(a) Chloral

• Chem. Abstr. Serv. Reg. No.: 75-87-6
• Chem. Abstr. Serv. Name: Trichloroacetaldehyde
• IUPAC Systematic Name: Chloral
• Synonyms: Anhydrous chloral; 2,2,2-trichloroacetaldehyde; trichloroethanal; 2,2,2-tri-chloroethanal

(b) Chloral hydrate

• Chem. Abstr. Serv. Reg. No.: 302-17-0
• Deleted Chem. Abstr. Serv. Reg. No.: 109128-19-0
• Chem. Abstr. Serv. Name: 2,2,2-Trichloro-1,1-ethanediol
• IUPAC Systematic Name: Chloral hydrate
• Synonyms: Chloral monohydrate; trichloroacetaldehyde hydrate; trichloroacetaldehyde monohydrate; 1,1,1-trichloro-2,2-dihydroxyethane

(a) Chloral

• C₂HCl₃O
• Relative molecular mass: 147.39

(b) Chloral hydrate

• C₂H₃Cl₃O₂
• Relative molecular mass: 165.40

(a) Chloral

• Description: Oily liquid. Pungent, irritating odour (O’Neil et al., 2006)
• Boiling-point: 97.8 °C (O’Neil et al., 2006)
• Melting-point: –57.5 °C (O’Neil et al., 2006)
• Density: 1.510 at 20 °C/relative to H₂O at 4 °C (O’Neil et al., 2006)
• Spectroscopy data: Infrared (prism [4626, 4426]), ultraviolet [5–3], nuclear magnetic resonance [8241] and mass [814] spectral data have been reported (Weast & Astle, 1985; Sadtler Research Laboratories, 1991)
• Solubility: Freely soluble in water, in which it is converted to chloral hydrate; soluble in diethyl ether and ethanol (O’Neil et al., 2006)
• Volatility: Vapour pressure, 10 kPa at 33.8 °C (Haynes, 2012)
• Stability: Polymerizes under the influence of light and in presence of sulfuric acid to form a white solid trimer called metachloral (O’Neil et al., 2006)
• Conversion factor: mg/m³ = 6.03 × ppm, calculated from: mg/m³ = (relative molecular mass/24.45) × ppm, assuming normal temperature (25 °C) and pressure (101 kPa).

(b) Chloral hydrate

• Description: Colourless, transparent, or white crystals with aromatic, penetrating and slightly acrid odour, slightly bitter, caustic taste. (O’Neil et al., 2006)
• Boiling-point: 96 °C, decomposes into chloral and water (Haynes, 2012)
• Melting-point: 57 °C (O’Neil et al., 2006)
• Density: 1.9081 at 20 °C/4 °C (Haynes, 2012)
• Spectroscopy data: Infrared (prism [5423]), nuclear magnetic resonance [10 362] and mass [1054] spectral data have been reported (Weast & Astle, 1985; Sadtler Research Laboratories, 1991)
• Solubility: Very soluble in water, olive oil. Freely soluble in acetone, methyl ethyl ketone (O’Neil et al., 2006)
• Volatility: Vapour pressure, 4.7 kPa at 20 °C; slowly evaporates on exposure to air (Jira et al., 1986; O’Neil et al., 2006)
• Octanol/water partition coefficient (P): Log P, 0.99 (Hansch et al., 1995)
• Conversion factor: mg/m³ = 6.76 × ppm, calculated from: mg/m³ = (relative molecular mass/24.45) × ppm, assuming normal temperature (25 °C) and pressure (101 kPa)

(a) Manufacturing processes

Chloral was first synthesized by J. von Liebig in 1832 by chlorination of ethanol (Jira et al., 2007).

Chloral is produced by chlorinating acetaldehyde or ethanol in acidic solution by gradually increasing the temperature from 0 °C to 90 °C (Jira et al., 2007). Antimony trichloride is sometimes used as a catalyst. Chloral is distilled from the reaction mixture as the hydrate. The hydrate is then mixed with concentrated sulfuric acid, the heavier acid layer is drawn off, and chloral is distilled through a fractionating column of moderate height.

