INTERNATIONAL PROGRAMME ON CHEMICAL SAFETY
ENVIRONMENTAL HEALTH CRITERIA 167
ACETALDEHYDE
This report contains the collective views of an international group of
experts and does not necessarily represent the decisions or the stated
policy of the United Nations Environment Programme, the International
Labour Organisation, or the World Health Organization.
First draft prepared by Mrs. J. de Fouw, National Institute of Public
Health and Enviromental Protection, Bilthoven, Netherlands
Published under the joint sponsorship of the United Nations
Environment Programme, the International Labour Organisation, and the
World Health Organization
World Health Organization
Geneva, 1995
The International Programme on Chemical Safety (IPCS) is a joint
venture of the United Nations Environment Programme, the International
Labour Organisation, and the World Health Organization. The main
objective of the IPCS is to carry out and disseminate evaluations of
the effects of chemicals on human health and the quality of the
environment. Supporting activities include the development of
epidemiological, experimental laboratory, and risk-assessment methods
that could produce internationally comparable results, and the
development of manpower in the field of toxicology. Other activities
carried out by the IPCS include the development of know-how for coping
with chemical accidents, coordination of laboratory testing and
epidemiological studies, and promotion of research on the mechanisms
of the biological action of chemicals.
WHO Library Cataloguing in Publication Data
Acetaldehyde.
(Environmental health criteria ; 167)
1.Acetadehyde - adverse effects 2.Enviromental exposure I.Series
ISBN 92 4 157167 5 (NLM Classification: QU 99)
ISSN 0250-863X
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CONTENTS
ENVIRONMENTAL HEALTH CRITERIA FOR ACETALDEHYDE
Preamble
Introduction
1. SUMMARY
1.1 Identity, physical and chemical properties,
and analytical methods
1.2 Sources of human and environmental exposure
1.3 Environmental transport, distribution, and
transformation
1.4 Environmental levels and human exposure
1.5 Kinetics and metabolism
1.5.1 Absorption, distribution, and elimination
1.5.2 Metabolism
1.5.3 Reaction with other components
1.6 Effects on organisms in the environment
1.6.1 Aquatic organisms
1.6.2 Terrestrial organisms
1.7 Effects on experimental animals and in vitro test
systems
1.7.1 Single exposure
1.7.2 Short- and long-term exposures
1.7.3 Reproduction, embryotoxicity, and
teratogenicity
1.7.4 Mutagenicity and related end-points
1.7.5 Carcinogenicity
1.7.6 Special studies
1.8 Effects on humans
1.9 Evaluation of human health risks and effects on the
environment
2. IDENTITY, PHYSICAL AND CHEMICAL PROPERTIES, AND ANALYTICAL
METHODS
2.1 Identity
2.2 Physical and chemical properties
2.3 Conversion factors
2.4 Analytical methods
3. SOURCES OF HUMAN AND ENVIRONMENTAL EXPOSURE
3.1 Natural occurrence
3.2 Anthropogenic sources
3.2.1 Production
3.2.1.1 Production levels and processes
3.2.1.2 Emissions
3.2.2 Uses
3.2.3 Waste disposal
3.2.4 Other sources
4. ENVIRONMENTAL TRANSPORT, DISTRIBUTION, AND TRANSFORMATION
4.1 Transport and distribution between media
4.2 Abiotic degradation
4.3 Biodegradation
5. ENVIRONMENTAL LEVELS AND HUMAN EXPOSURE
5.1 Environmental levels
5.1.1 Air
5.1.2 Water
5.1.3 Soil
5.1.4 Food
5.1.5 Cigarette smoke
5.2 General population exposure
5.3 Occupational exposure
6. KINETICS AND METABOLISM IN LABORATORY ANIMALS AND HUMANS
6.1 Absorption
6.2 Distribution
6.2.1 Animal studies
6.2.1.1 Distribution after inhalation
exposure
6.2.1.2 Distribution to the embryo and
fetus
6.2.1.3 Distribution to the brain
6.2.2 Human studies
6.3 Metabolism
6.3.1 Animal studies
6.3.1.1 Liver
6.3.1.2 Respiratory tract
6.3.1.3 Kidneys
6.3.1.4 Testes and ovaries
6.3.1.5 Embryonic tissue
6.3.1.6 Metabolism during pregnancy
6.3.2 Human studies
6.4 Elimination
6.5 Reaction with cellular macromolecules
6.5.1 Proteins
6.5.2 Nucleic acids
7. EFFECTS ON ORGANISMS IN THE ENVIRONMENT
7.1 Aquatic organisms
7.2 Terrestrial organisms
8. EFFECTS ON EXPERIMENTAL ANIMALS AND IN VITRO TEST SYSTEMS
8.1 Single exposure
8.1.1 LD50 and LC50 values
8.2 Short-term exposure
8.2.1 Oral
8.2.2 Inhalation
8.2.3 Dermal
8.2.4 Parenteral
8.3 Skin and eye irritation; sensitization
8.4 Long-term exposure
8.4.1 Oral
8.4.2 Inhalation
8.5 Reproductive and developmental toxicity
8.6 Mutagenicity and related end-points
8.6.1 Bacteria
8.6.2 Non-mammalian eukaryotic systems
8.6.2.1 Gene mutation assays
8.6.2.2 Chromosome alterations
8.6.3 Cultured mammalian cells
8.6.3.1 Gene mutation assays
8.6.3.2 Chromosome alterations and sister
chromatid exchange
8.6.4 In vivo assays
8.6.4.1 Somatic cells
8.6.4.2 Germ cells
8.6.5 Other assays
8.6.5.1 DNA single-strand breaks
8.6.5.2 DNA cross-linking
8.6.6 Cell transformation
8.7 Carcinogenicity bioassays
8.7.1 Inhalation exposure
8.7.2 Co-carcinogenicity and promotion studies
8.8 Neurological effects
8.9 Immunological effects
8.9.1 Direct effects on immune cells
8.9.2 Generation of antibodies reacting with
acetaldehyde-modified proteins
8.9.3 Related immunological effects
8.10 Biochemical effects
9. EFFECTS ON HUMANS
9.1 General population exposure
9.2 Occupational exposure
9.2.1 General observations
9.2.2 Clinical studies
9.2.3 Epidemiological studies
9.3 Effects of endogenous acetaldehyde
9.3.1 Effects of ethanol possibly attributable to
acetaldehyde or acetaldehyde metabolism
10. EVALUATION OF HUMAN HEALTH RISKS AND EFFECTS ON THE
ENVIRONMENT
10.1 Evaluation of human health risks
10.1.1 Exposure
10.1.2 Health effects
10.1.3 Approaches to risk assessment
10.2 Evaluation of effects on the environment
11. RECOMMENDATIONS FOR RESEARCH
REFERENCES
RESUME
RESUMEN
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This publication was made possible by grant number
5 U01 ES02617-15 from the National Institute of Environmental Health
Sciences, National Institutes of Health, USA, and by financial support
from the European Commission.
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WHO TASK GROUP ON ENVIRONMENTAL HEALTH CRITERIA FOR ACETALDEHYDE
Members
Mrs I. Arts, Department of Biological Toxicology, TNO Nutrition and
Food Research, Zeist, The Netherlands
Dr R.E. Barry, Faculty of Medicine, University of Bristol, Bristol
Royal Infirmary, Bristol, United Kingdom
Professor D. Beritic-Stahuljak, Andrija œtampar School of Public
Health, Faculty of Medicine, University of Zagreb, Zagreb, Croatia
Dr Sai Mei Hou, Karolinska Institute, Huddinge, Sweden
Dr M.E. Meek, Environmental Health Directorate, Priority Substances
Section, Health & Welfare Canada, Tunney's Pasture, Ottawa, Canada
(Chairperson)
Professor M.H. Noweir, Industrial Engineering Department, College of
Engineering, King Abdul Aziz University, Jeddah, Saudi Arabia
Professor G. Obe, University of Essen, Essen, Germany
Professor T.V.N. Persaud, Department of Anatomy, Faculty of
Medicine, University of Manitoba, Winnipeg, Manitoba, Canada
Mr D. Renshaw, Department of Health, Elephant & Castle, London, United
Kingdom
Dr A. Smith, Health and Safety Executive, Toxicology Unit, Bootle,
Merseyside, United Kingdom (Co-rapporteur)
Professor A. Watanabe, Toyama Medical and Pharmaceutical University,
Faculty of Medicine, Toyama, Japan
Dr S. Worrall, Department of Biochemistry, University of Queensland,
Brisbane, Queensland, Australia
Representatives from other organizations
Dr V. Krutovskikh, Programme of Multistage Carcinogenesis,
International Agency for Research on Cancer, Lyon, France
Secretariat
Mrs J. de Fouw, National Institute of Public Health and Environmental
Protection, Bilthoven, The Netherlands
(Co-rapporteur)
Professor F. Valic, IPCS Consultant, World Health Organization,
Geneva, Switzerland, also Vice-Rector, University of Zagreb,
Zagreb, Croatia (Responsible Officer and Secretary)
ENVIRONMENTAL HEALTH CRITERIA FOR ACETALDEHYDE
A WHO Task Group on Environmental Health Criteria for
Acetaldehyde met in Geneva from 6 to 10 December 1993. Professor F.
Valic opened the meeting on behalf of the three cooperating
organizations of the IPCS (UNEP/ILO/WHO). The Task Group reviewed and
revised the draft monograph and made an evaluation of the risks for
human health and the environment from exposure to acetaldehyde.
The first draft of this monograph was prepared by Mrs J. de Fouw,
National Institute of Public Health and Environmental Protection,
Bilthoven, The Netherlands.
Professor F. Valic was responsible for the overall scientific
content of the monograph and for the organization of the meeting, and
Mrs M.O. Head of Oxford for the technical editing of the monograph.
The efforts of all who helped in the preparation and finalization
of this publication are gratefully acknowledged.
INTRODUCTION
This monograph will deal mainly with the effects of direct
exposure to acetaldehyde. However, it should be borne in mind that
for most people exposure to acetaldehyde will occur through the
consumption of alcoholic beverages (IARC, 1988). These beverages
contain ethanol, which is metabolized to acetaldehyde by alcohol
dehydrogenase (ADH). ADH activity has been detected in nearly every
tissue including liver, kidney, muscle, intestine, ovary, and testes
(Buehler et al., 1983; Agarwal & Goedde, 1990).
However, data concerning metabolically formed acetaldehyde will
only be considered when no data are available on direct exposure.
The accurate determination of acetaldehyde in body fluid and
tissue samples is relatively difficult. Only the most recent
techniques take into account artifactual acetaldehyde formation in
biological samples, especially those containing ethanol (Eriksson &
Fukunaga 1993). As values for concentrations of acetaldehyde given in
older references may well have been overestimates, absolute values are
only given when necessary.
