UNITED NATIONS ENVIRONMENT PROGRAMME
INTERNATIONAL LABOUR ORGANISATION
WORLD HEALTH ORGANIZATION
INTERNATIONAL PROGRAMME ON CHEMICAL SAFETY
ENVIRONMENTAL HEALTH CRITERIA 196
Methanol
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.
Environmental Health Criteria 196
First draft prepared by Dr. L. Fishbein, Fairfax, Virginia, USA
Published under the joint sponsorship of the United Nations
Environment Programme, the International Labour Organisation, and the
World Health Organization, and produced within the framework of the
Inter-Organization Programme for the Sound Management of Chemicals.
World Health Organization
Geneva, 1997
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of the biological action of chemicals.
WHO Library Cataloguing in Publication Data
Methanol.
(Environmental health criteria ; 196)
1.Alcohol, Methyl - toxicity 2.Alcohol, Methyl - adverse effects
3.Environmental exposure I.Series
ISBN 92 4 157196 9 (NLM Classification: QV 83)
ISSN 0250-863X
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CONTENTS
ENVIRONMENTAL HEALTH CRITERIA FOR METHANOL
PREAMBLE
ABBREVIATIONS
1. SUMMARY
1.1. Identity, physical and chemical properties, analytical
methods
1.2. Sources of human exposure
1.3. Environmental levels and human exposure
1.4. Environmental distribution and transformation
1.5. Absorption, distribution, biotransformation and elimination
1.6. Effects on laboratory mammals and in vitro test systems
1.6.1. Systemic toxicity
1.6.2. Genotoxicity and carcinogenicity
1.6.3. Reproductive toxicity, embryotoxicity and
teratogenicity
1.7. Effects on humans
1.8. Effects on organisms in the environment
2. IDENTITY, PHYSICAL AND CHEMICAL PROPERTIES, AND ANALYTICAL
METHODS
2.1. Identity
2.2. Physical and chemical properties
2.2.1. Physical properties
2.2.2. Chemical properties
2.3. Conversion factors
2.4. Analytical methods
2.4.1. Environmental samples
2.4.1.1 Methanol in air
2.4.1.2 Methanol in fuels
2.4.1.3 Methanol in fuel emissions
2.4.1.4 Methanol in sewage and aqueous solutions
2.4.1.5 Methanol in soils
2.4.2. Foods, beverages and consumer products
2.4.3. Biological materials
2.4.3.1 Methanol in exhaled air
2.4.3.2 Methanol in blood
2.4.3.3 Methanol in urine
2.4.3.4 Methanol in miscellaneous biological
tissues
2.4.3.5 Methanol metabolites in biological
fluids
3. SOURCES OF HUMAN AND ENVIRONMENTAL EXPOSURE
3.1. Natural occurrence
3.2. Anthropogenic sources
3.2.1. Production levels and processes
3.2.1.1 Production processes
3.2.1.2 Production figures
3.2.2. Uses
3.2.2.1 Use as feedstock for chemical syntheses
3.2.2.2 Use as fuel
3.2.2.3 Other uses
3.2.2.4 Losses into the environment
4. ENVIRONMENTAL TRANSPORT, DISTRIBUTION AND TRANSFORMATION
4.1. Transport and distribution between media
4.2. Transformation
4.2.1. Biodegradation
4.2.1.1 Water and sewage sludge
4.2.1.2 Soils and sediments
4.2.2. Abiotic degradation
4.2.2.1 Water
4.2.2.2 Air
4.3.2. Bioconcentration
5. ENVIRONMENTAL LEVELS AND HUMAN EXPOSURE
5.1. Environmental levels
5.1.1. Air
5.1.2. Water
5.1.3. Food
5.1.4. Tobacco smoke
5.2. Occupational exposure
5.3. General population
6. KINETICS AND METABOLISM IN LABORATORY ANIMALS AND HUMANS
6.1. Absorption
6.1.1. Inhalation
6.1.2. Oral
6.1.3. Dermal
6.2. Distribution
6.3. Metabolic transformation
6.4. Elimination and excretion
6.5. Modelling of pharmacokinetic and toxicokinetic data
7. EFFECTS ON LABORATORY MAMMALS AND IN VITRO TEST SYSTEMS
7.1. Single exposure
7.1.1. Non-primates
7.1.2. Non-human primates
7.2. Short-term exposure
7.2.1. Inhalation exposure
7.3. Long-term exposure
7.4. Skin and eye irritation; sensitization
7.5. Reproduction toxicity, embryotoxicity and teratogenicity
7.5.1. Reproductive toxicity (effects on fertility)
7.5.2. Developmental toxicity
7.5.3. Behavioural effects
7.5.4. In vitro studies
7.6. Mutagenicity and related end-points
7.6.1. In vitro studies
7.6.2. In vivo studies
7.7. Carcinogenicity
7.8. Special studies
7.8.1. Effects on hepatocytes
7.8.2. Toxic interactions
7.8.3. Studies with exhaust emissions from methanol-
fuelled engines
7.9. Mechanism of ocular toxicity
8. EFFECTS ON HUMANS
8.1. General population and occupational exposure
8.1.1. Acute toxicity
8.1.2. Clinical features of acute poisonings
8.1.3. Repeated or chronic exposure
8.1.4. Reproductive and developmental effects
8.1.5. Chromosomal and mutagenic effects
8.1.6. Carcinogenic effects
8.1.7. Sensitive sub-populations
9. EFFECTS ON OTHER ORGANISMS IN THE LABORATORY AND FIELD
9.1. Aquatic organisms
9.1.1. Microorganisms
9.1.2. Algae
9.1.3. Aquatic invertebrates
9.1.4. Fish
9.2. Terrestrial organisms
9.2.1. Plants
10. EVALUATION OF EFFECTS ON HUMAN HEALTH AND THE ENVIRONMENT
10.1. Evaluation of human health risks
10.1.1. Exposure
10.1.2. Human health effects
10.1.3. Approaches to risk assessment
10.2. Evaluation of effects on the environment
11. RECOMMENDATIONS FOR PROTECTION OF HUMAN HEALTH AND THE
ENVIRONMENT
11.1. Protection of human health
11.2. Protection of the environment
12. FURTHER RESEARCH
13. PREVIOUS EVALUATIONS BY INTERNATIONAL BODIES
REFERENCES
RESUME
RESUMEN
NOTE TO READERS OF THE CRITERIA MONOGRAPHS
Every effort has been made to present information in the criteria
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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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Financial support for this Task Group meeting was provided by the
United Kingdom Department of Health as part of its contributions to
the IPCS.
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WHO TASK GROUP ON ENVIRONMENTAL HEALTH CRITERIA FOR METHANOL
Members
Dr D. Anderson, British Industry Biological Research Association
(BIBRA) Toxicology International, Carshalton, Surrey, United
Kingdom
Dr S.A. Assimon, Contaminants Standards Monitoring and Projects
Branch, US Food and Drug Administration, Washington DC, USA
Dr H.B.S. Conacher, Bureau of Chemical Safety, Ottawa, Ontario,
Canada
Professor J. Eells, Department of Pharmacology and Toxicology,
Medical College of Wisconsin Milwaukee, USA (Chairman)
Mr J. Fawell, National Centre for Environmental Toxicology,
Marlow, Essex, United Kingdom
Dr L. Fishbein, Fairfax, Virginia, USA (Joint Rapporteur)
Dr K. McMartin, Department of Pharmacology and Therapeutics,
Louisiana State University Medical Center, Shreveport,
Louisiana, USA
Mr H. Malcolm, Institute of Terrestrial Ecology, Monks Wood,
Huntingdon, United Kingdom (Joint Rapporteur)
Dr H.B. Matthews, National Institute of Environmental Health
Sciences, Research Triangle Park, North Carolina, USA
Professor M. Piscator, Karolinska Institute, Stockholm, Sweden
(Vice-Chairman)
Dr G. Rosner, Merzhausen, Germany
Representatives of other Organizations
Professor K.R. Butterworth, BIBRA Toxicology International,
Carshalton, Surrey, United Kingdom (representing the
International Union of Toxicology)
Mr M.G. Penman, ICI Chemicals & Polymers Limited,
Middlesbrough, Cleveland, United Kingdom (representing the
European Centre for Ecotoxicology and Toxicology of
Chemicals)
Secretariat
Dr E. Smith, International Programme on Chemical Safety, World
Health Organization, Geneva, Switzerland (Secretary)
Mr J.D. Wilbourn, Unit of Carcinogen Identification and
Evaluation, International Agency for Research on Cancer
(IARC), Lyon, France
ENVIRONMENTAL HEALTH CRITERIA FOR METHANOL
A WHO Task Group on Environmental Health Criteria for Methanol
met at the British Industrial Biological Research Association (BIBRA)
Toxicology International, Carshalton, Surrey, United Kingdom from 28
to 31 October 1996. Dr D. Anderson opened the meeting and welcomed
the participants on behalf of the host institute. Dr E. Smith, IPCS,
welcomed the participants on behalf of the Director, IPCS, and the
three IPCS cooperating organizations (UNEP/ILO/WHO). The Task Group
reviewed and revised the draft criteria monograph and made an
evaluation of the risks for human health and the environment from
exposure to methanol.