(b) Production volume

Estimated production and use of chloral in the Member States of the European Union in 1984 was 2500 tonnes (Environmental Chemicals Data and Information Network, 1993).

Chloral (anhydrous) is known to have been produced by 14 companies in China, seven companies in India and one company each in Brazil, France, Japan, Mexico, the Russian Federation and the USA. Chloral hydrate was produced by four companies in China, three companies in Germany, two companies in Japan and one company each in Mexico, the Russian Federation and Spain (Chemical Information Services, 2002).

Chloral hydrate has been produced for use as a hypnotic drug in relatively low and gradually declining volume for many years. As an indication of the scale, production in the USA for this purpose was about 135 tonnes in 1978 (Jira et al., 1986).

(a) Chloral

The principal historical use of chloral has been in the production of the insecticide dichlorodiphenyltrichloroethane (DDT) and, to a lesser extent, other insecticides such as methoxychlor, naled, trichlorfon, and dichlorvos and the herbicide trichloroacetic acid (Jira et al., 2007). In the USA in 1975, about 40% of chloral was used in the manufacture of DDT, about 10% in the manufacture of other pesticides and about 50% in other applications (IARC, 1995). After the banning of DDT in many countries, demand for chloral for this use has declined dramatically.

Chloral has also been used in the production of rigid polyurethane foam (IARC 1995; Boĭtsov et al., 1970) and to induce swelling of starch granules at room temperature (Whistler & Zysk, 1978).

(b) Chloral hydrate

Chloral hydrate has been used as a hypnotic drug since the 1870s, principally for the short-term treatment of insomnia. It was also used to allay anxiety and to induce sedation and/or sleep post-operatively, before electroencephalogram evaluations, and to treat the symptoms of withdrawal of alcohol and other drugs such as opiates and barbiturates. For many years chloral hydrate was widely used for the sedation of children before diagnostic, dental or medical procedures, but although still in use, it has largely been replaced by newer drugs with a lower risk of overdose (Pershad et al., 1999).

After oral administration, chloral hydrate is converted rapidly to trichloroethanol, which is largely responsible for its hypnotic action. Externally, chloral hydrate has a rubefacient action (producing redness of the skin) and has been used as a counter-irritant. It is administered by mouth as a liquid or as gelatin capsules. It has also been dissolved in a bland fixed oil and given by enema or as suppositories (Gennaro, 2000; Royal Pharmaceutical Society of Great Britain, 2002).

Chloral hydrate is also an ingredient in Hoyer's solution, which is used in microscopy to mount organisms such as bryophytes, ferns, seeds, and arthropods (Anderson, 1954).

(a) Air

No data are available on human exposure to chloral or chloral hydrate in air. The low volatility of chloral hydrate from a water solution precludes significant exposure by inhalation (EPA, 2000).

(b) Water

When raw water, containing natural organic material such as humic, tannic or amino acids, is treated by chlorination for use as drinking-water or in swimming pools, by-products resulting from disinfection can be formed, such as chloral, which is rapidly transformed into chloral hydrate (Miller & Uden, 1983; Sato et al., 1985; Trehy et al., 1986; Italia & Uden, 1988). Chloral hydrate is primarily produced by the treatment of water with chlorine or chloramine, but the use of pre-ozonation before chlorination or chloramination can favour its formation (Richardson et al., 2007).

Table 1.2 summarizes recent measurements of chloral hydrate in drinking-water and swimming pool water in several countries.

Table 1.2

Concentrations of chloral hydrate in drinking-water.

(a) Humans

Studies in humans have shown that chloral hydrate is rapidly absorbed after oral administration, with peak concentrations in blood occurring within 1 hour after dosing (Merdink et al., 2008). The extent of oral absorption has not been measured precisely in humans, but is likely to be high due to the rapidity of absorption. Recovery of metabolites in urine, which represents a lower boundary on absorption, has been reported to be between 47% and 60% for subjects followed for 1 week after administration of a single oral dose (Müller et al., 1974; Merdink et al., 2008). It should be noted that 1 week is insufficient for complete urinary excretion of the metabolites of chloral hydrate. With repeated oral dosing, Owens & Marshall (1955) reported high inter-individual variation in recovery, with average daily excretion ranging from 7% to 94% of the daily dose of chloral hydrate.