1. SUMMARY
1.1 Identity, physical and chemical properties, and analytical
methods
Acetaldehyde is a colourless, volatile liquid with a pungent
suffocating odour. The reported odour threshold is 0.09 mg/m3.
Acetaldehyde is a highly flammable and reactive compound that is
miscible in water and most common solvents.
Analytical methods are available for the detection of
acetaldehyde in air (including breath) and water. The principal
method is based on the reaction of acetaldehyde with
2,4-dinitrophenylhydrazine and subsequent analysis of the hydrazone
derivatives by high pressure liquid chromatography or gas
chromatography.
1.2 Sources of human and environmental exposure
Acetaldehyde is a metabolic intermediate in humans and higher
plants and a product of alcohol fermentation. It has been identified
in food, beverages, and cigarette smoke. It is also present in
vehicle exhaust and in wastes from various industries. Degradation of
hydrocarbons, sewage, and solid biological wastes produces
acetaldehyde, as well as the open burning and incineration of gas,
fuel oil, and coal.
More than 80% of the acetaldehyde used commercially is produced
by the liquid-phase oxidation of ethylene with a catalytic solution of
palladium and copper chlorides. Production in Japan was 323 thousand
tonnes in 1981. In the USA, production was 281 thousand tonnes in 1982
while in Western Europe it was 706 thousand tonnes in 1983. Most
acetaldehyde produced commercially is used in the production of acetic
acid. It is also used in flavourings and foods.
The annual emission of acetaldehyde from all sources in the USA
is estimated to be 12.2 million kg.
1.3 Environmental transport, distribution, and transformation
Because of its high reactivity, intercompartmental transport of
acetaldehyde is expected to be limited. Some transfer of acetaldehyde
to air from water and soil is expected because of the high vapour
pressure and low sorption coefficient.
It is suggested that the photo-induced atmospheric removal of
acetaldehyde occurs predominantly via radical formation. Photolysis is
expected to contribute another substantial fraction to the removal
process. Both processes cause a reported daily loss of about 80% of
atmospheric acetaldehyde emissions. Reported half-lives of
acetaldehyde in water and air are 1.9 h and 10-60 h, respectively.
Acetaldehyde is readily biodegradable.
1.4 Environmental levels and human exposure
Levels of acetaldehyde in ambient air generally average
5 µg/m3. Concentrations in water are generally less than
0.1 µg/litre. Analysis of a wide range of foodstuffs in the
Netherlands showed that concentrations, generally less than 1 mg/kg,
occasionally ranged up to several 100 mg/kg, particularly in some
fruit juices and vinegar.
By far, the main source of exposure to acetaldehyde for the
majority of the general population is through the metabolism of
alcohol. Cigarette smoke is also a significant source of exposure.
With respect to other media, the general population is exposed to
acetaldehyde principally from food and beverages, and, to a lesser
extent, from air. The contribution from drinking-water is negligible.
Available data are inadequate to determine the extent of exposure
to acetaldehyde in the workplace. Workers may be exposed in some
manufacturing industries and during alcohol fermentation, where the
principal route of exposure is most likely inhalation with possible
dermal contact.
1.5 Kinetics and metabolism
1.5.1 Absorption, distribution, and elimination
Available studies on toxicity indicate that acetaldehyde is
absorbed through the lungs and gastrointestinal tract; however, no
adequate quantitative studies have been identified. Absorption
through the skin is probable.
Following inhalation by rats, acetaldehyde is distributed to the
blood, liver, kidney, spleen, heart, and other muscle tissues. Low
levels were detected in embryos after maternal intraperitoneal (ip)
injection of acetaldehyde (mouse) and following maternal exposure to
ethanol (mouse and rat). Potential production of acetaldehyde has
also been observed in rat fetuses and in the human placenta,
in vitro.
Distribution of acetaldehyde to brain interstitial fluid, but not
to brain cells, has been demonstrated following ip injection of
ethanol. A high affinity, low Km ALDHa may be important in
maintaining low levels of acetaldehyde in the brain during the
metabolism of ethanol.
Acetaldehyde is taken up by red blood cells and, following
ethanol consumption in humans and in baboons, in vivo,
intracellular levels can be 10 times higher than plasma levels.
Following oral administration, virtually no unchanged
acetaldehyde is excreted in the urine.
1.5.2 Metabolism
The major pathway for the metabolism of acetaldehyde is by
oxidation to acetate under the influence of NADb-dependent ALDH.
Acetate enters the citric acid cycle as acetyl-CoA. There are several
isoenzymes of ALDH with different kinetic and binding parameters that
influence acetaldehyde oxidation rates.
ALDH activity has been localized in the respiratory tract
epithelium (excluding olfactory epithelium) in rats, in the renal
cortex and tubules in the dog, rat, guinea-pig, and baboon, and, in
the testes in the mouse.
Acetaldehyde is metabolized by mouse and rat embryonic tissue
in vitro. Acetaldehyde crosses the rat placenta, in spite of
placental metabolism.
Though there is some metabolism of acetaldehyde in human renal
tubules, the liver is the most important metabolic site.
Several isoenzymic forms of ALDH have been identified in the
human liver and other tissues. There is polymorphism for the
mitochondrial ALDH. Subjects who are homozygous or heterozygous for a
point mutation in the mitochondrial ALDH corresponding gene have low
activity of this enzyme, metabolize acetaldehyde slowly, and are
intolerant of ethanol.
The metabolism of acetaldehyde can be inhibited by
crotonaldehyde, dimethylmaleate, phorone, disulfiram, and calcium
carbamide.
a ALDH = acetaldehyde dehydrogenase.
b NAD = nicotinamide adenine dinocleotide.
1.5.3 Reaction with other components
Acetaldehyde forms stable and unstable adducts with proteins.
This can impair protein function, as evidenced by inhibition of enzyme
activity, impaired histone-DNA binding, and inhibition of
polymerization of tubulin.
Unstable adducts of acetaldehyde of undetermined significance
occur in vitro with nucleic acids.
Acetaldehyde can react with various macromolecules in the body,
preferentially those containing lysine residues, which can lead to
marked alterations in the biological function of these molecules.
1.6 Effects on organisms in the environment
1.6.1 Aquatic organisms
LC50s in fish ranged from 35 (guppy) to 140 mg/litre (species
not specified). An EC5 of 82 mg/litre and an EC50 of 42 mg/litre
were reported for algae and Daphnia magna, respectively.
1.6.2 Terrestrial organisms
Acetaldehyde in air appears to be toxic for some microorganisms
at relatively low concentrations.
Aphids were killed when exposed to acetaldehyde at a
concentration of 0.36 µg/m3 for 3 or 4 h.
Median lethal values were 8.91 mg/litre per h and 7.69 mg/litre
per h for the slug species, Arion hortensis and Agriolimax
reticulatus, respectively.
Inhibition of seed germination in the onion, carrot, and tomato
by acetaldehyde (up to 1.52 mg/litre) was reversible, whereas
inhibition of Amaranthus palmeri, similarly exposed, was
irreversible. Acetaldehyde at 0.54 µg/m3 damaged lettuce.
1.7 Effects on experimental animals and in vitro test systems
1.7.1 Single exposure
LD50s in rats and mice and LC50s in rats and Syrian hamsters
showed that the acute toxicity of acetaldehyde is low. Acute dermal
studies were not identified.
1.7.2 Short- and long-term exposures
In repeated dose studies, by both the oral and inhalation routes,
toxic effects at relatively low concentrations were limited
principally to the sites of initial contact. In a 28-day study in
which acetaldehyde at 675 mg/kg body weight (no-observedeffect level
(NOEL): 125 mg/kg body weight) was administered in the drinking-water
to rats, effects were limited to slight focal hyperkeratosis of the
forestomach. Following administration of a single dose level (0.05%
in the drinking-water) for 6 months (estimated by the Task Group to be
approximately 40 mg/kg body weight) in a biochemical study,
acetaldehyde induced synthesis of rat liver collagen, an observation
that was supported by in vitro data.
Following inhalation, NOELs for respiratory effects were
275 mg/m3 in rats exposed for 4 weeks and 700 mg/m3 in hamsters
exposed for 13 weeks. At lowest-observed-effect levels, degenerative
changes were observed in the olfactory epithelium in rats
(437 mg/m3) and the trachea (2400 mg/m3) in hamsters.
Degenerative changes in the respiratory epithelium and larynx were
observed at higher concentrations. No repeated dose dermal studies
were identified.
1.7.3 Reproduction, embryotoxicity, and teratogenicity
In several studies, parenteral exposure of pregnant rats and mice
to acetaldehyde induced fetal malformations. In the majority of these
studies, maternal toxicity was not evaluated. No data on reproductive
toxicity were identified.
1.7.4 Mutagenicity and related end-points
Acetaldehyde is genotoxic in vitro, inducing gene mutations,
clastogenic effects, and sister-chromatid exchanges (SCEs) in
mammalian cells in the absence of exogenous metabolic activation.
However, negative results were reported in adequate tests on
Salmonella. Following intraperitoneal injection, acetaldehyde induced
SCEs in the bone marrow of Chinese hamsters and mice. However,
acetaldehyde administered intraperitoneally did not increase the
frequency of micronuclei in early mouse spermatids. There is indirect
evidence from in vitro and in vivo studies to suggest that
acetaldehyde can induce protein-DNA and DNA-DNA cross-links.
1.7.5 Carcinogenicity
Increased incidences of tumours have been observed in inhalation
studies on rats and hamsters exposed to acetaldehyde. In rats, there
were dose-related increases in nasal adenocarcinomas and squamous cell
carcinomas (significant at all doses). However, in hamsters,
increases in nasal and laryngeal carcinomas were non-significant. All
concentrations of acetaldehyde administered in the studies induced
chronic tissue damage in the respiratory tract.
1.7.6 Special studies
Adequate studies on the potential neuro- and immunotoxicity of
acetaldehyde were not identified.
1.8 Effects on humans
In limited studies on human volunteers, acetaldehyde was mildly
irritating to the eyes and upper respiratory tract following exposure
for very short periods to concentrations exceeding approximately 90
and 240 mg/m3, respectively. Cutaneous erythema was observed in
patch testing with acetaldehyde, in twelve subjects of "Oriental
ancestry".
One limited investigation in which the incidence of cancer was
examined in workers exposed to acetaldehyde and other compounds has
been reported.
On the basis of indirect evidence, acetaldehyde has been
implicated as the putatively toxic metabolite in the induction of
alcohol-associated liver damage, facial flushing, and developmental
effects.