Dr L. Fishbein, Fairfax, Virginia, USA prepared the first draft
of this monograph. The second draft, incorporating comments received
following the circulation of the first draft to the IPCS Contact
Points for Environmental Health Criteria monographs, was also prepared
by Dr Fishbein.
Dr E.M. Smith and Dr P.G. Jenkins, both of the IPCS Central Unit,
were responsible for the overall scientific content and technical
editing, respectively.
The efforts of all who helped in the preparation and finalization
of the monograph are gratefully acknowledged.
ABBREVIATIONS
ATP adenosine triphosphate
BCF bioconcentration factor
BOD biochemical oxygen demand
COD chemical oxygen demand
CNS central nervous system
FID flame ionization detection
GC gas chromatography
MLD minimum lethal dose
MS mass spectrometry
MTBE methyl tertiary butyl ether
NAD nicotinamide adenine dinucleotide
NCAM neural cell adhesion molecule
NOAEL no-observed-adverse-effect level
THF tetrahydrofolate
TLV threshold limit value
1. SUMMARY
1.1 Identity, physical and chemical properties, analytical methods
Methanol is a clear, colourless, volatile flammable liquid with a
mild alcoholic odour when pure. It is miscible with water and many
organic solvents and forms many binary azeotropic mixtures.
Analytical methods, principally gas chromatography (GC) with
flame ionization detection (FID), are available for the determination
of methanol in various environmental media (air, water, soil and
sediments) and foods, as well as the determination of methanol and its
principal metabolite, formate, in body fluids and tissues. In addition
to GC-FID, enzymatic procedures with colorimetric end-points are
utilized for the determination of formate in blood, urine and tissues.
Determination of methanol in the workplace usually involves
collection and concentration on silica gel, followed by aqueous
extraction and GC-FID or GC-mass spectrometry analysis of the extract.
1.2 Sources of human exposure
Methanol occurs naturally in humans, animals and plants. It is a
natural constituent in blood, urine, saliva and expired air. A mean
urinary methanol level of 0.73 mg/litre (range 0.3-2.61 mg/litre) in
unexposed individuals and a range of 0.06 to 0.32 µg/litre in expired
air have been reported.
The two most important sources of background body burdens for
methanol and formate are diet and metabolic processes. Methanol is
available in the diet principally from fresh fruits and vegetables,
fruit juices (average 140 mg/litre, range 12 to 640 mg/litre),
fermented beverages (up to 1.5 g/litre) and diet foods (principally
soft drinks). The artificial sweetener aspartame is widely used and,
on hydrolysis, 10% (by weight) of the molecule is converted to free
methanol, which is available for absorption.
About 20 million tonnes of methanol were produced worldwide in
1991, principally by catalytic conversion of pressurized synthesis gas
(hydrogen, carbon dioxide and carbon monoxide). Worldwide capacity was
projected to rise to 30 million tonnes by 1995.
Methanol is used in the industrial production of many important
organic compounds, principally methyl tertiary butyl ether (MTBE),
formaldehyde, acetic acid, glycol methyl ethers, methylamine, methyl
halides and methyl methacrylate.
Methanol is a constituent of a large number of commercially
available solvents and consumer products including paints, shellacs,
varnishes, paint thinners, cleansing solutions, antifreeze solutions,
automotive windshield washer fluids and deicers, duplicating fluids,
denaturant for ethanol, and in hobby and craft adhesives. Potentially
large uses of methanol are in its direct use as a fuel, in gasoline
blends or as a gasoline extender. It should be noted that the highest
morbidity and mortality has been associated with deliberate or
accidental oral ingestion of methanol-containing mixtures.
Methanol has been identified in exhausts from both gasoline and
diesel engines and in tobacco smoke.
1.3 Environmental levels and human exposure
Emissions of methanol primarily occur from the miscellaneous
industrial and domestic solvent use, methanol production, end-product
manufacturing and bulk storage and handling losses.
Exposures to methanol can occur in occupational settings through
inhalation or dermal contact. Many national occupational health
exposure limits suggest that workers are protected from any adverse
effects if exposures do not exceed a time-weighted average of
260 mg/m3 (200 ppm) methanol for any 8-h day and for a 40-h working
week.
Current general population exposures through air are typically
10 000 times lower than occupational limits. The general population is
exposed to methanol in air at concentrations ranging from less than
0.001 mg/m3 (0.8 ppb) in rural air to nearly 0.04 mg/m3 (30 ppb) in
urban air.
Data on the occurrence of methanol in finished drinking-water is
limited, but methanol is frequently found in industrial effluents.
If the projected use of methanol as an alternate fuel or in
admixture with fuels increases significantly, it can be expected that
there will be widespread exposure to methanol via inhalation of
vapours from methanol-fuelled vehicles and/or siphoning or
percutaneous absorption of methanol fuels or blends.
1.4 Environmental distribution and transformation
Methanol is readily degraded in the environment by photo
oxidation and biodegradation processes. Half-lives of 7-18 days have
been reported for the atmospheric reaction of methanol with hydroxyl
radicals.
Many genera and strains of microorganisms are capable of using
methanol as a growth substrate. Methanol is readily degradable under
both aerobic and anaerobic conditions in a wide variety of
environmental media including fresh and salt water, sediments and
soils, ground water, aquifer material and industrial wastewater; 70%
of methanol in sewage systems is generally degraded within 5 days.
Methanol is a normal growth substrate for many soil
microorganisms, which are capable of completely degrading methanol to
carbon dioxide and water.
Methanol has a fairly low absorptive capacity on soils.
Bioconcentration in most organisms is low.
Methanol is of low toxicity to aquatic and terrestrial organisms,
and effects due to environmental exposure to methanol are unlikely to
be observed except in the case of a spill.
1.5 Absorption, distribution, biotransformation and elimination
Methanol is readily absorbed by inhalation, ingestion and dermal
exposure, and it is rapidly distributed to tissues according to the
distribution of body water. A small amount of methanol is excreted
unchanged by the lungs and kidneys.
Following ingestion, peak serum levels occur within 30-90 min,
and methanol is distributed throughout the body with a volume of
distribution of approximately 0.6 litre/kg.
Methanol is metabolized primarily in the liver by sequential
oxidative steps to formaldehyde, formic acid and carbon dioxide. The
initial step involves oxidation to formaldehyde by hepatic alcohol
dehydrogenase, which is a saturable rate-limiting process. The
relative affinity of alcohol dehydrogenase for ethanol and methanol is
approximately 20:1. In step 2, formaldehyde is oxidized by
formaldehyde dehydrogenase to formic acid/or formate depending on the
pH. In step 3, formic acid is detoxified to carbon dioxide by folate-
dependent reactions.