No data were available to the Working Group on absorption in humans via other routes of exposure.

(b) Experimental systems

Orally administered chloral hydrate is rapidly absorbed in rats and mice, with peak concentrations reported at 15 minutes after administration (Beland et al., 1998). The extent of oral absorption has not been measured precisely in experimental systems, but is likely to be high due to the rapidity of absorption. The percentage absorption was not estimated since no studies on urinary excretion or mass balance for orally administered chloral hydrate were available to the Working Group.

No experimental data on absorption via other routes of exposure were available to the Working Group.

(a) Humans

In humans, orally administered chloral hydrate enters the liver where it undergoes extensive metabolism (see Section 4.1.3); only a limited amount enters the systemic circulation (Merdink et al., 2008). Chloral hydrate in the blood is rapidly eliminated by metabolism, as shown by Zimmermann et al. (1998), who reported a half-life of less than 1 hour, and Merdink et al. (2008), who reported a rapid initial decline with a terminal half-life of about 10 hours. [Although no data on distribution in human tissues were available, the Working Group noted that due to the extensive first-pass effect, it was likely that concentrations of chloral hydrate were much lower in extra-hepatic tissues than in the liver of humans exposed orally.]

No data on distribution via other routes of administration were available to the Working Group.

(b) Experimental systems

In rats and mice, orally administered chloral hydrate enters the liver where it undergoes extensive metabolism (see Section 4.1.3); only a limited amount enters the systemic circulation (Beland et al., 1998). Chloral hydrate in the blood is rapidly eliminated, probably by metabolism, with levels reported to be below the limit of detection within 3 hours after oral administration (Beland et al., 1998). [Although no data on tissue distribution were available, the Working Group noted that due to the extensive first-pass effect, it was likely that concentrations of chloral hydrate in extrahepatic tissues were much lower than those in the liver of rodents exposed orally.]

The half-life of chloral hydrate after intravenous administration has been reported to be about 5–20 minutes in mice and rats (Abbas et al., 1996; Merdink et al., 1999), indicating rapid elimination from the systemic circulation. Abbas et al. (1996) reported concentrations of chloral hydrate in the liver to be about one quarter to one half of concentrations in the blood. No data on distribution to other tissues after intravenous administration were available to the Working Group.

No data on distribution via other routes of administration were available to the Working Group.

(a) Humans

The metabolic pathways of chloral hydrate and its metabolites in humans are depicted in Fig. 4.1.

Fig. 4.1

Metabolism of chloral hydrate

Multiple studies in humans have reported that the metabolites of chloral hydrate include trichloroethanol, its glucuronide, and trichloroacetic acid. For instance, Owens & Marshall (1955), Breimer et al. (1974), and Merdink et al. (2008) measured free trichloroethanol, total trichloroethanol (free plus glucuronidated), and trichloroacetic acid in blood and/or urine after administration of chloral hydrate to human volunteers. The terminal half-lives of trichloroethanol and trichloroacetic acid after oral exposure to chloral hydrate have been measured to be about 8–13 hours for trichloroethanol and 4–5 days for trichloroacetic acid (Breimer, 1977; Zimmermann et al., 1998; Merdink et al., 2008).

Merdink et al. (2008) also examined the importance of enterohepatic recirculation, by which trichloroethanol-glucuronide is excreted with bile into the intestine, where trichloroethanol is regenerated and reabsorbed in the gut. In this study, the subjects consumed a high-fat meal 4 hours after dosing with chloral hydrate, to “synchronize” the secretion of bile and subsequent excretion of trichloroethanol-glucuronide into the intestine. The resulting cyclic variation in the amount of trichloroethanol-glucuronide and the complex kinetic behaviour of plasma trichloroacetic acid (e.g. two distinct peak concentrations) are consistent with enterohepatic recirculation.

Considerable amounts of dichloroacetic acid were reported as a urinary metabolite of chloral hydrate in children (Henderson et al., 1997a). However, only trace amounts were detected after administration of chloral hydrate to adults (Merdink et al., 2008). [The Working Group noted that due to uncertainties as to artefactual (ex vivo) formation of dichloroacetic acid in biological samples (Ketcha et al., 1996), it was unclear whether this difference was due to life stage or to the analytical methodologies used.]