1.9 Evaluation of human health risks and effects on the environment
The acute toxicity of acetaldehyde by the inhalation or oral
route in studies conducted on animals was low. According to studies
on humans and animals, acetaldehyde is mildly irritating to the eyes
and the upper respiratory tract. In limited studies on human
volunteers, acetaldehyde was mildly irritating to the eyes and upper
respiratory tract (section 1.8). Cutaneous erythema has also been
observed in the patch testing of humans. Although a possible mechanism
has been identified, available data are inadequate to assess the
potential of acetaldehyde to induce sensitization.
Available data on the effects of acetaldehyde following ingestion
are limited. Following oral administration of 675 mg/kg body weight
per day to rats, a borderline increase in hyper-keratosis of the
forestomach was observed (NOEL: 125 mg/kg body weight). In rats
exposed to a dose level of approximately 40 mg acetaldehyde/kg body
weight in the drinking-water for 6 months, there was an increase in
collagen synthesis in the liver, the significance of which is unclear.
On the basis of studies on rats and hamsters, the target tissue
in inhalation studies is the upper respiratory tract. In available
studies, the lowest concentration at which effects were observed was
437 mg/m3 following administration for 5 weeks. The NOELs
identified for respiratory effects were 275 mg/m3 in rats exposed
for 4 weeks and 700 mg/m3 in hamsters exposed for 13 weeks.
At concentrations that induced tissue damage in the respiratory
tract, increased incidences were observed of nasal adenocarcinomas and
squamous cell carcinomas in the rat and laryngeal and nasal carcinomas
in the hamster.
There is evidence to suggest that acetaldehyde causes genetic
damage to somatic cells in vivo.
Available data are inadequate for the assessment of the potential
reproductive, developmental, neurological, or immunological effects
associated with exposure to acetaldehyde in the general, or
occupationally exposed, populations.
On the basis of data on irritancy in humans, a tolerable
concentration of 2 mg/m3 has been derived. Since the mechanism of
induction of tumours by acetaldehyde has not been well studied, two
approaches were adopted for the provision of guidance with respect to
this end-point, i.e., the development of a tolerable concentration
based on division of an effect level for irritancy in the respiratory
tract of rodents by an uncertainty factor, and, estimation of lifetime
cancer risk based on linear extrapolation. The tolerable
concentration is 0.3 mg/m3. The concentrations associated with a
10-5 excess lifetime risk are 11-65 µg/m3.
The limited available data preclude definitive conclusions
concerning the potential risks of acetaldehyde for environmental
biota. However, on the basis of the short half-lives of acetaldehyde
in air and water and the fact that it is readily biodegradable, the
impact of acetaldehyde on organisms in the aquatic and terrestrial
environments is expected to be low, except, possibly, during
industrial discharges or spills.
2. IDENTITY, PHYSICAL AND CHEMICAL PROPERTIES, AND ANALYTICAL METHODS
2.1 Identity
Chemical formula: C2H4O
Chemical structure: CH3-CHO
Common name: acetaldehyde
Common synonyms: ethanal; acetic aldehyde; acetylaldehyde;
ethylaldehyde; diethylacetal;
1,1-diethyoxy ethane
CAS chemical name: acetaldehyde
CAS registry number: 75-07-0
RTECS registry number: AB 1925000
2.2 Physical and chemical properties
The most important physical and chemical properties of
acetaldehyde are given in Table 1.
Acetaldehyde is a volatile liquid with a pungent, suffocating
odour that is fruity in dilute concentrations. The odour threshold for
acetaldehyde is reported to be 0.09 mg/m3 (0.05 ppm). This was a
geometric average of all available literature data (Amoore & Hautala,
1983). In the case of carbon dioxide solutions in acetaldehyde, the
acetaldehyde odour is weakened by the carbon dioxide (Hagemeyer,
1978).
Acetaldehyde is a highly reactive compound that undergoes
numerous condensation, addition, and polymerization reactions. It
decomposes at temperatures above 400°C, forming principally methane
and carbon monoxide. Acetaldehyde is highly flammable when exposed to
heat or flame, and, in air, it can be explosive. Acetaldehyde can
react violently with acid anhydrides, alcohols, ketones, phenols,
NH3, HCN, H2S, P, halogens, isocyanates, strong alkalies, and
amines. It is miscible in all proportions with water and the most
common organic solvents. In aqueous solutions, acetaldehyde exists in
equilibrium with the hydrate, CH3 CH(OH)2. The enol form, vinyl
alcohol (CH2=CHOH) exists in equilibrium with acetaldehyde to the
extent of approximately one molecule per 30 000 (Hagemeyer, 1978).
Table 1. Physical and chemical properties of acetaldehydea
Colour colourless
Relative molecular mass 44.1
Boiling point at 101.3 kPa 20.2°C
Melting point -123.5°C
Octanol/water partition coefficient as 0.63
log Pow
Flash point, closed cup -38°C
Autoignition temperature 185-193°C
Explosion limits of mixtures with air 4.5-60.5 vol % acetaldehyde
Vapour pressure at -50°C 2.5 kPa
0°C 44.0 kPa
20.16°C 101.3 kPa
Specific gravity (20/4) 0.778
Relative vapour density 1.52
Refractive index 20/D 1.33113
Dissociation constant at 0°C, Ka 0.7 × 10-14
Solubility miscible in water and most
common solvents
a From: Hagemeyer (1978); IPCS/CEC (1990).
Commercial acetaldehyde should have the following typical
specifications: purity, 99% min; acidity (as acetic acid), 0.1% max,
and a specific gravity of 0.804-0.811 (0°/20°C) (US NRC, 1981).
2.3 Conversion factors
1 mg acetaldehyde/m3 air = 0.56 ppm at 25°C and 101.3 kPa
(760 mmHg).
1 ppm = 1.8 mg acetaldehyde/m3 air.
2.4 Analytical methods
Several analytical procedures used for the sampling and
determination of acetaldehyde in various media are summarized in
Table 2.
Table 2. Sampling, preparation, and determination of acetaldehydea
Medium Sampling method Analytical method Detection Sample Comments Reference
limit size
Air collection in a midget HPLC with 18 µg/m3 20 litre designed for analysis of Lipari & Swarin
impinger containing 2,4-DNPH spectrophotometric automobile exhaust (1982)
in acetonitrile with detection
perchloric acid as catalyst
Air collection in a tube containing HPLC with 0.9 µg/m3 2 litre suitable for analysis of Jarke et al.
a thermal stable organic polymer spectrophotometric indoor and outdoor air (1981)
based on 2,6-diphenyl-p- detection
phenylene oxide
Air adsorption on a silica gel GC-FTD 0.09-0.45 50-100 suitable for analysis of Aoyama & Yashiro
treated with 2,4-DNPH µg/m3 litre smog and automobile (1983)
exhaust
Air collection in a 2,4-DNPH HPLC with < 18 µg/m3 < 20 litre suitable for long-term Tejada (1986)
coated Sep-PAK cartridge, spectrophotometric sampling at low µg/m3
acidified with HCl detection (ppb) levels in ambient
air, or, for short-term
sampling at low mg/m3
(ppm) levels in diluted
automotive exhaust
emissions
Table 2 (contd).
Medium Sampling method Analytical method Detection Sample Comments Reference
limit size
Air collection in annular denuders HPLC with UV 0.36 µg/m3 100 litre suitable for analysis of Possanzini et
coated with 2,4-DNPH absorbance or outdoor and indoor al. (1987)
detection voltametric
pollution
Air collection and derivatization HPLC with UV 90 µg/m3 5 litre suitable for personal Binding et al.
on 2,4-DNPH coated detection monitoring of 5-min, (1986)
Chromosorb P short-term values as
well as for continuous
sampling over a whole
work shift
Air collection on DNPH-coated C18 HPLC with UV 12 ng per suitable for ambient Grosjean (1991)
cartridge detection cartridge monitoring
Air collection and derivatization HPLC with UV 32 mg/m3b 60 litre suitable for short-term US NIOSH (1987)
in midget bubblers containing detection exposure sampling;
Girard T solution interference with other
aldehydes and volatile
ketones should be
considered
Table 2 (contd).
Medium Sampling method Analytical method Detection Sample Comments Reference
limit size
Air collection in a Chromosorb 104 GC-FID 0.1 µg/litre 1.5 litre suitable for monitoring Watanabe (1988)
tube installed in an automated of outdoor and indoor
sampler pollution
Air collection on a XAD-2 sorbent GC-FID 1.3 mg/m3c 3 litre suitable for short-term US NIOSH (1989)
coated with 2-(hydroxymethyl)- exposure sampling and for
piperidine analysis of field samples
Water derivatization in a two-phase HPLC with 21 µg -- designed for analysis of Facchini et
system by addition of 2,4-DNPH electrochemical fog and rain water al. (1986)
and isooctane detection
Water purging with nitrogen gas and sweeping by rapid 200 µg/ 5 ml suitable for analysis of Spingarn et
collection on a Tenax GC heating of trap litre aqueous solution and al. (1982)
sorbent and silica gel trap into GC-MS industrial effluent
Water derivatization with 2,4-DNPH HPLC; the reaction < 10 µg per 1 ml suitable for routinely Steinberg &
(in acetonitrile) mixture is analysed sample monitoring rain, fog, Kaplan (1984)
directly, without and mist samples
prior separation of
the DNPH-derivatives
Table 2 (contd).
Medium Sampling method Analytical method Detection Sample Comments Reference
limit size
Water collection in a PTFE-cartridge HPLC with UV 0.3 µg/litre 500 ml designed for analysis of Takami et
packed with sulfonated cation detection water samples at the low al. (1985)
exchange resin charged with µg/litre levels
2,4-DNPH
Water collection of aqueous solution HS-GC-FID 25 µg/litre 10 ml designed for the Gramiccioni
in vials, no special treatments quantification of et al. (1986)
released from plastics acetaldehyde
into aqueous foods
Water collection on cyanogen bromide spectrophotometric 0.6 mg/ 30 µlitre immobilized aldehyde Almuaibed &
activated Sepharose 4B detection litre dehydrogenase makes the Townshend (1987)
containing aldehyde determination more
dehydrogenase; soluble aldehyde economic and simpler
dehydrogenase injected in the
sampler flow stream using a
double injection technique
Beverage collection of the 2,6- HPLC with 0.01 µg per 15 ml designed for analysis of Okamoto et
dimethylpyridine derivative spectrophotometric sample wine al. (1981)
on a 3-aminopropyl- detection
triethoxysiloxane or a
Nucleosil 5NH2 treated silica
gel with propionaldehyde as
internal standard
Table 2 (contd).