Elimination of methanol from the blood via the urine and exhaled
air and by metabolism appears to be slow in all species, especially
when compared to ethanol. Clearance proceeds with reported half-times
of 24 h or more with doses greater than 1 g/kg and half-times of
2.5-3 h for doses less than 0.1 g/kg. It is the rate of metabolic
detoxification, or removal of formate that is vastly different between
rodents and primates and is the basis for the dramatic differences in
methanol toxicity observed between rodents and primates.
1.6 Effects on laboratory mammals and in vitro test systems
1.6.1 Systemic toxicity
The acute and short-term toxicity of methanol varies greatly
between different species, toxicity being highest in species with a
relatively poor ability to metabolize formate. In such cases of poor
metabolism of formate, fatal methanol poisoning occurs as a result of
metabolic acidosis and neuronal toxicity, whereas, in animals that
readily metabolize formate, consequences of CNS depression (coma,
respiratory failure, etc.) are usually the cause of death. Sensitive
primate species (humans and monkeys) develop increased blood formate
concentrations following methanol exposure, while resistant rodents,
rabbits and dogs do not. Humans and non-human primates are uniquely
sensitive to the toxic effects of methanol. Overall methanol has a low
acute toxicity to non-primate animals. The LD50 values and minimal
lethal doses after oral exposure range from 7000 to 13 000 mg/kg in
the rat, mouse, rabbit and dog and from 2000 to 7000 mg/kg for the
monkey.
Rats exposed to levels of methanol up to 6500 mg/m3 (5000 ppm)
for 6 h/day, 5 days/week for 4 weeks, exhibited no exposure-related
effects except for increased discharges around the nose and eyes.
These were considered reflective of upper respiratory irritation.
Rats exposed to methanol vapour levels up to 13 000 mg/m3
(10 000 ppm) for 6 h/day, 5 days/week for 6 weeks, failed to
demonstrate pulmonary toxicity.
In the rabbit, methanol is a moderately irritant to the eye. It
was not skin-sensitizing in a modified maximization test.
Toxic effects found in methanol-exposed primates include
metabolic acidosis and ocular toxicity, effects that are not normally
found in folate-sufficient rodents. The differences in toxicity are
due to differences in the rate of metabolism of the methanol
metabolite formate. For instance, the clearance of formate from the
blood of exposed primates is at least 50% slower than for rodents.
Monkeys receiving methanol doses higher than 3000 mg/kg by gavage
demonstrated ataxia, weakness and lethargy within a few hours of
exposure. These signs tended to disappear within 24 h and were
followed by transient coma in some of the animals.
In monkeys exposed to methanol for 6 h/day for 5 days a week, 20
repeated exposures to 6500 mg/m3 (5000 ppm) methanol failed to elicit
ocular effects.
1.6.2 Genotoxicity and carcinogenicity
Methanol has given negative results for gene mutation in bacteria
and yeast assays, but it did induce chromosomal malsegregation in
Aspergillus. It did not induce sister chromatic exchanges in Chinese
hamster cells in vitro but caused significant increases in mutation
frequencies in L5178Y mouse lymphoma cells.
Methanol inhalation did not induce chromosomal damage in mice.
There is some evidence that oral or intraperitoneal administration
increased the incidence of chromosomal damage in mice.
There is no evidence from animal studies to suggest that methanol
is a carcinogen, although the lack of an appropriate animal model is
recognized.
1.6.3 Reproductive toxicity, embryotoxicity and teratogenicity
Conflicting results have been reported on the effects of
inhalation of methanol for up to six weeks on gonadotropin and
testosterone concentrations.
The inhalation of methanol by pregnant rodents throughout the
period of embryogenesis induces a wide range of concentration-
dependent teratogenic and embryolethal effects. Treatment-related
malformations, predominantly extra or rudimentary cervical ribs and
urinary or cardiovascular defects, were found in fetuses of rats
exposed 7 h/day for 7-15 days of gestation to 26 000 mg/m3
(20 000 ppm) methanol. Slight maternal toxicity was found at this
exposure level, and no adverse effects to the mother or offspring were
found in animals exposed to 6500 mg/m3 (5000 ppm), which was
interpreted as the no-observed-adverse-effect level (NOAEL) for this
test system.
Increased incidences of exencephaly and cleft palate were found
in the offspring of CD-1 mice exposed 7 h/day, on days 6-15 of
gestation, to methanol levels of 6500 mg/m3 (5000 ppm) or more. There
was increased embryo/fetal death at 9825 mg/m3 (7500 ppm) or more and
an increasing incidence of full-litter resorptions. Reduced fetal
weight was observed at 13 000 and 19 500 mg/m3 (10 000 or 15 000
ppm). The NOAEL for developmental toxicity was 1300 mg/m3 (1000 ppm)
methanol. There was no evidence of maternal toxicity at methanol
exposure levels below 9000 mg/m3 (7000 ppm).
When litters of pregnant CD-1 mice were given 4 g methanol/kg by
gavage, the incidences of adverse effects on resorption, external
defects including cleft palate, and fetal weight were similar to those
found in the 13 000 mg/m3 (10 000 ppm) inhalation exposure group,
presumably due to the greater rate of respiration of the mouse. The
mouse is more sensitive than the rat to developmental toxicity
resulting from inhaled methanol.
Transient neurological signs and reduced body weights were found
in CD-1 dams exposed to 19 500 mg/m3 (15 000 ppm) for 6 h/day
throughout organogenesis (gestational days 6-15). Fetal malformations
found at 13 000 and 19 500 mg/m3 (10 000 and 15 000 ppm) included
neural and ocular defects, cleft palate, hydronephrosis and limb
anomalies.
1.7 Effects on humans
Humans (and non-human primates) are uniquely sensitive to
methanol poisoning and the toxic effects in these species is
characterized by formic acidaemia, metabolic acidosis, ocular
toxicity, nervous system depression, blindness, coma and death. Nearly
all of the available information on methanol toxicity in humans
relates to the consequences of acute rather than chronic exposures. A
vast majority of poisonings involving methanol have occurred from
drinking adulterated beverages and from methanol-containing products.
Although ingestion dominates as the most frequent route of poisoning,
inhalation of high concentrations of methanol vapour and percutaneous
absorption of methanolic liquids are as effective as the oral route in
producing acute toxic effects. The most noted health consequence of
longer-term exposure to lower levels of methanol is a broad range of
ocular effects.
The toxic properties of methanol are based on factors that govern
both the conversion of methanol to formic acid and the subsequent
metabolism of formate to carbon dioxide in the folate pathway. The
toxicity is manifest if formate generation continues at a rate that
exceeds its rate of metabolism.
The lethal dose of methanol for humans is not known for certain.
The minimum lethal dose of methanol in the absence of medical
treatment is between 0.3 and 1 g/kg. The minimum dose causing
permanent visual defects is unknown.
The severity of the metabolic acidosis is variable and may not
correlate well with the amount of methanol ingested. The wide
interindividual variability of the toxic dose is a prominent feature
in acute methanol poisoning.
Two important determinants of human susceptibility to methanol
toxicity appear to be (1) concurrent ingestion of ethanol, which slows
the entrance of methanol into the metabolic pathway, and (2) hepatic
folate status, which governs the rate of formate detoxification.
The symptoms and signs of methanol poisoning, which may not
appear until after an asymptomatic period of about 12 to 24 h, include
visual disturbances, nausea, abdominal and muscle pain, dizziness,
weakness and disturbances of consciousness ranging from coma to clonic
seizures. Visual disturbances generally develop between 12 and 48 h
after methanol ingestion and range from mild photophobia and misty or
blurred vision to markedly reduced visual acuity and complete
blindness. In extreme cases death results. The principal clinical
feature is severe metabolic acidosis of the anion-gap type. The
acidosis is largely attributed to the formic acid produced when
methanol is metabolized.
