004) Nutritional Significance and Metabolism of Very Long Chain Fatty Alcohols and Acids from Dietary Waxes. Exp Biol Med 229:

Lassen N et al (2005) Molecular cloning, baculovirus expression and tissue distribution of the zebrafish aldehyde dehydrogenase 2. Drug Metabol Disposit 33(5): 649-656

Lech JJ and Bend JR (1980) Relationship Between Biotransformation and the Toxicity and Fate of Xenobiotic Chemicals in Fish. Environmental Health Perspectives 34: 115-131.

Lech JJ and Melancon MJ (1980) Uptake, metabolism, and elimination of 14c‐labeled 1,2,4‐trichlorobenzene in rainbow trout and carp. J Toxicol Environ health 6(3): 645-658

Lehninger AL (1970) Biochemistry. Worth Publishers, Inc.

Linfield WM et al (1984) Enzymatic fat hydrolysis and synthesis. J Am Oil Chem Soc 61(2): 191-195

Mattson FH and Volpenhein RA (1972) 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

Nendz, M and Müller M (2007) Literature Study: Effects of Molecular Size and Lipid Solubility on

Bioaccumulation Potential. Testing laboratory: Fraunhofer Institute for Molecular Biology and Applied Ecology, Schmallenberg, Germany and Analytisches Laboratorium für Umweltuntersuchungen und Auftragsforschung, Luhnstedt, Germany. Report no.: FKZ 360 01 043. Owner company: Umweltbundesamt, Dessau, Germany. Report date: 2007-02-15.

Nendza M and Müller M (2010) Screening for low aquatic bioaccumulation (1): Lipinski’s “Rule of 5” and molecular size. SAR and QSAR in Environ Res 21(5-6): 495-512

Nilsson GE (1990) Distribution of aldehyde dehydrogenase and alcohol dehydrogenase in summer-acclimatized crucian carp, Carassius carassius L. J Fish Biol 36(2): 175-179

Nilsson GE (1988) A comparative study of aldehyde dehydrogenase and alcohol dehydrogenase activities in crucian carp and three other vertebrates: apparent adaptations to ethanol production. J Comp Physiol 158(4): 479-485

Reimers et al (2004) Two Zebrafish Alcohol Dehydrogenases Share Common Ancestry with Mammalian Class I, II, IV, and V Alcohol Dehydrogenase Genes but Have Distinct Functional Characteristics. J Biol Chem 279: 38303-38312.

Rizzo WB et al (1987) Fatty alcohol metabolism in cultured human fibroblasts. Evidence for a fatty alcohol cycle. J Biol Chem 262: 17412-17419

Sand DM et al (1973) Wax ester in fish: Absorption and metabolism of oleyl alcohol in the gourami (Trichogaster cosby). J Nutr 103(4): 600-607

Sund H and Theorell H (1963) Alcohol dehydrogenases. The Enzymes 7: 25-83

Watabiki T et al (1999) Intralobular Distribution of Class I Alcohol Dehydrogenase and Aldehyde Dehydrogenase 2 Activities in the Hamster Liver. Alc: Clinic Experimental Res 23: 52-55

Wheelock CE et al (2008) Applications of Carboxylesterase Activity in Environmental Monitoring and Toxicity Identification Evaluations (TI Es). Rev Environ Contam Toxicol 195: 117-178