Medium Sampling method Analytical method Detection Sample Comments Reference
limit size
Beverage steam distillation followed by HPLC with UV ± 5 µg/ 500 ml designed for analysis of Piendl et
liquid liquid extraction, detection litre beer al. (1981)
derivatization with p-nitrobenzyl-
oxyamine-hydrochloride with T-2
undecenal as internal standard
Beverage conversion of acetaldehyde HS-GC-FID ± 1 mg/ 5 ml designed as a rapid Jones et al.
acetals and bisulfite addition litre means by which the (1986)
products to free acetaldehyde by acetaldehyde production
a series of 1-min acid, base, and consumption
and iodine treatments followed pattern of different
by a 10-min equilibration period wines can be predicted
Breast collection of volatile compounds thermal -- 60 ml designed for Pellizari et
milk on a Tenax cartridge after desorption determination in breast al. (1982)
warming milk and purging with into GC-MS milk
helium
Blood precipitation of protein with GC headspace 4.4 µg per - designed for analysis of Eriksson et
perchloric acid analysis sample blood in order to reduce al. (1982)
artificial formation
of acetaldehyde
Blood derivatization with 2,4-DNPH HPLC with UV 4.4 ng per 2 ml designed for analysis of Pezzoli et
with butyraldehyde as internal detection sample blood al. (1984)
standard and perchloric acid
(for protein precipitation)
Table 2 (contd).
Medium Sampling method Analytical method Detection Sample Comments Reference
limit size
Blood rapid separation plasma: HPLC with > 0.9 ng 0.5 ml designed for analysis of Di Padova et
plasma: deproteinization and spectrophotometric per sample plasma plasma and red blood al. (1986)
derivatization with 2,4-DNPH detection cells
haemolysate: deproteinization haemolysate:
and mixed with semicarbazide HS-GC-FID
hydrochloride
Blood separation of plasma and HPLC with 11 µg per 1 ml suitable for clinical Peterson &
haemolysate plasma: fluorescence sample plasma use Polizzi (1987)
1,3-cyclo-hexanedione and detection or RBC
isooctane haemolysate: haemolysate
1,3-cyclo-hexadione both in
presence of ammonium ion
Blood reaction with 1,3-cyclo-hexanone HPLC with 4.4 µg per 50-100 designed for microassays Ung-Chhun &
in the presence of ammonium ion fluorescence sample µlitre with negligible Collins (1987)
propionaldehyde used as internal detection interference
standard
Blood collection in an organic solution HPLC with > 0.13 µg 1 ml designed for analysis of Rideout et
of 2-diphenylacetyl- spectrofluometric per sample blood; with minor al. (1986)
1,3-indandione-1-hydrazone, and detection modifications also
forming fluorescent azine suitablefor analysis of
derivative-precipitation of beverages, breath, and
proteins tissue
Table 2 (contd).
Medium Sampling method Analytical method Detection Sample Comments Reference
limit size
Blood reaction with methanolic solution HPLC; the 4.4 µg/litre 1 ml blood suitable for assessment Lynch et
and of 2,4-DNPH, with acetaldehyde adduct 1 g tissue of acetaldehyde levels in al. (1983)
tissue dinitrophenyl-[14C]-formaldehyde was identified by clinical and experimental
as internal standard co-chromatography studies of ethanol
with the authentic metabolism and alcoholic
derivative and by beverages
mass spectrometry
a 2,4-DNPH = 2,4-dinitrophenylhydrazine; HPLC = high pressure liquid chromatography; GC-FID = gas chromatography with flame ionization
detection; GC-FTD = gas chromatography with flame thermionic detection; GC-MS = gas chromatography with mass spectrometric detection;
HS-GC-FID = head space gas chromatography with flame ionization detection.
b minimum working range (estimated LOD: 1.6 mg/m3).
c minimum working range (estimated LOD: 0.67 mg/m3).
The most specific and sensitive analytical method, widely used to
date, is based on the reaction of acetaldehyde with
2,4-dinitro-phenylhydrazine (2,4-DNPH) and the subsequent analysis of
the hydrazone derivatives by high pressure liquid chromatography
(HPLC) or gas chromatography (GC). Methods mentioned by US NIOSH are
based on derivatization with Girard T solution followed by HPLC
analysis with UV detection (US NIOSH, 1987), or, on derivatization
with 2-(hydroxymethyl)piperidine followed by GC analysis with a flame
ionization detector (FID) (US NIOSH, 1989). In the method based on
Girard T derivation, other volatile aldehydes compete for the Girard T
reagent. Chromatographic conditions may be adjusted to resolve
acetaldehyde from other aldehydes.
Spingarn et al. (1982) determined volatile organic compounds in
aqueous solutions, including acetaldehyde, using a technique in which
the compounds were purged from the solution by bubbling with an inert
gas into a trap containing a Tenax sorbent and silica gel. The
analytes were separated by GC and detected with either specific
ionization detection or MS. An improvement in detection limits,
compared with those of the widely used spectrophotometric method of
analysing carbonyls in aqueous solution, was obtained by Facchini et
al. (1986) by means of an electrochemical detector.
In the determination of acetaldehyde in blood, two main
difficulties exist. The first is related to its disappearance from
blood prior to measurement and the second is related to the formation
of acetaldehyde in blood after collection. According to Pezzoli et
al. (1984), the addition of butyraldehyde to blood, as an internal
standard, immediately after withdrawal, obviates some of the
inconveniences in the determination of acetaldehyde in blood. The
addition of butyric acid makes it possible to obtain results both for
the interaction of the aldehyde group of the acetaldehyde with amino
groups, and for the formation and extraction of the derivative
compound. However, Di Padova et al. (1986) stated that the addition
of butyraldehyde was not specific for the determination of the
acetaldehyde but was related to the aldehyde group reactivity.
Therefore, they described an improved procedure for measuring
acetaldehyde in plasma, based on rapid separation, 2,4-DNPH
derivatization, and HPLC analysis, and a procedure for measuring
acetaldehyde in red blood cells, based on the use of a semicarbazide
solution and analysis by head space gas chromatography.
3. SOURCES OF HUMAN AND ENVIRONMENTAL EXPOSURE
3.1 Natural occurrence
Acetaldehyde is a metabolic intermediate in humans and higher
plants and it is a product of alcohol fermentation (IARC, 1985). It
has been identified as a volatile component of mature cotton leaves
and cotton blossoms (Berni & Stanley, 1982) and as a component in the
essential oil of alfalfa at a concentration of about 0.2% (Kami,
1983). It occurs in food, various fruits, and several spices (see
section 5.1.4) and in oak and tobacco leaves (Furia & Bellanca, 1975;
US NRC, 1985).
Acetaldehyde is formed in the atmosphere in a variety of ways.
It is generated by the oxidation of non-methane hydrocarbons both in
the background troposphere and in photochemical smog (Grosjean, 1982).
3.2 Anthropogenic sources
3.2.1 Production
3.2.1.1 Production levels and processes
Until 1968, most acetaldehyde produced in the USA was made by the
partial oxidation of ethanol over a silver catalyst; however,
currently less than 5% of US production is based on this process. The
liquid-phase oxidation of ethylene using a catalytic solution of
palladium and copper chlorides was first used commercially in the USA
in 1960 and more than 80% of the world production of acetaldehyde is
made by this process. The remainder is produced by the oxidation of
ethanol and the hydration of acetylene. Acetaldehyde is produced by a
limited number of companies over the world. The total production of
acetaldehyde in the USA in 1982 amounted to 281 thousand tonnes.
Total acetaldehyde production in Western Europe in 1982 was 706
thousand tonnes, and the production capacity was estimated to have
been nearly 1 million tonnes. In Japan, the estimated production in
1981 was 323 thousand tonnes (Hagemeyer, 1978; IARC, 1985).
3.2.1.2 Emissions
Eimutis et al. (1978) estimated that the annual atmospheric
emissions of acetaldehyde in the USA amounted to 12.2 thousand tonnes
(Table 3). Emissions of acetaldehyde in the Netherlands in the year
1980 were reported to be 584 tonnes (Guicherit & Schulting, 1985).
Table 3. Emission and sources of acetaldehyde in the USA
Source Emissions
(tonnes/year)
Residential external combustion of wood 5056.4
Coffee roasting 4411.4
Acetic acid manufacture 1460.9
Vinyl acetate manufacture from ethylene 1094.6
Ethanol manufacture 57.8
Acrylonitrile manufacture 51.6
Acetic acid manufacture from butane 20.8
Crotonaldehyde manufacture 4.5
Acetone and phenol manufacture from cumene 1.9
Acetaldehyde manufacture by hydration of ethylene 0.5
Polyvinyl chloride manufacture 0.2
Acetaldehyde manufacture by oxidation of ethanol 0.1
3.2.2 Uses
Acetaldehyde is an important intermediate in the production of
acetic acid, ethyl acetate, peracetic acid, pentaerythritol, chloral,
glyoxal, alkylamines, and pyridines (Hagemeyer, 1978). The use
pattern for the estimated 281 thousand tonnes of acetaldehyde produced
in the USA in 1982 was as follows: acetic acid 61%, pyridine and
pyrine bases 9%, peracetic acid 8%, pentaerythritol 7%, 1,3-butylene
glycol 2%, chloral 1%, and other applications (including use as a food
additive and exports) 12%. The use pattern for the estimated 706
thousand tonnes of acetaldehyde produced in Western Europe was as
follows: acetic acid 62%, ethyl acetate 19%, pentaerythritol 5%,
synthetic pyridines 3%, and all other uses 11% (IARC, 1985).
Acetaldehyde is used for the flavourings: berry, butter,
chocolate, apple, apricot, banana, grape, peach, black walnut, and
rum, and it is used in the following foods: beverages, ice cream and
ices, candy, baked goods, gelatin desserts, and chewing gum (Furia &
Bellanca, 1975; US NRC, 1981, 1985). Acetaldehyde is also used in
perfumes, aniline dyes, plastics, in the manufacture of synthetic
rubber, in the silvering of mirrors, in the hardening of gelatin
fibres, and in the laboratory (Verschueren, 1983).
3.2.3 Waste disposal
Degradation of hydrocarbons, sewage, and solid biological wastes
produces acetaldehyde. It has been detected in effluents from
sewage-treatment plants and chemical plants (US EPA, 1975; Shackelford
& Keith, 1976).
Acetaldehyde has been identified as a constituent in the wastes
from petroleum refining, coal processing, the oxidation of alcohols,
saturated hydrocarbons, or ethylene, and the hydration of acetylene
(IARC, 1985).
3.2.4 Other sources
Acetaldehyde is detected as a combustion product of plastics and
polycarbonate and polyurethane foams of western European origin
(Hagen, 1967; Boettner et al., 1973).
Acetaldehyde occurs in vehicle exhaust at levels of
1.4-8.8 mg/m3 in gasoline exhaust, about 5.8 mg/m3 in diesel
exhaust (Verschueren, 1983), and 51.6% acetaldehyde/ n-hexane GC peak
area ratio in exhaust gas oxygenates (Hugues & Hum, 1960). It also
occurs in the open burning and incineration of gas, fuel oil, and
coal, and evaporation products of perfumes (Verschueren, 1983).