The normal blood concentration of methanol from endogenous
sources is less than 0.5 mg/litre (0.02 mmol/litre), but dietary
sources may increase blood methanol levels. Generally, CNS effects
appear above blood methanol levels of 200 mg/litre (6 mmol/litre);
ocular symptoms appear above 500 mg/litre (16 mmol/litre), and
fatalities have occurred in untreated patients with initial methanol
levels in the range of 1500-2000 mg/litre (47-62 mmol/litre).
Acute inhalation of methanol vapour concentrations below
260 mg/m3 or ingestion of up to 20 mg methanol/kg by healthy or
moderately folate-deficient humans should not result in formate
accumulation above endogenous levels.
Visual disturbances of several types (blurring, constriction of
the visible field, changes in colour perception, and temporary or
permanent blindness) have been reported in workers who experienced
methanol air levels of about 1500 mg/m3 (1200 ppm) or more.
A widely used occupational exposure limit for methanol is
260 mg/m3 (200 ppm), which is designed to protect workers from any of
the effects of methanol-induced formic acid metabolic acidosis and
ocular and nervous system toxicity.
No other adverse effects of methanol have been reported in
humans except minor skin and eye irritation at exposures well above
260 mg/m3 (200 ppm).
1.8 Effects on organisms in the environment
LC50 values in aquatic organisms range from 1300 to
15 900 mg/litre for invertebrates (48-h and 96-h exposures), and
13 000 to 29 000 mg/litre for fish (96-h exposure).
Methanol is of low toxicity to aquatic organisms, and effects due
to environmental exposure to methanol are unlikely to be observed,
except in the case of a spill.
2. IDENTITY, PHYSICAL AND CHEMICAL PROPERTIES, ANALYTICAL METHODS
2.1 Identity
Chemical formula: CH3OH
Chemical structure: H
'
H - C - OH
'
H
Relative molecular mass: 32.04
CAS chemical name: methanol
CAS registry number: 67-56-1
RTECS number: PC 1400000
Synonyms: methyl alcohol, carbinol, wood
alcohol, wood spirits, wood
naphtha, Columbian spirits,
Manhattan spirits, colonial spirit,
hydroxymethane, methylol,
methylhydroxide,
monohydroxymethane, pyroxylic
spirit
Impurities in commercial methanol include acetone, acetaldehyde,
acetic acid and water.
2.2 Physical and chemical properties
2.2.1 Physical properties
Methanol is a colourless, volatile, flammable liquid with a mild
alcoholic odour when pure. However, the crude product may have a
repulsive pungent odour. Methanol is miscible with water, alcohols,
esters, ketones and most other solvents and forms many azeotropic
mixtures. It is only slightly soluble in fats and oils (Clayton &
Clayton, 1982; Windholz, 1983; Elvers et al., 1990).
Important physical constants and properties of methanol are
summarized in Table 1.
Table 1. Some physical properties of methanola
Appearance clear colourless liquid
Odour slight alcoholic when pure;
crude material pungent
Boiling point 64.7°C
Flash point 15.6°C (open cup)
12.2°C (closed cup)
Freezing point -97.68°C
Specific gravity 0.7915 (20/4°C)
0.7866 (25°C)
Vapour pressure
at 30°C 160 mmHg
at 20°C 92 mmHg
Henry's Law Constant (25°C) 1.35 x 10-4atm.m3/mole
Log P (octanol/water) -0.82; -0.77; -0.68
Partition constant -0.66; -0.64
Ignition temperature 470°C
Explosive limits in air lower 5.5
(% by volume) upper 44
Refractive index n20 1.3284
a Data from: Clayton & Clayton, 1982; Elvers et al., 1990;
Grayson, 1981; Howard, 1990; Windholz, 1983.
In the USA, sales grade methanol must normally meet the
following specifications:
methanol content (weight %) minimum 99.85
acetone and aldehydes (ppm) maximum 30
acid (as acetic acid) (ppm) maximum 30
water content (ppm) maximum 1.500
specific gravity (d2020) 0.7928
permanganate time, minimum 30
odour characteristic
distillation range at 101 kPa 1°C, must include
64.6°C
colour, platinum-cobalt scale, maximum 5
appearance clear-colourless
residual on evaporation, g/100 ml 0.001
carbonizable impurities, colour 30
platinum-cobalt scale, maximum 5
Grade AA differs in specifying an acetone maximum (20 ppm), a
minimum for ethanol (10 ppm), and in having a more stringent water
content specification (1.000 ppm, maximum) (Grayson, 1981).
2.2.2 Chemical properties
Methanol undergoes reactions that are typical of alcohols as a
chemical class. The reactions of particular industrial importance
include the following: dehydrogenation and oxidative dehydrogenation
over silver or molybdenum-iron oxide to form formaldehyde; the
acid-catalysed reaction with isobutylene to form methyl tertiary butyl
ether (MTBE); carbonylation to acetic acid catalysed by cobalt or
rhodium; esterification with organic acids and acid derivatives;
etherification; addition to unsaturated bonds and replacement of the
hydroxyl group (Grayson, 1981; Elvers et al., 1990).
2.3 Conversion factors
1 ppm = 1.31 mg/m3 (25°C, 1013hPa) 1 mmol/litre = 32 mg/litre
1 mg/m3 = 0.763 ppm (25°C, 1013hPa) 1 mg/litre =31.2 µmol/litre
(Adapted from Clayton & Clayton, 1982)
2.4 Analytical methods
Prior to the advent of sensitive gas chromatographic techniques,
the analysis of methanol in environmental, consumer and biological
samples was performed by procedures involving isolation of the
volatile alcohol and titrimetry. This was followed later by more
sensitive spectrophotometric methods based on the oxidation of
methanol to formaldehyde with potassium permanganate then reaction
with Schiff's reagent or rosaniline solution to produce an easily
recognizable and stable colour (Gettler, 1920; Boos, 1948; Skaug,
1956; Hindberg & Wieth, 1963; NIOSH, 1976).
The earliest procedures for the determination of methanol in
blood and urine were based on the initial distillation to isolate the
volatile alcohol (Gettler, 1920). Feldstein & Klendshog (1954)
determined methanol in biological fluids by initial microdiffusion
followed by oxidation to formaldehyde and subsequent reaction with
chromotropic acid (1,8-dihydroxy naphthalene-3,6-disulfonic acid). The
recovery ranged from 80 to 85% for less than 0.10 mg methanol. In the
procedure of Harger (1935), methanol was determined by oxidation with
bichromate to carbon dioxide and water followed by titration with a
mixture of ferrous sulfate and methyl orange. Jaselkis & Warriner
(1966) determined methanol in aqueous solution by titrimetry employing
xenon trioxide oxidation. Methanol was determined at a level of
0.03 mg with a relative standard deviation of 4%.
2.4.1 Environmental samples
The determination of methanol by primarily GC-FID procedures has
been frequently reported in ambient air, workplace air, fuels, fuel
emissions, sewage and aqueous solutions, soils, coal-gasification
condensate water and tobacco smoke.
The measurement of methanol in ambient and workplace air, usually
involves a preconcentration step in which the sample is passed through
a solid absorbent containing silica gel, Tenax GC, Porapak or
activated charcoal (NIOSH, 1976,1977,1984; CEC, 1988). It can also be
accomplished by on-column cryogenic trapping or can be analysed
directly. Direct reading infrared instruments with gas cuvettes can be
used for continuous monitoring of methanol in air (Lundberg, 1985).
2.4.1.1 Methanol in air
The use of absorption tubes to trap methanol from ambient and
workplace air with subsequent liquid or thermal desorption prior to
gas chromatographic analysis has been reported frequently. The US
National Institute of Occupational Safety and Health (NIOSH,
1977,1984) recommended the use of a glass tube (7 cm × 4 mm internal
diameter) containing two sections of 20-40 mesh silica gel separated
by a 2-mm portion of urethane foam (front=100 mg, back=50 mg). Water
is used to extract the methanol, which is separated on a 2 m × 2 mm
internal diameter glass column containing 60-80 mesh Tenax GC or the
equivalent using flame ionization detection (FID). The working range
is 25 to 900 mg/m3 (19 to 690 ppm) methanol for a 5-litre air sample.