Bronley-DeLancey et al. (2006) used cryogenically preserved human hepatocytes to simultaneously evaluate the kinetics of the metabolism of chloral hydrate and alcohol dehydrogenase/aldehyde dehydrogenase (ADH/ALDH) genotype. Thirteen samples of human hepatocytes were examined, and large inter-individual variation in the V_max values for formation of trichloroethanol and trichloroacetic acid was reported. In this sample of limited size, no correlation with ADH/ALDH genotype was apparent. Furthermore, despite the large variation in V_max values between individuals, disposition of chloral hydrate into downstream metabolites was found to be relatively constant.

(b) Experimental systems

A similar spectrum of metabolites has been reported in experimental studies. Studies in rats, mice, and dogs have all identified trichloroacetic acid, trichloroethanol-glucuronide, and trichloroethanol as the major metabolites of chloral hydrate (Breimer et al., 1974; Abbas et al., 1996; Beland et al., 1998; Merdink et al., 1998, 1999).

The importance of enterohepatic recirculation in the kinetics of chloral hydrate metabolites has been examined by Merdink et al. (1999) through comparison between intact and bile-cannulated rats given chloral hydrate, trichloroethanol, and trichloroacetic acid via the jugular vein. A statistically significant difference in the kinetics of trichloroethanol and its glucuronide was reported for between intact and bile-cannulated rats given trichloroethanol. Kinetic differences for chloral hydrate, trichloroethanol and its glucuronide, or trichloroacetic acid after administration of chloral hydrate were evident only at the highest dose of chloral hydrate (192 mg/kg bw, compared with 12 and 48 mg/kg bw). Moreover, chloral hydrate, trichloroethanol and its glucuronide, and trichloroacetic acid were all detected in bile, with trichloroethanol-glucuronide exhibiting the highest peak concentrations. Therefore, while the data are consistent with enterohepatic recirculation of trichloroethanol and its glucuronide occurring in rats, this process appears to have little impact on the kinetics of chloral hydrate or its metabolites in rats, except at higher doses.

Abbas et al. (1996) reported detecting dichloroacetic acid in mice given chloral hydrate by oral administration, but it was later determined that these data were probably confounded by artefactual formation of dichloroacetic acid during sample preparation (Ketcha et al., 1996; Abbas & Fisher, 1997; Merdink et al., 1998). Later studies (Beland et al., 1998; Merdink et al., 1998, 1999) reported only trace or undetectable amounts of dichloroacetic acid in mice or rats given chloral hydrate. Merdink et al. (1998) have suggested that it is likely that some dichloroacetic acid is being formed as a short-lived intermediate, but that the extremely rapid elimination kinetics of dichloroacetic acid relative to its formation do not allow for accumulation (and detection) of dichloroacetic acid in the blood.

Chloral hydrate was shown to be an inhibitor of ALDH (Wang et al., 1999), suggesting that production of trichloroacetic acid from chloral hydrate may not increase in a linear fashion with dose. An inhibitory effect of chloral hydrate on liver ADH was also reported in studies in mice (Sharkawi et al., 1983). In a short-term study in rats, Poon et al. (Poon et al., 2002) showed that exposure to drinking-water containing chloral hydrate led to statistically significant reduction in activity of liver ALDH, while the activity of liver aniline hydroxylase (a marker for CYP2E1) was significantly elevated in males and females receiving chloral hydrate at 200 ppm. In the same study, the findings of Wang et al. (1999) were confirmed, showing that chloral hydrate is a potent inhibitor of liver ALDH in vitro, with an IC₅₀ of 8 μM, while trichloroacetic acid was weakly inhibitory, and trichloroethanol was without effect.