Acetaldehyde has been identified in fresh tobacco leaves and in
tobacco smoke (concentrations ranging from 2.1 to 4.6 mg/litre smoke)
(Buyske et al., 1956; Osborne et al., 1956; Mold & McRae, 1957).
When Lipari et al. (1984) measured aldehyde emissions from
wood-burning fireplaces, they ranged from 0.08 to 0.20 g/kg of wood
burned, based on tests with cedar, jack pine, red oak, and green ash.
Acetaldehyde emissions from wood-burning furnaces and
stoves were also measured in a Swedish study (Rudling et al., 1981)
and in a Norwegian study (Ramdahl et al., 1982). In the Swedish
study, the emissions ranged from 1-72 mg/kg wood in prechamber ovens
to 9-710 mg/kg wood in fireplace stoves. In the Norwegian study, the
reported emissions from stoves were 14.4 mg/kg dry wood under normal
burning conditions and up to 992 mg/kg dry wood under low-efficiency
combustion.
4. ENVIRONMENTAL TRANSPORT, DISTRIBUTION, AND TRANSFORMATION
4.1 Transport and distribution between media
Acetaldehyde can enter the atmosphere during production of the
compound itself, as a product of incomplete combustion, and also as a
by-product of fermentation (Grosjean, 1982).
Photochemical oxidation of acetaldehyde has been shown to be an
important process in the chemistry of photochemical smog (Bagnall &
Sidebottom, 1984; Leone & Seinfeld, 1984). Present theories ascribe
the importance of acetaldehyde to its being a precursor of
peroxyacetylnitrate (PAN) in polluted atmospheres (Kopczynski et al.,
1974; Grosjean et al., 1983; Bagnall & Sidebottom, 1984; Moortgat &
McQuigg, 1984). Acetaldehyde is likely to be a precursor of acetic
acid, which is a component of natural precipitation and contributes to
its acidity (Moortgat & McQuigg, 1984).
Intercompartmental transport of acetaldehyde is expected to be
limited, because of its high reactivity. However, because of the high
vapour pressure of acetaldehyde, some transfer to air from water and
soil can be expected.
The tendency of acetaldehyde to adsorb on soil particles can be
expressed in terms of Koc, the ratio of the amount of chemical
adsorbed per unit weight of organic carbon to the concentration of the
chemical in solution at equilibrium. On the basis of the available
empirical relationships derived for estimating Koc, a low soil
adsorption potential is expected (Lyman et al., 1982). Koch & Nagel
(1988) calculated a soil sorption coefficient of 0.90 for
acetaldehyde, and, therefore, acetaldehyde was classified as a
compound with a very low sorption tendency.
4.2 Abiotic degradation
It is suggested that photo-induced atmospheric removal of
acetaldehyde occurs predominantly via radical formation. Singh et al.
(1982) reported that photolysis and reaction with hydroxyl radicals
cause a daily loss rate of about 80% of atmospheric acetaldehyde
emissions. Grosjean et al. (1983) reported that the reaction with
hydroxyl radicals could remove 50-300 tonnes of carbonyls from the Los
Angeles air over a 12-h daytime period and, thus, is considered to be
a major removal process for all aldehydes. The absolute rate constant
for the reaction of the hydroxyl radical with acetaldehyde was
determined over a temperature range of 26-153°C by Atkinson & Pitts
(1978). At 26°C, they obtained a rate constant of (1.60 ± 0.16) ×
10-11 cm3 per molecule per second. This results in a half-life
for acetaldehyde of 10 h, using a 12-h daytime average hydroxyl
radical concentration of 2 × 10-15 mol/litre (Lyman et al., 1982).
Hustert & Parlar (1981) reported that 49.5% acetaldehyde was
photochemically degraded (reaction with hydroxyl radicals) after a 2-h
radiation (lambda > 230 nm) at 25°C, which, contrary to Atkinson &
Pitts (1978), shows a half-life of 2 h. Atkinson et al. (1984)
obtained a rate constant of (1.34 ± 0.28) × 10-15 for the gas-phase
reaction of nitrate radicals with acetaldehyde at 25°C. This results
in a half-life for acetaldehyde of 59.6 h using a 12-h nighttime
average nitrate radical concentration of 4.0 × 10-12 mol/litre
(Atkinson et al., 1987).
There is a considerable amount of evidence that acetaldehyde in
aqueous solution is in equilibrium with its hydrated form
CH3CH(OH)2. The degree of hydration decreases with increasing
temperature (e.g., at 0°C, the fraction of acetaldehyde hydrated is
0.73; at 25°C, it is 0.59) (Bell & Clunie, 1952).
Von Burg & Stout (1991) reported a half-life of 1.9 h for
acetaldehyde in river water; no other details were provided.
4.3 Biodegradation
Several studies have revealed significant degradation of
acetaldehyde by mixed cultures obtained from sludges and settled
sewage. Hatfield (1957) reported the ability of acclimatized sludge
to oxidize acetaldehyde (major portion of the biological and chemical
oxygen demand (BOD and COD) removed within a 4-h aeration period).
Ludzack & Ettinger (1960) determined the BOD for acetaldehyde in
activated sludge at 20°C and found that 93% of the acetaldehyde was
removed after an observation period of 1/3-5 days and an
acclimatization period of 30 days. Thom & Agg (1975) and Speece
(1983) also reported that acetaldehyde was easily biodegradable by
biological sewage treatment (additional information was not provided).
However, Gerhold & Malaney (1966) reported little degradation of
acetaldehyde by unacclimatized municipal sludge with a BOD of 27.6% as
a percentage of the theoretical oxygen demand in 24 h.
Acetaldehyde is also degraded by anaerobic biological treatment
with unacclimatized acetate-enriched cultures. A COD-removal of 97%
was obtained at the end of a 90-day acclimatization period in
completely mixed reactors with a 20-day hydraulic retention time, no
solids recycle, and a final daily feed concentration of
10 000 mg/litre (Chou & Speece, 1978).
Acetaldehyde is reported to be readily biodegradable using the
biodegradability MITI test, defined in OECD Guidelines for testing of
chemicals (OECD, 1992).
5. ENVIRONMENTAL LEVELS AND HUMAN EXPOSURE
5.1 Environmental levels
5.1.1 Air
The concentrations of acetaldehyde in uncontaminated Arctic air
masses, determined over a 24-h period, ranged from not detected to
0.54 µg/m3 (Cavanagh et al., 1969).
In samples collected during April 1981, the levels of
acetaldehyde in the air in Pittsburg (PA) and Chicago (Il) were
0.36-4.68 µg/m3 and 1.62-6.12 µg/m3, respectively (Singh et al.,
1982). In samples collected at 7 other locations in the USA between
1975 and 1978, mean concentrations in ambient air were 5-124 µg/m3
(Brodzinsky & Singh, 1982).
Schulam et al.(1985) determined the levels of acetaldehyde in air
(June-August 1983) in the urban location of Schenectady (NY) and the
rural location of Whiteface Mountain (NY). Concentrations of
acetaldehyde were similar in the two locations (the levels of
acetaldehyde varied from 0.36 to 1.44 µg/m3, detection limit:
0.29 µg/m3).
The average ambient atmospheric level of acetaldehyde, measured
during the four seasons at Brookhaven National Laboratory (Upton, Long
Island, NY) from July 1982 to May 1983, was 5.2 µg/m3, with a mean
minimum concentration in winter of 1.8 µg/m3 and a mean maximum
value in summer of 15.1 µg/m3 (Tanner & Meng, 1984). Concentrations
of acetaldehyde in the air in Tulsa, OK (sampled in July 1978), Rio
Blanco County, CO (sampled in July 1978), and the Great Smoky
Mountains, TN (sampled in September 1978), ranged up to 14.9, 16.9,
and 23.9 µg/m3, respectively (Arntz & Meeks, 1981).
Mean concentrations of acetaldehyde in the air in Tokyo during
four seasons in 1985-86 ranged from 2.2 to 7.3 µg/m3 (Watanabe,
1987). Seasonal trends were not noted. Concentrations of
acetaldehyde in an environmental survey conducted by the Japan
Environment Agency in 1987 ranged from 0.9 to 22 µg/m3 (number of
sites sampled unspecified) (Japan Environment Agency, 1989).
The mean concentration of acetaldehyde was 2 µg/m3 at three
locations in the Netherlands, namely, the island of Terschelling (one
of the least polluted areas of the country), Delft (suburban), and
Vlaardingen (heavily industrialized area) (Guicherit & Schulting,
1985).
Grosjean (1991) reported levels of acetaldehyde in ambient air,
sampled every sixth day over a one-year period, at six locations in
Southern California between September 1988 and September 1989.
Concentrations ranged up to 23.3 µg/m3 (13 ppb) with average values
at the various locations ranging from 5.2 to 8.6 µg/m3
(2.9-4.8 ppb).
The mean concentration of acetaldehyde in fog samples taken in
November, 1985 in the Po Valley (Italy) was 21 µg/litre (Facchini et
al., 1986). At urban locations in California (Los Angeles) and Alaska
(Fairbanks), concentrations of acetaldehyde ranged from 0.007 to
0.13 µg/ml in ice fog (Alaska), 0 to 0.11 µg/ml in rain (CA), 0 to
0.59 µg/ml in cloud (CA), 0.10 to 0.11 µg/ml in mist (CA), and 0.006
to 0.17 µg/ml in fog (CA) (Grosjean & Wright, 1983).
5.1.2 Water
No quantitative data on concentrations of acetaldehyde in raw
water supplies were identified.
Acetaldehyde has been detected in drinking-water from
Philadelphia and Seattle at levels of up to 0.1 µg/litre (Keith et
al., 1976). No other information was provided.
5.1.3 Soil
Data on concentrations of acetaldehyde in soil were not
identified.
5.1.4 Food
Acetaldehyde has been detected in a wide range of foodstuffs (US
NRC, 1981, 1985; Horvath et al., 1983; Feron et al., 1991), though few
quantitative data are available. In a variety of foodstuffs analysed
in the Netherlands including fruits and juices, vegetables, milk
products, bread, eggs, fish, meat, and alcoholic beverages,
concentrations were generally less than 1 mg/kg, but occasionally
ranged up to several hundred mg/kg, particularly in some fruit juices
and alcoholic beverages; in vinegar, a maximum value of 1060 mg/kg was
reported (Maarse & Visscher, 1992). Acetaldehyde has been identified
in alcoholic beverages, such as beer and wine (Okamoto et al., 1981;
Piendl et al., 1981; Jones et al., 1986); levels in 18 English beers
ranged from 2.6 to 13.5 mg/litre (Delcour et al., 1982). Levels of
0.2 to 1.2 mg/litre were found in wine samples in Japan (Okamato et
al., 1981), while Margeri et al. (1984) reported levels of
acetaldehyde in wines ranging between about 30 and 80 µg/litre.