The limit of detection has been reported to be 1.05 mg/m3 in a
3-litre air sample (NIOSH, 1976). At high concentrations of methanol
or at high relative humidity, a large silica gel tube is required
(700 mg silica gel front section). The injection, detector and column
temperatures are 200°C, 250-300°C and 80°C respectively. Positive
identification by mass spectrometry may be necessary in some cases,
and alternative gas chromatographic columns, e.g., SP-1000, SP-2100 or
FFAP, are also conformation aides.
Although GC-FID provides greater sensitivity than GC-MS, the
latter is generally considered more reliable for the measurement of
methanol in samples containing other alcohols or low molecular weight
oxygenates.Analysis of methanol in workplace air has been carried out
by head-space GC-FID using a column containing 15% Carbowax 1500 on
diatomaceous earth, 70-100 mesh operated at 100°C. The detection limit
was below 5 ml/m3 ( Heinrich & Angerer, 1982). Methanol in workplace
air was initially collected in silica gel tubes and the methanol
concentrations analysed by GC-FID equipped with a 50 m silica
capillary column containing Carbowax 20M. Additionally, methanol
vapour concentrations in the workplace have been analysed by a Miron-B
analyser with detection at a wavelength of 9.70 µm.
Methanol and other low molecular weight oxygenates have been
determined in ambient air by cryogradient sampling and two-dimensional
gas chromatography (Jonsson & Berg, 1983). Samples were initially
separated on a packed column (1,2,3-tris (2-cyanoethoxy)propane on
Chromosorb W-AW), then refocused on-line in a fused-silica capillary
cold trap, followed by on-line splitless reinjection onto a 50 m ×
0.3 mm internal diameter fused silica capillary column. The detection
limit for a typical oxygenate (3-methylbutanol) was 0.1 µg/m3 using a
3-litre sample. The detection limit for methanol was slightly higher.
Spectrophotometric methods have also been employed for the
determination of methanol in air. Aqueous potassium permanganate
acidified with phosphoric acid was used to absorb methanol from air
with the simultaneous oxidation to formaldehyde. After the addition of
p-aminoazobenzene and sulfur dioxide, the resulting pink dye was
determined spectrophotometrically at 505 nm. The limit of detection
was 5 µg methanol/ml air (Verma & Gupta, 1984).
Methanol from air was absorbed by acidified potassium
permanganate producing formaldehyde which on reaction with
4-nitroaniline produced a yellow dye determined spectroscopically at
395 nm (Upadhyay & Gupta, 1984).
Infrared spectrometry and infrared lasers have also been employed
for the determination of methanol in air (Diaz-Rueda et al., 1977;
Sweger & Travis, 1979). Methanol together with acetone, toluene and
ethyl acetate were recovered from 10 litres of air at a flow rate of
11 ml/min by passage through a tube containing 150 mg of activated
charcoal. The carbon disulfide extracts of the organic compounds were
determined by infrared at 1300 cm-1 using caesium bromide windows.
The minimum concentration of methanol detected quantitatively was
0.77 mg/m3 (0.60 ppm) and the minimum concentration required for
identification was 0.24 mg/m3 (0.18 ppm) (Diaz-Rueda et al., 1977).
Infrared lasers have been used to detect trace organic gases
including methanol. An air sample at 8 Tor was introduced to a
20-litre capacity sample cell, and laser radiation was detected
synchronously by a mercury-cadmium Te detector. The laser line
employed was P (34), the electric field was 1.40 kV/cm and the
measurement time was 2 min. The detection limit for methanol was
0.105 mg/m3 (0.08 ppm) (Sweger & Travis, 1979).
Methanol in the workplace can be measured by portable direct
reading instruments, real-time continuous monitoring systems and
passive dosimeters (NIOSH, 1976,1977,1984; Liesivouri & Savolainen,
1987; Kawai et al., 1990).
Kawai et al. (1990) described a personal diffusive badge type
that could absorb methanol vapour in linear relation to the exposure
duration up to 10 h and to exposure concentrations up to 1050 mg/m3
(800 ppm) the maximum duration and concentration tested respectively.
Additionally it was shown that the response to short-term peak
exposure was rapid enough and that no spontaneous desorption would
occur.
2.4.1.2 Methanol in fuels
Agarawal (1988) determined methanol quantitatively in commercial
gasoline via an initial extraction with ethylene glycol then by GC
utilizing a GB-1 fused silica capillary column (OV-1 equivalent, 60 m
× 0.32 mm internal diameter) and FID. The recovery of 4% methanol in
gasoline by this procedure was 95.4 ± 2.34% (SD).
In the procedure of Tackett (1987), gasoline samples were
injected directly on a Carbowax 20M column operated at 50°C for 3.0
min and then programmed to rise to 150°C at a rate of 10°C per min.
The calibration curve is linear up to 10% (v/v) methanol and the
detection limit was 0.2% employing a thermal conductivity detector.
Low molecular weight alcohols and MTBE were determined in
gasoline by GC-FID utilizing dual columns: 4.6 m × 3.2 mm o.d. column
packed with 30% m/m ethylene glycol succinate on Chromosorb P (85-100
mesh) and a 2.7 m × 3.2 mm o.d. stainless steel column packed with
Porapak P (80-100 mesh) operated at 150°C (Luke & Ray, 1984).
Gas chromatographic analyses of methanol, ethanol and tert-
butanol in gasoline have been reported by Pauls & McCoy (1981). The GC
column was 150 cm × 3 mm in o.d. stainless steel packed with Porapak R
(80-100 mesh) operated at 175°C and the injector and FID detector
temperatures were maintained at 250°C.
A direct liquid chromatographic method for the determination of
C1-C3 alcohols and water in gasoline-alcohol blends was described by
Zinbo (1984). The separation was performed on either one or two
microparticulate size-exclusion columns of ultrastyragel with toluene
as the mobile phase. The quantification of alcohols and water in the
effluent was achieved by a differential refractometer at 30°C. The
lower limits of detection for C1-C3 alcohols was 0.005 vol %. Methanol
in gasoline-alcohol blends has been determined by nuclear magnetic
resonance (Renzoni et al., 1985). The method takes advantage of a
window in the proton nuclear magnetic resonance spectrum of gasoline
that extends from a chemical shift of 2.8 to 6.8 ppm. Methanol was
quantified in gasoline by integration of the methyl singlet at
3.4 ppm. The method gave linear calibration curves in the range of
0-25% (v/v) methanol with a detection limit of less than 0.1%.
2.4.1.3 Methanol in fuel emissions
Methanol has been detected in motor vehicle emissions at levels
of 0.9 mg/m3 (0.69 ppm) and in ambient air by GC-FID utilizing a
360 cm × 0.27 cm internal diameter stainless steel column packed with
Porapak Q (50-80 mesh) operated at 150°C (Bellar & Sigsby, 1970).
Seizinger & Dimitriades (1972) determined methanol in simple
hydrocarbon fuel emissions utilizing GC with time-of-flight mass
spectrometry. The analytical procedure involved concentration of the
exhaust oxygenates drawn through a Chromosorb bed followed by GC-FID
initially on a 30 in by 1/4 in o.d. column packed with 10% 1,2,3-tris
(2-cyanoethoxy) propane (TCEP) programmed from -20°C to 110°C at
4°C/min. The second-stage column was a 45 m × 0.05 cm internal
diameter by 0.03 o.d Carbowax 20M support coated on tubular (SCOT)
column programmed from 60°C to 210°C at 10°C/min. The column effluent
was split for parallel detection with FID and mass spectrometry.
Methanol was found at levels of 0.1-0.8 mg/m3 (0.1-0.6 ppm) in
the exhaust of simple hydrocarbon fuels.