(a) Humans

The primary known excretion route for chloral hydrate in humans is as the metabolites trichloroethanol-glucuronide and trichloroacetic acid in urine (Owens & Marshall, 1955; Merdink et al., 2008). Owens & Marshall (1955) found that recovery of urinary metabolites was not complete, with average daily excretion ranging from 7% to 94% of daily doses of chloral hydrate, according to individual. Therefore, it is possible that other excretion routes exist that have not been well characterized. For instance, low concentrations of chloral hydrate have been found in breast milk. Although breastfeeding infants may be sedated by chloral hydrate in breast milk, the highest concentration measured in milk (about 15 μg/mL) is considerably lower than that measured in blood after administration of chloral hydrate at a clinically active dose (100 μg/mL) (Bernstine et al., 1956; Wilson, 1981).

(b) Experimental systems

In mice and rats, chloral hydrate appears to be excreted primarily in urine as the metabolites trichloroethanol-glucuronide and trichloroacetic acid (Merdink et al., 1998, 1999); however, there were no comprehensive studies of mass balance available to the Working Group. Therefore, the extent of recovery represented by urinary excretion is unknown, although it is often assumed to be 100% (Beland et al., 1998).

Few notable differences have been found in the excretion of chloral hydrate in rats and mice. While Beland et al. (1998) noted statistically significant differences in the half-lives of trichloroethanol and its glucuronide in rats and mice, all estimated half-lives were very short (< 1 hour). Beland et al. (1998) also reported that with the same regime of repeated doses (12 doses of chloral hydrate at 50 or 200 mg/kg bw over 16 days), the area under the concentration–time curve (AUC) of trichloroacetic acid in rats was greater than that in mice.

(a) DNA binding and damage

There has been limited analysis of the DNA-binding potential of chloral hydrate (Keller & Heck, 1988; Ni et al., 1995; Von Tungeln et al., 2002). Keller & Heck (1988) conducted experiments in B6C3F₁ mice in vitro and in vivo. The mice were pretreated by gavage with trichloroethylene at a dose of 1500 mg/kg bw per day for 10 days, and then given [¹⁴C]-labelled chloral intraperitoneally at a dose of 800 mg/kg bw. No detectable covalent binding of the radiolabel to DNA in the liver was observed.

Keller & Heck (1988) investigated the potential of chloral hydrate to form DNA–protein crosslinks in rat liver nuclei. No statistically significant increase in the frequency of DNA–protein crosslinks was observed with chloral hydrate at concentrations of 25, 100, or 250 mM [3.7, 14.7, 36.8 mg/mL]. DNA and RNA isolated from the nuclei treated with [C¹⁴]-labelled chloral did not have any detectable bound radiolabel; however, concentration-dependent binding of the radiolabel to proteins from choral-treated nuclei was observed.

Incubation of chloral hydrate with liver microsomes from male B6C3F₁ mice resulted in increases in the amounts of lipid-peroxidation products (malondialdehyde and formaldehyde). This effect was inhibited by free radical scavengers, α-tocopherol or menadione (Ni et al., 1994). Ni et al. (1995) subsequently observed malondialdehyde adducts in calf thymus DNA in the presence of chloral hydrate and liver microsomes from male B6C3F₁ mice. In another study in B6C3F₁ mice, exposure in vivo to nonradiolabelled chloral hydrate at a concentration of 2000 nmol [330 μg] resulted in an increase in malondialdehyde-derived adducts and 8-oxo-2′-deoxyguanosine adducts in liver DNA, indirect indicators of oxidative DNA damage (Von Tungeln et al., 2002).

Kiffe et al. (2003) carried out the single-cell gel electrophoresis assay (comet assay) in Chinese hamster ovary K5 cells under standard assay conditions or with modifications involving collection of all cells, concurrent treatment with ethyl methanesulfonate (for detecting crosslinking properties) and/or analysis of subcellular DNA breakage. Chloral hydrate gave negative results except at the highest concentration (5000 μg/mL), at which cell viability was about 30%. Zhang et al. (2012) conducted the single-cell gel electrophoresis assay in HepG2 cells to assess the DNA-damaging potential of chloral hydrate. A statistically significant increase (P < 0.01) in DNA damage was reported after treatment with chloral hydrate at 20 μM [3.3 μg/mL] for 4 hours, with cell viability exceeding 75%. Cell viability decreased below 75% at concentrations of 80 μM [13.2 μg/mL] and higher. Liviac et al. (2010) carried out the single-cell gel electrophoresis assay in TK6 cells and reported statistically significant increases in DNA damage at concentrations of 100 μM [16.5 μg/mL] and higher, with no change in cell viability up to the highest concentration tested (10 mM) [1654 μg/mL]. Liviac et al. (2010) also examined DNA-repair kinetics, reporting that induced DNA damage was repaired after 45 minutes. Therefore, the degree of DNA damage appears to depend heavily on the type of cell used in the experimental system.