Acetaldehyde has been detected, but not quantified, in breast
milk in the USA (detection limit not reported) (Pellizari et al.,
1982).
5.1.5 Cigarette smoke
Acetaldehyde is present in tobacco leaves and in cigarette smoke
(Furia & Bellanca, 1975; US NRC, 1985). Hoffman et al. (1975)
detected acetaldehyde in the smoke of tobacco (980 µg per cigarette)
and marijuana (1200 µg/cigarette). The concentration in smoke from
several cigarettes ranged from 0.87 to 1.22 mg per cigarette or from
1.14 to 1.37 mg/cigarette, depending on the method of detection. The
concentration of acetaldehyde in three types of low-tar cigarettes
ranged from 0.09 to 0.27 mg/cigarette (Manning et al., 1983).
5.2 General population exposure
Acetaldehyde is a metabolic product of ethanol. On the basis of
the assumptions that a standard drink contains 10 g of ethanol and
that about 90% of imbibed alcohol is metabolized to acetaldehyde,
alcoholic beverages are generally by far the most significant source
of exposure to acetaldehyde for the general population.
On the basis of the content of acetaldehyde in cigarettes
reported in section 5.1.5, it is likely that cigarettes contribute
significantly to the total intake of acetaldehyde by smokers.
Assuming that smoke contains about 1 mg acetaldehyde per cigarette,
that 20 cigarettes are smoked per day, and a mean adult body weight of
64 kg (WHO, in press), intake from mainstream smoke would be about
300 µg/kg body weight per day.
On the basis of the average dietary intake of food groups in
different regions of the world (WHO, in press) and the contents of
acetaldehyde in foodstuffs and non-alcoholic beverages in the
Netherlands (Maarse & Visscher, 1992), food (particularly fruit
juices) may be one of the principal sources of exposure to
acetaldehyde in the general environment. More representative data on
mean concentrations in foodstuffs have not been identified, but, on
the basis of the ranges of concentrations determined in the Dutch
survey, intake in food is estimated to range from just less than 10 to
several hundred µg/kg body weight per day.
Data from recent studies in various locations in the world
indicate that mean concentrations of acetaldehyde in ambient air range
from 2 to 8.6 µg/m3 (Guicheret & Schulting, 1985; Watanabe, 1987;
Grosjean, 1991) (section 5.1.1). Data on concentrations of
acetaldehyde in indoor air were not identified. On the basis of a
daily inhalation volume for adults of 22 m3, a mean body weight for
males and females of 64 kg (WHO, in press), and the assumption that
mean concentrations are approximately 5 µg/m3, the mean intake of
acetaldehyde from ambient air for the general population is estimated
to be 1.7 µg/kg body weight per day.
Limited identified data on concentrations of acetaldehyde in
drinking-water indicate that they are generally less than 0.1 µg/litre
(Keith et al., 1976). Assuming a daily volume of ingestion for adults
of 1.4 litres and a mean body weight for males and females of 64 kg
(WHO, in press), and that levels are less than 0.1 µg/litre, the
estimated intake of acetaldehyde from drinking-water for the general
population would not exceed 0.002 µg/kg body weight per day.
5.3 Occupational exposure
Workers are exposed to acetaldehyde in the organic chemicals
industry and in the fabricated rubber, plastic, and fermentation
industries (US NIOSH, 1980, 1981). Concentrations of acetaldehyde
were below the detection limits (1-3.4 mg/m3) in five studies in
which the workroom air of plants, such as those in textile finishing,
propylene bottle production, and ureaformaldehyde foam-insulation
manufacturing, was monitored (Rosensteel & Tanaka, 1976; Ahrenholz &
Gorman, 1980; Herrick, 1980; Chrostek & Shoemaker, 1981; Chrostek,
1981). Bittersohl (1975) reported levels of acetaldehyde of
1-7 mg/m3 in the hydrogenation unit of a chemical factory after
equipment leakages.
Concentrations of acetaldehyde to which workers may be exposed
near aircraft with low-smoke combustor engines were found to range
from 139 to 394 µg/m3 (Miyamoto, 1986).
6. KINETICS AND METABOLISM IN LABORATORY ANIMALS AND HUMANS
6.1 Absorption
No studies are available on animals or humans concerning the
absorption of acetaldehyde. However, the results of toxicity studies
indicate that absorption via the lungs and gastrointestinal tract does
occur. The physical and chemical properties of acetaldehyde indicate
that absorption via the skin is also possible.
6.2 Distribution
6.2.1 Animal studies
6.2.1.1 Distribution after inhalation exposure
Distribution studies were conducted on overnight-starved, male
Sprague-Dawley rats exposed (whole-body) to unknown concentrations of
acetaldehyde vapour for 1 h. Acetaldehyde was recovered in total
blood, liver, kidneys, spleen, heart muscle, and skeletal muscle. No
other tissues were studied. The concentration of acetaldehyde in the
liver was relatively low (Hobara et al., 1985; Watanabe et al., 1986).
This can be attributed to rapid metabolism by hepatocytes.
6.2.1.2 Distribution to the embryo and fetus
No studies are available concerning routes of relevance to
humans.
Acetaldehyde was detected in the embryo up to 2 h after maternal
ip injection of 200 mg acetaldehyde/kg body weight in CD-1 mice on day
10 of gestation; acetaldehyde was measured within 5 min of injection.
Following maternal ip injection of 79 mg ethanol/kg body weight,
acetaldehyde was measured up to 12 h after injection; however, levels
were low and approached the limit of sensitivity (Blakley & Scott,
1984b).
Several other studies have demonstrated the presence of
acetaldehyde in the embryos of rats (Espinet & Argiles, 1984; Gordon
et al., 1985; Guerri & Sanchis, 1985; Clarke et al., 1986),
guinea-pigs, and ewes (Clarke, 1988) exposed to ethanol. Embryological
and cytogenic studies with ethanol and acetaldehyde in preimplantation
mouse embryos in vitro showed that acetaldehyde is three times more
toxic than ethanol. It has been suggested that the preimplantation
mouse embryo is able to convert ethanol to acetaldehyde, and that the
enzyme involved is alcohol dehydrogenase (ADH) (Lau et al., 1991)
6.2.1.3 Distribution to the brain
In the only study involving a route of relevance to humans,
following a single intragastric administration of 4500 mg ethanol/kg
body weight to male and female Wistar rats, acetaldehyde was detected
in the blood and in brain interstitial fluid collected from the
caudate nucleus and the thalamushypothalamus region. Following
administration of disulfiram (an inhibitor of the aldehyde
dehydrogenase (ALDH)-catalysed oxidation of acetaldehyde to acetate)
20 h prior to exposure to ethanol, there was a 6-fold increase in the
concentration of acetaldehyde in the blood and brain. Although
acetaldehyde was found in interstitial fluid, none was detected in
whole brain (Westcott et al., 1980).
In albino rats treated first with pyrazole, an inhibitor of ADH,
injected (ip) with a solution of acetaldehyde in saline (200 mg/kg
body weight per day) for 10 days, and then sacrificed 30 min after
receiving the last injection, acetaldehyde was detected in the brain,
liver, and blood (Prasanna & Ramakrishnan, 1984b, 1987).
A study by Pettersson & Kiessling (1977) indicated the importance
of ALDH activity, with a low Michaelis constant, in maintaining a low
level of brain acetaldehyde during ethanol metabolism. They detected
acetaldehyde and ethanol in the cerebrospinal fluid of rats after
intraperitoneal administration of ethanol alone or of ethanol followed
by acetaldehyde.
6.2.2 Human studies
The percentage of acetaldehyde retained by 8 volunteers inhaling
acetaldehyde vapour (100-800 mg/m3) from a recording respirometer
ranged from 45 to 70%, at different respiratory rates. Total
respiratory tract retention was the same whether the vapour was
inhaled through the nose or the mouth. A direct relationship was
found between the contact time and uptake, independent of rate. Thus,
the critical factor in determining acetaldehyde uptake is the duration
of the ventilatory cycle (Egle, 1970).
Baraona et al. (1987) used the blood of 5 healthy individuals, 6
alcoholic patients, and 2 baboons to show that, after alcohol
consumption, most of the blood acetaldehyde was found in the red blood
cells. In vivo, the acetaldehyde concentration in red cells was
about 10 times higher than that in the plasma. No significant
variations were seen between the 3 groups.
Studies using the perfused human placental cotyledon indicated
that the human placenta has the potential to produce acetaldehyde,
which can enter the fetal circulation. Furthermore, partial transfer
of acetaldehyde from maternal to fetal blood may occur (Karl et al.,
1988).
6.3 Metabolism
6.3.1 Animal studies
The main pathway for the metabolism of acetaldehyde is shown in
Fig. 1.
6.3.1.1 Liver
The main pathway for the metabolism of acetaldehyde is by rapid
oxidation to acetate, which enters the citric acid cycle in an
activated form as acetyl-CoA and is metabolized to CO2 and H2O.
Although catalase and other oxidases may contribute to metabolism
(Brien & Loomis, 1983), because of its high affinity, at least 90% of
acetaldehyde is oxidized by mitochondrial ALDH (Hellström-Lindahl &
Weiner, 1985) reducing NAD+ to NADH in the process. This step can
be blocked by disulfiram.
There are multiple molecular forms of ALDH with different kinetic
properties that influence the rate of removal of acetaldehyde
(Marjanen, 1973; Parilla et al., 1974; Teschke et al., 1977).
Acetaldehyde is a highly reactive molecule that can react with
many other large or small molecules by adduction, condensation, or
polymerization. These pathways may have little quantitative
significance in acetaldehyde metabolism, but the by-products may have
biological significance (Collins et al., 1979; Sorrell & Tuma, 1985).
Acetaldehyde is the primary metabolic product of ethanol
oxidation. Since ethanol is oxidized to acetaldehyde mole for mole,
and, since the exposure to exogenous acetaldehyde is small, endogenous
acetaldehyde resulting from the metabolism of ingested ethanol is
likely to be the most important source of exposure for most people.
Oxidation of ethanol to acetaldehyde occurs predominantly under the
influence of ADH, of which there are many isoenzymic forms. Like
ALDH, ADH is also NAD dependent. The inseparable metabolism of
ethanol and acetaldehyde results in the reduction of NAD+, thus,
affecting the redox state of the liver causing secondary metabolic
consequences.