Methods for the quantification of evaporative emissions (running
losses, hot soak, diurnal and refuelling) from methanol-fuelled motor
vehicles (methanol/gasoline fuel mixtures of 100, 85, 50, 15 and 0%
methanol) have been described (Snow et al., 1989; Federal Register,
1989; Gabele & Knapp, 1993).
Methanol emissions from methanol-fuelled cars were determined by
GC employing a Quadrex 007 methyl silicone 50 m × 0.53 mm internal
diameter column with 5.0 µm film thickness. The separation was
affected isothermally at 75°C (limit of detection 0.25 µg/ml)
(Williams et al., 1990).
2.4.1.4 Methanol in sewage and aqueous solutions
Fox (1973) determined methanol at levels of 0.5-100 mg/litre
(0.5-100 ppm) in sewage or other aqueous solutions by GC-FID employing
a 0.5 m × 3.175 mm o.d. stainless steel column packed with Tenax GC
60/80 mesh and operated at 70°C isothermal.
C1-C4 alcohols in aqueous solution were determined
quantitatively by GC-FID using a 1 m × 0.32 cm stainless steel column
packed with 5% w/w Carbowax 20M on Chromosorb 101 (80-100 mesh) with a
column temperature of 65°C for methanol and ethanol and 100°C for n-
propanol and n-butanol (Sims, 1976).
Methanol and ethanol at the mg/litre level in aqueous solution
were determined by Komers & Sir (1976) utilizing a combination of
stripping and GC-FID technique. The alcohols were analysed as their
corresponding volatile nitrite on a 170 cm × 0.4 cm internal diameter
glass column containing Chromosorb 102 (80-120 mesh) operated at
104°C. Approximately 1 µg of the individual alcohol could be
determined in sample volumes of about 5 ml.
Mohr & King (1985) determined methanol in coal-gasification
condensate water by GC. Condensate water was injected directly on a 45
× 0.32 cm Porapak R column programmed from 80-200°C at 20°C/min.
A standard method for the analysis of methanol in raw, waste and
potable waters has been published by the UK Standing Committee of
Analysts (1982). The method is based on direct injection GC-FID using
a 2 m stainless steel column with 15% carbowax 1540 m chromosorb
W80-100 DMCS. The limit of detection is 0.11 mg/litre.
2.4.1.5 Methanol in soils
The biodegradation of methanol in gasolines by various soils was
determined by Novak et al. (1985). Methanol extracted in water (25%
v/v) was measured by direct injection GC-FID using a 2.1 m × 3 mm
stainless steel column packed with 0.2% Carbowax 1500 0n 80/100 mesh
Carbopak C at 120°C isothermal.
2.4.2 Foods, beverages and consumer products
Lund et al. (1981) determined methanol in orange and grapefruit
juice, fresh and canned, by GC-FID using a 1.5 × 3 mm column packed
with 50/80 mesh Porapak Q at 100°C with injector port and detector
block at 200°C.
Greizerstein (1981) utilized GC-FID and GC-MS for the analysis of
alcohols, aldehydes and esters in commercial beverages (beers, wines,
distilled spirits). Separations were carried out using a 3 m × 2 mm
internal diameter glass column packed with 30% Carbowax 20 M at 150°C.
A more satisfactory separation of methanol from the other congeners
was achieved using a 180-cm Porapak P column. Methanol was found at
levels of 6-27 mg/litre beer; 96-321 mg/litre in wines and
10-220 mg/litre in distilled spirits. Methanol in distilled liquors
and cordials has been determined by GC-FID (AOAC, 1990).
Rastogi (1993) analysed methanol content of 26 model and hobby
glues and found methanol in 12 of them by head-space GC-FID employing
capillary columns of different polarity. The polar GC column was a
Supelcowax 10, 60 m × 0.32 mm internal diameter; and the non-polar
column was a CP-Sil-5 CB, 50 m × 0.32 mm. The detection limit for
methanol was 20 mg/litre.
Methanol in wine vinegars was determined by GC-MS (Blanch et al.,
1992). Methanol with many other minor volatile components was
fractionated using a simultaneous distillation extraction technique
before GC analysis on a 4 m × 0.85 mm internal diameter micropacked
column coated with a mixture of Carbowax and bis-(2-ethylhexyl)-
sebecate (92:8), 4% on desilanized Volaspher A-2. The column
temperature was 60°C and the injector and FID detector were at 180°C.
2.4.3 Biological materials
A variety of primarily gas chromatographic methods have been
utilized for the determination of methanol in biological samples from
normal, poisoned and occupationally exposed individuals. Methanol
exposure has been measured in exhaled breath, blood and urine samples.
2.4.3.1 Methanol in exhaled air
Prior to analysis, expired air samples are normally collected in
sampling bags or glass containers or after preconcentration on Tenax
or other solid sorbents in adsorbent tubes and thermally desorbed, or
utilizing cryotraps (Franzblau et al., 1992a).
Free methanol has been detected and measured by GC in the expired
air of normal healthy humans with separations made on 1.52 m × 0.3 cm
columns filled with Anakrom ABS, 70-80 mesh coated with 2% N,N,-N,-N-
tetramethyl azeleamide and 8% behenyl alcohol at 86°C. The
concentration of methanol in nine subjects ranged from
0.06-0.32 µg/litre (Eriksen & Kulkarni, 1963). Methanol was only
infrequently detected in samples of human expired air and saliva by
Larsson (1965) employing GC-FID and a 1.75 mx 3.5 mm internal diameter
glass column containing polyethylene glycol (M=1500) 20% on Chromosorb
W.
Methanol in expired air and in head-space analysis of plasma was
determined as the nitrite ester utilizing GC-MS (Jones et al., 1983).
Condensed expired air samples were analysed on Porapak Q and the assay
of methanol nitrite ester was accomplished on a 2 m × 2 mm internal
diameter silanized glass column containing Tenax GC (30-60 mesh) at
60°C.
Krotosynski et al. (1977) analysed expired air from normal
healthy subjects using for sample preconcentration a 18 cm × 6 mm o.d.
stainless steel column containing Tenax GC. Sample analysis was
performed using GC-FID and a 91 m × 6 mm stainless steel column coated
with Emulphoron-870. Apart from methanol, 102 organic compounds were
detected.
Alveolar air of workers exposed to methanol was first collected
in gas sampling tubes and then analysed by GC-FID using a Porapak Q
(80-100 mesh) column at 150°C (Baumann & Angerer, 1979).
The detection of methanol and other endogenous compounds in
expired air by GC-FID with on-column concentration of sample and
separation on a 1.5 m × 3 mm o.d. stainless steel column packed with
Porapak Q, 80-100 mesh maintained at 35°C was described by Phillips &
Greenberg (1987).
The expired air of volunteer subjects exposed for periods of
about 90 min to atmospheres artificially contaminated with low levels
of methanol (ca. 130 mg/m3 (100 ppm)) was monitored during and
after the exposure using an atmospheric pressure ionization mass
spectrometer (API/MS) fitted with a direct breath analysis system
(Benoit et al., 1985).
A transportable Fourier Transform Infrared (FTIR) spectrometer
was utilized for the analysis of methanol vapour in alveolar and
ambient air in humans exposed to methanol vapour. The infrared
spectrum region used for methanol quantification was in the 950-1100
cm region. For the analysis of methanol in alveolar air with FTIR the
limit of detection for methanol was 0.4 mg/m3 (0.32 ppm), and for
methanol in ambient air the detection limit was 0.13 mg/m3 (0.1 ppm)
(Franzblau et al., 1992a).
2.4.3.2 Methanol in blood
A number of methods have been used to extract methanol from blood
prior to analysis including purge-and-trap, head-space analysis and
solvent extraction.
Baker et al. (1969) reported the simultaneous determination of
lower alcohols, acetone and acetaldehyde in blood by GC-FID utilizing
a 183 cm × 5 mm internal diameter column containing Porapak Q operated
at 100°C. The method did not require precipitation of protein prior to
analysis.