(b) Mutations

Chloral hydrate induced gene mutation in Salmonella typhimurium TA100 and TA104 strains, but not in most other strains assayed. Four out of six studies of reverse mutation in S. typhimurium TA100 and two out of two studies in S. typhimurium TA104 gave positive results (Haworth et al., 1983; Ni et al., 1994; Giller et al., 1995; Beland, 1999). Waskell (1978) studied the effect of chloral hydrate (dose range, 1.0–13 mg/plate) on gene mutation in different S. typhimurium strains (TA98, TA100, TA1535) in the Ames assay. No revertant colonies were observed in strains TA98 or TA1535 either in the presence and absence of metabolic activation by S9 (9000 × g rat liver supernatant). Similar results were obtained by Leuschner & Leuschner (1991); however, in TA100, a dose-dependent statistically significant increase in the frequency of revertant colonies was obtained in the presence and absence of metabolic activation. It should be noted that the chloral hydrate used (obtained from Sigma), recrystallized one to six times from chloroform, was described as “crude.” However, this positive result was consistent with those of other studies in this strain, as noted above (Waskell, 1978). Giller et al. (1995) studied the genotoxicity of chloral hydrate in three short-term tests. Chloral induced mutations in strain TA100 of S. typhimurium (fluctuation test). Similar results were obtained by Haworth et al. (1983). [The Working Group noted that these results were consistent with those of several studies with trichloroethylene, in which low, but positive, responses were observed in the TA100 strain in the presence of metabolic activation from S9, even when genotoxic stabilizers were not present.]

A statistically significant increase in the frequency of mitotic segregation was observed in Aspergillus nidulans treated with chloral hydrate at a concentration of 5 [827 μg/mL] or 10 mM [1654 μg/mL] (Crebelli et al., 1985). Studies of mitotic crossing-over in A. nidulans gave negative results, while these same studies gave positive results for aneuploidy (Crebelli et al., 1985, 1991; Käfer, 1986; Kappas, 1989).

Two studies in Saccharomyces cerevisiae investigated chromosomal malsegregation after exposure to chloral hydrate (Sora & Agostini Carbone, 1987; Albertini, 1990). Chloral hydrate (1–25 mM) [165–4135 μg/mL] was dissolved in the sporulation medium and the frequencies of various meiotic events such as recombination and disomy were analysed. Chloral hydrate inhibited sporulation as a function of dose and increased the frequency of diploid and disomic clones. Chloral hydrate was also tested for mitotic chromosome malsegregation in S. cerevisiae D61.M (Albertini, 1990). The test strain was exposed at a dose range of 1–8 mg/mL. An increase in the frequency of chromosomal malsegregation was observed after exposure to chloral hydrate.

Two studies of mutagenicity with chloral hydrate were performed in Drosophila (Zordan et al., 1994; Beland, 1999). In these two studies, chloral hydrate gave positive results in the wing spot test for somatic mutation (Zordan et al., 1994), while the results of a test for induction of sex-linked lethal mutation were equivocal when chloral hydrate was administered in the feed, but negative when chloral hydrate was administered by injection (Beland, 1999).