6.3.1.2 Respiratory tract
ALDH localization in the respiratory tract of Fischer-344 rats
was studied by Bogdanffy et al. (1986). Histochemical studies
indicated activity principally in the nasal respiratory epithelium,
especially in the supranuclear cytoplasm of ciliated epithelial cells.
Activity was also high in the Clara cells of the lower bronchioles.
The tracheal epithelia possessed only low levels of ALDH. The
olfactory epithelium was almost devoid of ALDH activity.
Casanova-Schmitz et al. (1984) characterized at least 2
isoenzymes of ALDH in rat nasal mucosa homogenates.
6.3.1.3 Kidneys
In an in vitro study, Michoudet & Baverel (1987a,b) studied the
metabolism of acetaldehyde in isolated dog, rat, guinea-pig, and
baboon kidney-cortex tubules.
Acetaldehyde was found to be metabolized by the tubules at high
rates and in a dose-dependent manner in all four species. It was
noted that, at all acetaldehyde concentrations, most of the
acetaldehyde removed was recovered as acetate in dog, guinea-pig, and
baboon, but not in rat kidney tubules.
6.3.1.4 Testes and ovaries
There are no studies on the capacity of the testes or ovaries to
mediate the biotransformation of acetaldehyde. However, ALDH activity
has been identified in the testes of Swiss-Webster mice (Anderson et
al., 1985).
6.3.1.5 Embryonic tissue
In an in vitro study, the ability of CBA/beige mouse (10 days
old) and Wistar rat (12 days old) embryos to metabolize acetaldehyde
was reported by Priscott & Ford (1985).
6.3.1.6 Metabolism during pregnancy
After intravenous administration of acetaldehyde (10 mg/kg body
weight) blood acetaldehyde levels were higher in pregnant rats than in
virgin rats. Acetaldehyde at high concentrations was able to cross
the placental barrier very rapidly. At low maternal concentrations,
it was metabolized by aldehyde dehydrogenase activity in the placenta
and fetal liver, and acetaldehyde was not detected in fetal blood.
Above the acetaldehyde threshold, the metabolic capacity of the
feto-placental unit was surpassed and acetaldehyde was detected in
fetal blood (Zorzano & Herrera, 1989).
6.3.2 Human studies
No high quality studies of the in vivo metabolism of
acetaldehyde in humans have been identified. Accurate assays for
acetaldehyde in blood and tissues have only recently become available
(Harade et al., 1978a,b).
Human liver ALDH consists of at least 4 main isoenzymes, which
are also present in many other tissues (Koivula, 1975; Goedde et al.,
1979). Mitochondrial ALDH is inactive in at least 40% of the Oriental
population. The frequently observed intolerance to alcohol (the
"flushing" reaction) is linked to this deficiency, which is produced
by an inherited positive mutation in the corresponding gene (Yoshida
et al., 1984; Goedde & Agarwal, 1986, 1987; Hsu et al., 1988).
Subjects with phenotypic deficiency have always shown the presence of
at least one mutant gene (heterozygous or homozygous) (Crabb et al.,
1989; Goedde et al., 1989; Singh et al., 1989).
In vitro, acetaldehyde (0.04-0.88 g/litre) was metabolized at
high rates and in a dose-dependent manner in isolated human
kidney-cortex tubules (Michoudet & Baverel, 1987b).
6.4 Elimination
In dogs, urinary excretion of acetaldehyde was essentially
non-existent following administration of a single dose of 600 mg
acetaldehyde/kg body weight, via a stomach tube (Booze & Oehme, 1986).
6.5 Reaction with cellular macromolecules
6.5.1 Proteins
Acetaldehyde can react with nucleophilic groups, such as amino,
hydroxyl, and sulfydryl groups, through nucleophilic attack on the
carbonyl carbon atom of acetaldehyde to give both stable and unstable
adducts (Tuma et al., 1984). Several adduct structures, formed when
acetaldehyde reacts with proteins in vitro, have been identified,
but have not yet been described fully.
The best characterized nucleophiles able to form adducts with
acetaldehyde are amino groups, notably the alpha-amino terminus of
peptides and proteins and the epsilon-amino group on the side-chain of
lysine residues. These reactions are shown in Fig. 2.
The structure of 2-methylimidazolidin-4-one adducts has been
confirmed by proton NMR (Gidley et al., 1981) and 13C-NMR
spectroscopy (San George & Hoberman, 1986). N-ethylation lysine
residues have been demonstrated by Tuma et al. (1984).
In a series of studies on lysine-dependent enzymes, Mauch et al.
(1986, 1987) were able to demonstrate that incubation of purified
enzymes with acetaldehyde for 1 h at 37°C led to inhibition of their
catalytic activity. Lysine non-dependent enzymes were not affected by
this treatment. A similar study involving the incubation of rat liver
histone H1 with physiological concentrations of acetaldehyde showed
that spontaneously stable adducts were formed on lysine residues at
the carboxy terminus, a site crucial for its function as a eukaryotic
repressor (Niëmela et al., 1990). This acetaldehyde-modified histone
H1 showed impaired DNA binding activity.
Tuma et al. (1987) have characterized the interaction of
acetaldehyde with a "highly reactive" lysine residue in purified
alpha-tubulin, which is only available in the monomeric form. They
found that modification of this residue was the critical factor in the
inhibition of tubulin polymerization by acetaldehyde, and that
modification of 5% of these residues was enough to inhibit tubulin
polymerization completely in vitro. Crebelli et al. (1989) found
similar effects: 0.075% v/v (13.5 mmol/litre) acetaldehyde partially
inhibited the in vitro polymerization of cattle brain tubulin, and
0.15% (27 mmol/litre) caused complete inhibition.
Incubation of calf brain microtubular proteins also resulted in
decreased polymerization, in an analogous manner to tubulin (McKinnon
et al., 1987a,b). Thus, acetaldehyde modification can impair the
molecular function of macromolecules, which can lead to marked
alterations in biological function.
No data are available on the formation of acetaldehydemodified
proteins in animals or humans directly exposed to acetaldehyde.
However, some data are available on proteins modified by acetaldehyde
derived from ethanol metabolism. In these studies, proteins carrying
acetaldehyde adducts were shown to be present in the liver cytosol of
rats fed ethanol for periods of 3 weeks, 12 months, or 27 months
(Worrall et al., 1991a). Acetaldehyde-modified proteins have also been
detected in the plasma (Liu et al., 1990) and haemoglobin of
alcoholics (Niemela & Israel, 1992). Furthermore, a limited
immunohistochemical study has demonstrated the presence of
acetaldehyde-modified proteins in the livers of some alcoholics
(Niemela et al., 1991). These studies demonstrate that acetaldehyde
adducts can form in the body. The possible immunological consequences
of adduct formation will be discussed in section 8.9.
6.5.2 Nucleic acids
No data are available from in vivo studies on the generation of
DNA adducts.
Acetaldehyde reacts with nucleosides and deoxynucleosides at pH
6.5 and 37°C in vitro to form unstable adducts by binding to the
exocyclic amino groups of adenine, cytosine, and guanine (Hemminki &
Suni, 1984). Addition of a reducing agent (sodium borohydride) leads
to the formation of stable adducts, of which the main one was
identified as N2-ethylguanosine using mass spectrometry and NMR.
Similar data for the formation of unstable adducts formed by reacting
acetaldehyde with ribonucleosides and deoxyribonucleosides was
reported by Fraenkel-Conrat & Singer (1988). When ethanol was present
in the reaction mixture, a different type of adduct was formed, which
was identified by fast atom bombardment and proton NMR to be a mixed
acetal (-NH-CH(CH3)-OR). These adducts were found to have half-lives
varying from 2.5 to 24 h at pH 7.5 and 37°C, depending on the base
involved.
7. EFFECTS ON ORGANISMS IN THE ENVIRONMENT
7.1 Aquatic organisms
An LC50 (semi-static study) of 35 mg/litre was found for
acetaldehyde in the guppy (Poecilia reticulata; 10 laboratory-reared
and acclimatized fish, 2-3 months old) (Deneer et al., 1988). Grahl
(1983) reported an LC50 (48-96 h) of 124 mg/litre for acetaldehyde
in fish (no additional information was provided). Juhnke & Luedemann
(1978) presented the results for fish obtained in the Golden Orfe
test, and found an LC50 of 140-124 mg/litre for acetaldehyde (no
additional information was provided). An LC50 (static conditions;
96-h) of 53 mg/litre was reported for the bluegill (Lepomis
macrochirus) by Von Burg & Stout (1991).
Acetaldehyde had a depressing effect on the aggressive behaviour
of the fish cichlid (Cichlasoma nigro fasciatum) at concentrations
that did not cause locomotor decrements in this species (Peeke &
Figler, 1981).
An EC5 (48-h; population growth) of 82 mg/litre and an EC50
(48-h; static conditions; immobilization) of 42 mg/litre were reported
for protozoa (Chilomonas paramecium) and the waterflea (Daphnia
magna), respectively (Von Burg & Stout, 1991).
7.2 Terrestrial organisms
Aharoni & Barkai-Golan (1973) studied the effects of acetaldehyde
vapours on the germination and colony-forming potential of two fungi
species, Alternaria tenuis and Stemphylium botryosum. The rate of
growth inhibition increased with both concentration and time of
exposure. The exposure of the spores was conducted at room
temperature. A. tenuis, the more sensitive species, was inactivated
by 0.54 µg acetaldehyde/m3 applied for 5 h, whereas 1.08 µg
acetaldehyde/m3 for 2 h, was needed to inactivate S. botryosum
spores.
Pittevils et al. (1979) reported the activity of acetaldehyde
against the fungi affecting stored apples and pears (Colletotrichum
gloeosporioides, Cryptosporiopsis malicorticis, Phlyctaena
vagabunda, Botrytis cinerea, and Alternaria tenuis). Acetaldehyde
was rapidly lethal at low concentrations: after a 24-h treatment
period, the lethal concentration of acetaldehyde ranged from
0.036 µg/m3 (A. tenuis) to 0.09 µg/m3 (C. gloeosporioides).
Acetaldehyde remained lethal for the five fungi, even when the
treatment lasted only 20 min (0.90 µg/m3 for P. vagabunda, C.
malicorticis, and A. tenuis, and 0.36 µg/m3 for C. gloeosporioides
and B. cinerea).
The fungi Botrytis cinerea, Penicillium expansum, Rhizopus
stolonifer, Monilinia fructicola, Erwinia carotovora, and
Pseudomonas fluorescens were killed, when exposed to
acetaldehyde vapours at concentrations ranging from 0.045 to
3.6 µg/m3, applied for 0.5 to 120 min at room temperature (Aharoni &
Stadelbacher, 1973).