Methanol in whole blood and serum was analysed by GC-FID
employing 1.2 m and 1.8 m × 3 mm internal diameter glass columns
packed with 20% Hallcomid or 10% Carbowax on 60-80 mesh Diatopor TW
operated at 70°C (Mather & Assimos, 1965).
Blood serum was deproteinized and acetone and aliphatic alcohols
including methanol were determined by GC-FID using a pre-column of 3%
OV-1 on Gas Chrom Q and an analytical 30-m capillary column packed
with SPB-1 and operated at 35°C. Methanol and other alcohols were
separated in less than 3 min (Smith, 1984).
Methanol in deproteinized blood samples from occupationally
exposed workers was quantified by GC-FID employing a 1.8 m × 4 mm
internal diameter glass column packed with 60-80 mesh Carbopak B/5%
Carbowax 20M at 60°C. The detection limit for methanol was about
0.4 µg/ml (Lee et al., 1992).
Methanol in blood of occupationally exposed workers was
determined by head-space GC-FID utilizing a column containing 15%
Carbowax 1599 on diatomaceous earth, 70-80 mesh and operated at 70°C.
The detection limit was 0.6 mg/litre (Heinrich & Angerer, 1982).
The simultaneous determination of methanol, ethanol, acetone,
isopropanol and ethylene glycol in plasma by GC-FID was accomplished
using a 180 cm × 4 mm internal diameter glass column packed with
Porapak Q, 50-80 mesh. The column temperature was programmed from
199-210°C at 2°C/min, and the injection port and detector temperatures
were 210°C and 240°C respectively. The detection limit for methanol
was 0.1 nmol/ml. The procedure was recommended for methanol and
ethylene glycol intoxication cases (Cheung & Lin, 1987).
Methanol in blood from occupationally exposed workers was
determined directly without further pretreatment by GC-FID using a 4 m
× 3 mm glass column packed with 10% SBS 100 on Shimalite TPA, 60-80
mesh. The detector and oven were heated at 180°C and 60°C,
respectively (Kawai et al., 1991a).
Head-space GC-FID on methanol in blood from workers exposed at
sub-occupational exposure limits was reported by Kawai et al. (1992).
A 30 m × 0.53 mm capillary column coated with 1.0 um DB-Wax was used
with the injection port and detector heated at 200°C and the oven
temperature kept at 40°C for 1 min after the injection and then
elevated at a rate of 5°C/min to 110°C for 15 min. The detection limit
for methanol in blood was 100 µg/litre.
Leaf & Zatman (1952) utilized a colorimetric procedure for the
determination of methanol in air as well as in the blood and urine of
occupationally exposed workers in a methanol synthesis plant. The
procedure involved acid permanganate oxidation of methanol to
formaldehyde, which was then determined with a modified Schiff's
reagent. Concentrations of methanol up to 150 mg/litre were determined
to within 3%.
Determination of methanol in patients with acute methanol
poisoning was accomplished with a colorimetric procedure following
permanganate oxidation to formaldehyde and the subsequent reaction
with chromotropic acid (1,8-dihydroxy naphthalene 3,6-disulfonic
acid). Quantitative recovery of 100% was found for methanol following
the analysis of 3 ml of plasma, which required 45 min (Hindberg &
Wieth, 1963).
Accumulation of methanol in blood was detected in alcoholic
subjects during a 10-15 day period of chronic alcohol intake using
GC-FID and a 1.8 m column packed with Porapak Q, 80-100 mesh, or
Chromosorb 101 operated at 140°C (Majchrowicz & Mendelson, 1971). The
identity of methanol was also confirmed chemically using the
specificity of the colour reaction between permanganate and
formaldehyde.
Head-space GC was used to determine the concentrations of
methanol and ethanol in blood samples from 519 individuals suspected
of drinking and driving in Sweden. Methanol was determined in whole
blood without prior dilution with an internal standard. Carbopack C
(0.2% Carbowax 1500) was used as the stationary phase and the oven
temperature was 80°C (Jones & Lowinger, 1988).
Methanol in whole blood of poisoned patients was determined
without pretreatment by GC-FID using a 1800 mm × 4 mm internal
diameter glass column packed with 80-100 mesh Carbopack C/0.2% CW 1500
operated at 80°C; the detector temperature was 120°C (Jacobsen et al.,
1982a).
Serum methanol concentrations in men after oral administration of
the sweetening agent aspartame were determined by GC-MS utilizing a
fused silica capillary column 26 m × 0.22 mm internal diameter of
CPWAX 57 CB operated at 50°C isothermally (Davoli et al., 1986).
Methanol and formate in blood and urine of rats administered
methanol intravenously was determined by HPLC employing a REZEX-ROA-
organic acid column (300 mm × 7.8 mm internal diameter) and a
similarly packed pre-column (50 mm × 4.6 mm internal diameter). The
mobile phase was 0.043 N sulfuric acid with 10% acetonitrile at a flow
rate of 1 ml/min (Horton et al., 1992).
Methanol in serum has also been determined by high-field (500
MHZ) proton nuclear magnetic resonance at the 3.39 singlet peak. For
serum containing 20-500 mg of added methanol/litre, peak area was a
linear function of concentration (r=0.998). This NMR technique
permitted the determination of methanol and acetone in blood serum at
a level of less than 1mM (Bock, 1982).
Pollack & Kawagoe (1991) determined methanol in deproteinized
whole blood of rats by capillary GC-FID with direct column injection
utilizing a 15 m × 0.54 mm internal diameter fused silica capillary
column coated with Carbowax and operated at 35°C. The limit of
detection was 2 µg/ml.
2.4.3.3 Methanol in urine
Sedivec et al. (1981) determined methanol in urine in five
volunteers exposed to methanol vapour for 8 h. Head-space GC-FID was
used with a 120 cm × 3 mm column packed with Chromosorb 102, 60-80
mesh at 120°C. The detection limit of methanol was 0.1 mg/litre. The
methanol content in urine of 20 subjects occupationally exposed to
methanol was determined by head-space GC-FID utilizing a column
containing Porapak QS, 80-100 mesh and operated at 130°C. The
detection limit was 0.6 mg/litre (Heinrich & Angerer, 1982).
Methanol in the urine of exposed workers was determined by
head-space GC-FID using a 4.1 m × 3.2 mm glass column containing 10%
SBS-100 on Shimalite TPA, 60-80 mesh. The oven and injection port
temperatures were 60°C and 180°C respectively. The limit of detection
for methanol in urine was 0.1 mg/litre (Kawai et al., 1991b, 1992).
Urinary methanol as a measure of occupational exposure was
determined by GC-FID utilizing a 2 m glass column packed with Porapak
Q, 80-100 mesh. The detection limit for methanol was 0.32 mg/litre
(Liesivouri & Savolainen, 1987).
Urine concentrations of methanol in volunteers who had ingested
small amounts of methanol was determined by head-space GC-FID using
Tenax GC as the column packing (Ferry et al., 1980).
2.4.3.4 Methanol in miscellaneous biological tissues
Methanol and other alcohols have been determined in tissue
homogenates either per se or as their nitrite esters by GC-FID
employing a 1.8 m × 6 mm o.d. glass column packed with Chromosorb 101
operated at 145°C. The sensitivity was 8 µg per g of tissue (Gessner,
1970).
2.4.3.5 Methanol metabolites in biological fluids
The principal metabolite of methanol in humans and monkeys is
formate and it has been shown that accumulation of blood formate at
higher levels of methanol exposure coincides with the development of
metabolic acidosis and visual system toxicities (Clay et al., 1975;
McMartin et al., 1975; Baumbach et al., 1977; Tephly, 1991). Formate
is an endogenous product of single carbon metabolism and is normally
found in the urine of healthy individuals.
Formate has been analysed in blood and urine samples primarily by
enzymatic methods with a colorimetric or fluorimetric end-point or by
derivatization followed by analysis by GC-FID. Formate in plasma has
also been determined by isotachophoresis (Sejersted et al., 1983).