In a mammalian system, Harrington-Brock et al. (1998) noted that increases in mutant frequency observed in Tk ^+/–  mouse lymphoma cell lines treated with chloral hydrate were not statistically significant. The mutants were primarily small-colony Tk mutants, indicating that most mutants induced by chloral hydrate resulted from chromosomal mutation rather than point mutation. It should be noted that cytotoxicity was observed at most concentrations tested (350–1600 μg/mL). Percentage cell survival ranged from 96% to 4%. Fellows et al. (2011) and Liviac et al. (2011) also tested for mutagenicity with chloral hydrate in Tk ^+/–  mouse lymphoma cell lines without metabolic activation from S9. Fellows et al. (2011) reported a statistically significant increase in mutant frequency at the highest tested concentration of 3.4 mM [562 μg/mL], at which a high degree of cytotoxicity was observed. Liviac et al. (2011) tested chloral hydrate at lower concentrations of 1 μM to 1 mM [0.16–165 μg/mL], at which relative total growth was at least 66%, and did not observe any statistically significant increases in mutant frequency. [The Working Group noted that reported increases in mutant frequencies in mammalian systems were not statistically significant, except at highly cytotoxic concentrations.]

(c) Chromosomal effects

Exposure to chloral hydrate induces the formation of micronuclei in most test systems, including assays in vitro and in vivo in mammalian species (Degrassi & Tanzarella, 1988; Leuschner & Leuschner, 1991; Bonatti et al., 1992; Russo et al., 1992; Russo & Levis, 1992a, b; Van Hummelen & Kirsch-Volders, 1992; Leopardi et al., 1993; Lynch & Parry, 1993; Seelbach et al., 1993; Allen et al., 1994; Marrazzini et al., 1994; Giller et al., 1995; Nutley et al., 1996; Parry et al., 1996; Grawé et al., 1997; Harrington-Brock et al., 1998; Beland, 1999; Nesslany & Marzin, 1999; Ikbal et al., 2004; Arkhipchuk & Garanko, 2005; Liviac et al., 2010). For instance, chloral hydrate has been shown to induce micronucleus formation, but not structural chromosomal aberrations in mouse bone-marrow cells. Micronuclei induced by non-clastogenic agents are generally believed to represent intact chromosomes that have failed to segregate into either daughter-cell nucleus at cell division (Xu & Adler, 1990; Russo et al., 1992). Furthermore, micronuclei induced by chloral hydrate in mouse bone-marrow cells (Russo et al., 1992) and in cultured mammalian cells (Degrassi & Tanzarella, 1988; Bonatti et al., 1992) have been shown to be predominantly kinetochore-positive in composition upon analysis with immunofluorescence methods. The presence of a kinetochore in a micronucleus is considered evidence that the micronucleus contains a whole chromosome lost at cell division (Degrassi & Tanzarella, 1988; Hennig et al., 1988; Eastmond & Tucker, 1989). In contrast, Allen et al. (1994) harvested spermatids from male C57B1/6J mice given chloral hydrate by intraperitoneal administration 49 days before harvesting, and found statistically significantly increased frequencies of kinetochore-negative micronuclei; however, no dose–response relationship was observed. The results of this study contrast with those of studies described above (Degrassi & Tanzarella, 1988; Bonatti et al., 1992) which demonstrated predominantly kinetochore-positive micronuclei.

Chloral hydrate induced aneuploidy in vitro in multiple Chinese hamster cell lines (Furnus et al., 1990; Natarajan et al., 1993; Warr et al., 1993) and in human lymphocytes (Vagnarelli et al., 1990; Sbrana et al., 1993), but not in mouse lymphoma cells (Harrington-Brock et al., 1998). Studies performed in vivo in various mouse strains led to increased aneuploidy in spermatocytes (Russo et al., 1984; Liang & Pacchierotti, 1988; Miller & Adler, 1992), but not in oocytes (Mailhes et al., 1993) or bone-marrow cells (Xu & Adler, 1990; Leopardi et al., 1993).

The potential of chloral hydrate to induce aneuploidy in mammalian germ cells has been of particular interest since Russo et al. (1984) first demonstrated that treatment of male mice with chloral hydrate results in statistically significant increases in the frequencies of hyperploidy in metaphase II cells. This hyperploidy was thought to have arisen from chromosomal nondisjunction in premeiotic/meiotic cell division, and may be a consequence of chloral hydrate interfering with spindle formation [reviewed by Russo et al. (1984) and Liang & Brinkley (1985)]. Chloral hydrate also causes meiotic delay, which may be associated with aneuploidy (Miller & Adler, 1992).