Aharoni et al. (1979) studied acetaldehyde as a fumigant for
control of the green peach aphid (Myzus persicae) on head lettuce
(Lactuca sativa). When aphids were placed on the lettuce prior to
fumigation, 0.36 µg acetaldehyde/m3 and a 3-4 h exposure were
required for 100% mortality. A similar treatment (0.27-0.36 µg/m3
for 4 h) was found to cause 100% mortality of aphids on lettuce by
Stewart et al. (1980).
The fumigant effect of acetaldehyde was tested on the garden slug
(Arion hortensis; weight range, 0.2-0.5 g) and the grey field slug
(Agriolimax reticulatus; weight range, 0.3-0.6 g). It caused both
species to close the pulmonary aperture and to secrete excess
'irritation' mucus. Medial lethal values of 7.69 ± 0.21 mg/litre
per h for A. reticulatus and of 8.91 ± 0.81 mg/litre per h for
A. hortensis were found (Henderson, 1970).
The seed germination of the onion (Allium cepa L.), carrot
(Daucus carota L.), Palmer Amaranth (Amaranthus palmeri S Wats.),
and tomato (Lycopersicon esculentum Mill.) after exposure to
acetaldehyde (up to 1.52 mg/litre), was examined by Bradow & Connick
(1988). After a 3-day exposure, acetaldehyde inhibited the seed
germination of all four plants by more than 50%. Seeds inhibited by a
3-day exposure to acetaldehyde followed by a 4-day recovery period
germinated to the same extent as the controls after seven days, except
for the Palmer Amaranth, which remained inhibited.
Acetaldehyde at concentrations of 0.54-1.08 µg/m3 affected head
lettuce (Lactuca sativa), as evidenced by dark-green, water-soaked,
necrotic areas on the outer leaves of the lettuce. Concentrations of
up to 0.36 µg/m3 did not affect the lettuce (Aharoni et al., 1979;
Stewart et al., 1980).
8. EFFECTS ON EXPERIMENTAL ANIMALS AND IN VITRO TEST SYSTEMS
8.1 Single exposure
8.1.1 LD50 and LC50 values
Relevant data are summarized in Table 4.
Oral LD50s for acetaldehyde in rats and mice ranged from 660 to
1930 mg/kg body weight. LC50s (0.5-4 h) in rats and Syrian hamsters
ranged from 24 to 37 g/m3. It is, therefore, concluded that the
acute toxicity of acetaldehyde is low. LD50 values by the dermal
route were not available.
LD50s for intratracheal, subcutaneous, intraperitoneal, and
intravenous administration are also presented in Table 4.
Table 4. LD50/LC50 values for acetaldehyde
Species Route of LD50/LC50 Reference
administration
Rat oral 1930 mg/kg body weight Smyth et al. (1951)
Rat oral 660 mg/kg body weight Sprince et al. (1974)
Mouse oral 1230 mg/kg body weight US NRC (1977)
Dog oral > 600 mg/kg body weight Booze & Oehme (1986)
Rat inhalation 24 g/m3; 4 h Appelman et al. (1982)
Rat inhalation 37 g/m3; 0.5 h Skog (1950)
Syrian
hamster inhalation 31 g/m3; 4 h Kruysse (1970)
Syrian
hamster intratracheal 96.1 mg/kg body weight Feron & De Jong (1971)
Rat subcutaneous 640 mg/kg body weight Skog (1950)
Mouse subcutaneous 560 mg/kg body weight Skog (1950)
Mouse intraperitoneal 500 mg/kg body weight Truitt & Walsh (1971)
Mouse intravenous 165 mg/kg body weight O'Shea & Kaufman
(pregnant) (1979)
8.2 Short-term exposure
8.2.1 Oral
Oral administration in the drinking-water of 675 mg
acetaldehyde/kg body weight to Wistar rats, daily for 4 weeks,
resulted in slight to moderate focal hyperkeratosis of the forestomach
in 8/10 males and 8/10 females. No effects were observed at lower
dose levels of 25 and 125 mg/kg body weight. In the control group,
very slight focal hyperkeratosis of the forestomach was noted in 6/20
females and 3/20 males (1/20 slight). At the top dose (675 mg/kg),
relative kidney weights were slightly increased in males, urinary
production was decreased, and there were variations in serum
biochemistry, most of which were attributable to reduced water intake.
There were no effects in the liver. The no-observed-effect level
(NOEL) was 125 mg/kg, the lowest-observed-effect level (LOEL) was
675 mg/kg (Til et al., 1988).
8.2.2 Inhalation
Male Sprague-Dawley rats were continuously exposed to
acetaldehyde vapour for 22 days at levels gradually increasing from
750 mg/m3 for a few days up to 2500 mg/m3 for the last few days.
By gradually increasing the concentrations, mortality in the early
period following exposure to 2000-2500 mg/m3 was prevented,
presumably because of metabolic adaptation; sudden, high, blood
acetaldehyde levels inducing vagal reflex reactions may result in
respiratory inhibition, and, as a consequence, death (Lamboeuf et al.,
1987; Latge et al., 1987).
Groups of 10 male and 10 female Wistar rats were exposed to
acetaldehyde at 0, 720, 1800, 3950, or 9000 mg/m3 (0, 400, 1000,
2200, or 5000 ppm) for 6 h/day, 5 days/week, for 4 weeks. Mortality
was slightly increased at 3950 and 9000 mg/m3, whereas growth was
retarded at 1800 mg/m3 and above in males, and at 9000 mg/m3 in
females. At 9000 mg/m3, relative liver weight decreased, and
relative lung weight in males increased. No treatment-related
histopathological changes were observed in the liver. Degenerative
changes of the nose were observed after exposure to all concentrations
(720 mg/m3-9000 mg/m3), with hyperplasia and metaplasia occurring
at concentrations of 3950 mg/m3 or more. A NOEL was not identified
(LOEL: 720 mg/m3) (Appelman et al., 1982).
Groups of 10 male Wistar rats were exposed to acetaldehyde, for
6 h/day, 5 days/week, for 4 weeks, in three different patterns: (a)
as a continuous daily exposure of 6 h to 0, 270, or 900 mg/m3 (0,
150, or 500 ppm), (b) as two daily exposures of 3 h to similar
concentrations with an intervening 1.5-h period with no exposure, or
(c) as two daily 3-h periods of exposure to similar concentrations
with an intervening 1.5-h period with eight short (5-min) peaks of 6
times the basic concentration, resulting in time-weighted average
concentrations of 0, 255, or 1050 mg/m3, respectively. Though there
were no indications of toxicity following continuous or interrupted
exposures to 270 and 900 mg/m3 and intermittent high/low exposure to
255 mg/m3, intermittent high/low exposure to 1050 mg/m3 induced
growth retardation (Appelman et al., 1986).
At 900 mg/m3, the observed effects were very similar to the
ones reported earlier by Appelman et al. (1982) at 720 mg/m3.
Variation of the pattern of exposure, by including a 1.5-h break, or
by additionally including eight 5-min, 6-fold higher peak exposures,
did not alter the observed degenerative effects. No effects were
observed in Wistar rats exposed to a lower concentration, 5 days/week
for 4 weeks, either as a "continuous" (6 h/day) exposure of
270 mg/m3, or as a time-weighted average of 255 mg/m3 after the
described intermittent low-high exposure. The NOEL was 255 mg/m3,
6-h TWA (LOEL = 1050 mg/m3, 6-h time weighted average) (Appelman et
al., 1986).
In another study, groups of 12 male Wistar rats were exposed to 0
or 437 mg acetaldehyde/m3 (0 or 243 ppm), 8 h/day, 5 days per week,
for 5 weeks. Hyperplasia of the olfactory epithelium and nasal
inflammation were observed in exposed animals, and on the basis of
lung function tests, residual volume and functional residual capacity
were increased, indicating some (unspecified) damage of the distal
airways (Saldiva et al., 1985).
8.2.3 Dermal
No relevant studies were identified.
8.2.4 Parenteral
Effects in the liver have been reported in several studies, but
only at very high doses. Intraperitoneal injection of male albino
rats with 200 mg acetaldehyde/kg body weight, daily, for 10 days, with
additional pyrazole treatment to inhibit the conversion of
acetaldehyde to ethanol, caused fatty accumulation in the liver, as
indicated by accumulation of total lipids, triacyl glycerols, and
total cholesterol, increased glycogenolysis, and a shift in metabolism
from the citric acid cycle towards the pentose phosphate pathway in
the liver. Serum triacyl glycerol, total cholesterol, and free fatty
acid levels were also increased. Changes were similar in rats not
receiving pyrazole pretreatment (Prasanna & Ramakrishnan, 1984a,
1987). The same treatment altered thyroid function, as indicated by
lower serum T4 and decreased iodine uptake in male albino rats, though
these effects may have been secondary to the observed hepatic changes
(Prasanna et al., 1986) and histopathological changes of the pancreas,
with resulting changes in trypsinogen levels and amylase secretion and
activity in female Sprague-Dawley rats (Majumdar et al., 1986).
8.3 Skin and eye irritation; sensitization
No relevant data were identified.
8.4 Long-term exposure
8.4.1 Oral
In rats exposed to 0.05% acetaldehyde in drinking-water
(estimated by the Task Group to be approximately 40 mg/kg body weight)
for 6 months, there was an increase in collagen synthesis in the liver
(Bankowski et al., 1993). The toxicological significance of this
observation is not known; no other effects were examined.
8.4.2 Inhalation
Non-neoplastic effects observed in carcinogenicity studies are
discussed in section 8.7.1.
Groups of 20 Syrian hamsters were exposed to acetaldehyde vapour
at 0, 700, 2400, or 8200 mg/m3 (0, 390, 1340, or 4560 ppm) for
6 h/day, 5 days/week, for 13 weeks. Increased relative lung and heart
weights as well as growth retardation were reported after exposure to
8200 mg/m3, though there were no increases in mortality in any of
the exposed groups (Kruysse et al., 1975). At the highest
concentration, there were severe degenerative, hyperplastic, and
metaplastic changes in the epithelium as well as subepithelial glands
and turbinate bones. Rhinitis was observed, with abundant nasal
discharge and salivation. The epithelium of the larynx, trachea, and
lungs was damaged, with some focal hyperplasia and metaplasia,
accompanied by tracheitis and focal bronchopneumonia. Changes in the
tracheal epithelium were also observed at 2400 mg/m3. At
700 mg/m3, no significant effects were observed (NOEL: 700 mg/m3;
LOEL: 2400 mg/m3).
8.5 Reproductive and developmental toxicity
Studies on reproductive effects have not been identified. A
number of studies on developmental effects have been conducted,
primarily to investigate the role of acetaldehyde in ethanol-induced
teratogenicity. However, in all of these studies, acetaldehyde was
administered by injection rather than by the principal routes of
exposure in the occupational and general environments (i.e., ingestion
and inhalation). Results of identif