Several studies have included analysis of chromosomal aberration in vitro and in vivo after exposure to chloral hydrate; there have been some positive results in vitro (Furnus et al., 1990; Harrington-Brock et al., 1998; Beland, 1999). In mouse lymphoma cell lines and Chinese hamster embryo cells treated with chloral hydrate, there was no significant increase in chromosomal aberration (Harrington-Brock et al., 1998; Furnus et al., 1990). Other studies of chromosome aberration in vivo have mostly reported negative results (Liang & Pacchierotti, 1988; Xu & Adler, 1990; Leuschner & Leuschner, 1991; Russo & Levis, 1992a, b; Mailhes et al., 1993), with the exception of one study (Russo et al., 1984) in mice of an F1 cross, C57B1/Cne × C3H/Cne.

Positive results for sister-chromatid exchange were observed by Beland (1999) in Chinese hamster ovary cells exposed in vitro with and without an exogenous metabolic activation system.

(d) Cell transformation

Chloral hydrate gave positive results in three studies designed to measure cellular transformation in Syrian hamster cells (dermal and/or embryo) exposed to chloral hydrate (Gibson et al., 1995; Parry et al., 1996; Pant et al., 2008).

(a) Cell proliferation

(b) Cell communication

(c) Activation of peroxisome proliferator-activated receptor α (PPARα)

(a) Cytotoxicity

(a) Humans

One study has shown that sedative doses of chloral hydrate to newborns increase the likelihood of hyperbilirubinaemia (Lambert et al., 1990).

(b) Experimental systems

Long-term (104 weeks) exposure of male mice to chloral hydrate at a dose equivalent to 166 mg/kg bw per day resulted in no changes in organ weights except in the liver, which showed an approximate 40% increase in absolute liver weight and in the liver-to-body weight ratio (Daniel et al., 1992a). The same type of study was conducted in male and female rats with the only non-cancer effect reported as focal hepatocellular necrosis in 2 out of 10 male rats (Daniel et al., 1992b).

Similar studies in mice and rats by George et al. (2000) also showed no effects in rats and only proliferative lesions in the liver of mice at doses up to approximately 150 mg/kg bw per day. The National Toxicology Program (NTP, 2002a) conducted studies of long-term exposure in male and female mice given chloral hydrate at doses of up to 71 mg/kg bw per day, and observed no non-neoplastic effects. A long-term (approximately 125 weeks) study by Leuschner & Beuscher (1998) in rats given chloral hydrate at doses of up to 135 mg/kg bw per day showed no increase in non-neoplastic lesions.

The short-term toxicity of chloral hydrate has been studied in CD1 mice and Sprague-Dawley rats. In mice, administration of chloral hydrate at daily doses of 14.4 or 144 mg/kg bw by gavage for 14 consecutive days resulted in an increase in relative liver weight and a decrease in spleen size. No other changes were seen. Mice given drinking-water containing chloral hydrate at a concentration of 0.07 or 0.7 mg/mL for 90 days showed dose-related hepatomegaly in males only and significant changes in hepatic microsomal enzymes in both males and females, indicative of hepatic toxicity (Sanders et al., 1982).

Male Sprague-Dawley rats were given drinking-water containing chloral hydrate at a concentration of 0.13, 1.35, or 13.5 mg/L for 7 days (Poon et al., 2000). No changes were observed in body or organ weights.

(a) Humans

Choral hydrate has long been used in both human and veterinary medicine as a sedative and hypnotic drug. The effects on the central nervous system are due to the metabolite, trichloroethanol (Shapiro et al., 1969; Miller & Greenblatt, 1979).

(b) Experimental systems

Exposure of female CD1 mice to chloral at a dose of 100 ppm [603 μg/L] for 6 hours induced deep anaesthesia, which was fully reversible on cessation of exposure (Odum et al., 1992).

(a) Humans

No other data in humans were available to the Working Group.

(b) Experimental systems

The National Toxicity Program (NTP, 2002a) reported an increase in the incidence of glomerulosclerosis in mice fed ad libitum with diets containing chloral hydrate at a concentration of 25 or 100 mg/kg. Other long-term studies did not report any lesions in the kidney (Daniel et al., 1992a; Leuschner & Beuscher, 1998).