Ferry et al. (1980) measured formic acid as an ethyl ester formed
by the treatment of urine with 30% sulfuric acid in ethanol. The
samples were analysed by head-space GC-FID on a column packed with 10%
silar 10C on Chrom Q.
The analysis of formic acid in blood was performed via an initial
transformation of formic acid by concentrated sulfuric acid into water
and carbon monoxide, the latter being reduced to methane on a
catalytic column and analysed directly by GC-FID (Angerer & Lehnert,
1977; Baumann & Angerer, 1979; Heinrich & Angerer, 1982).
Urinary formic acid was determined after the methylation of the
acid and its conversion to N,N-dimethylformamide with GC-FID equipped
with a 50-m silica capillary column containing Carbowax 20M liquid
phase. The detection limit was 2.3 mg/litre (Liesivouri & Savolainen,
1987).
Franzblau et al. (1992b) found that urinary formic acid in
specimens collected 16 h following cessation of methanol exposure and
analysed by head-space GC-FID may not be an appropriate approach to
assess methanol exposure biologically.
Enzymatic methods for the determination of formate are based
primarily on the enzyme-catalysed conversion of formate to carbon
dioxide in the presence of nicotinamide adenine dinucleotide (NAD),
generating NADH as the other reaction product. NADH formation can be
subsequently measured directly or reacted in a coupled reaction to
generate a fluorescent or coloured complex.
A specific assay for formic acid in body fluids based on the
reaction of formate with bacterial formate dehydrogenase coupled to a
diaphorase-catalysed reduction of the non-fluorescent dye resazurin to
the fluorescent substance resorufin was reported by Makar et al.
(1975) and Makar & Tephly (1982). This permitted the accurate
determination of about 6 mg formate/litre blood at excitation
wavelength of 565 nm and an emission wavelength of 590 nm (Makar et
al., 1975; Makar & Tephly, 1982).
A serum formate enzymic assay based on modifications of the
formate dehydrogenase (FDH)-diaphorase procedure using NAD-diaphorase-
iodonitrotetrazolium violet to develop a red-coloured complex, which
is measured at 500 nm, was described by Grady & Osterloh (1986). The
calibration curve was linear over the formate range of 0 to
400 mg/litre.
Formate in plasma was determined by Lee et al. (1992) employing
an enzymatic procedure (Grady & Osterloh, 1986; Buttery & Chamberlin,
1988) and measured spectrophotometrically at 510 nm. The detection
limit was about 3 µg/ml.
Lee et al. (1992) determined that formate associated with acute
methanol toxicity in humans does not accumulate in blood when
atmospheric methanol exposure concentrations are below the
occupational threshold limit value of 260 mg/m3 (200 ppm) for 6 h in
exposed healthy volunteers.
d'Alessandro et al. (1994) found that serum and urine formate
determinations were not sensitive biological markers of methanol
exposure at the threshold limit value (TLV) in human volunteers.
Formate in serum was analysed by the enzymatic-colorimetric procedure
of Grady & Osterloh (1986). The sensitivity of the method was
0.5 mg/litre of formate in serum.
Buttery & Chamberlin (1988) developed an enzymatic method for the
determination of abnormal levels of formate in plasma requiring no
deproteinization and utilizing a stable colour reagent consisting of
phenazine methosulfate, p-iodonitrotetrazolium and NAD to produce a
stable red formazan colour. The precision at 1.0 and 5.0 mmol/litre
formate was 2.9% and 1.7%, respectively, within-day and 5.5% and 2.3%,
respectively, between days.
Urinary formic acid was determined using formate dehydrogenase
(FDH) in the presence of NAD. The detection limit was 0.5 mg/litre.
The normal formic acid excretion in urine is between 2.0 and
30 mg/litre (Triebig & Schaller, 1980).
3. SOURCES OF HUMAN AND ENVIRONMENTAL EXPOSURE
3.1 Natural occurrence
Methanol occurs naturally in humans, animals and plants (Axelrod
& Daly, 1965; CEC, 1988). It is a natural constituent of blood, urine
and saliva (Leaf & Zatman, 1952) and expired air (Erikssen & Kulkarni,
1963; Larsson, 1965; Krotosynski et al., 1979; Jones et al., 1990),
and has also been found in mother's milk (Pellizzari et al., 1982).
Humans have a background body burden of 0.5 mg/kg body weight (Kavet
& Nauss, 1990).
Levels of methanol in expired air are reported to range from 0.06
to 0.49 µg/litre (46-377 ppb) (Eriksen & Kulkarni, 1963). Methanol has
been detected in the expired air of normal, healthy non-smoking
subjects at a mean level of 0.5 ng/litre (Krotosynski et al., 1979).
It is believed that dietary sources are only partial contributors
to the total body pool of methanol (Stegink et al., 1981). It has been
suggested that methanol is formed by the activities of the intestinal
microflora or by other enzymatic processes (Axelrod & Daly, 1965). The
methanol-forming enzyme was shown to be protein carboxylmethylase, an
enzyme that methylates the carboxyl groups of proteins (Kim, 1973;
Morin & Liss, 1973).
Natural emission sources of methanol include volcanic gasses,
vegetation, microbes and insects (Owens et al., 1969; Holzer et al.,
1977; Graedel et al., 1986). Isidorov et al. (1985) identified
methanol emissions of evergreen cyprus in the forests of Northern
Europe and Asia. Methanol was identified as one of the volatile
components emitted by alfalfa (Owens et al., 1969) and it is formed
during biological decomposition of biological wastes, sewage and
sludges (US EPA, 1975; Howard, 1990; Nielsen et al., 1993).
3.2 Anthropogenic sources
The major anthropogenic sources of methanol include its
production, storage and use, principally its use as a solvent, as a
chemical intermediate, in the production of glycol ethers, and in the
manufacture of charcoal, and exhaust from vehicle engines (US EPA,
1976a,b, 1980a,b; CEC, 1988).
3.2.1 Production levels and processes
3.2.1.1 Production processes
The earliest important source of methanol ("wood alcohol") was
the dry distillation of wood at about 350°C, which was employed from
around 1830 to 1930. In countries where wood is plentiful and wood
products form an important industry, methanol is still obtained by
this procedure (ILO, 1983).
In 1880, about 1.5 million litres of wood alcohol were produced
in the USA while in 1910 the amount had increased to over 3 million
litres (Tyson & Schoenberg, 1914). However methanol produced from wood
contained more contaminants, primarily acetone, acetic acid and allyl
alcohol, than the chemical-grade methanol currently available
(Grayson, 1981; Elvers et al., 1990). Methanol was also produced as
one of the products of the non-catalytic oxidation of hydrocarbons (a
procedure discontinued in the USA in 1973), and as a by-product of
Fischer-Tropsch synthesis, which is no longer industrially important
(Grayson, 1981).
Modern industrial scale methanol production is based exclusively
on the catalytic conversion of pressurized synthesis gas (hydrogen,
carbon monoxide and carbon dioxide) in the presence of metallic
heterogenous catalysts. All carbonaceous materials such as coal, coke,
natural gas, petroleum and fractions obtained from petroleum (asphalt,
gasoline, gaseous compounds) can be employed as starting materials for
synthesis gas production (Grayson, 1981; Elvers et al., 1990).
The required synthesis pressure is dependant upon the activity of
the particular metallic catalyst employed, with copper-containing zinc
oxide-alumina catalysts being the most effective in industrial
methanol plants (Elvers et al., 1990). By convention the processes are
classified according to the pressure used: low-pressure processes,
50-100 atmospheres; medium-pressure processes, 100-250 atmospheres;
and high-pressure processes, 250-350 atmospheres. Low-pressure
technology is the most widely employed globally and accounted for 55%
of the USA methanol capacity in 1980 (Grayson, 1981).
Almost all the methanol produced in the USA is made from natural
gas. This is steam reformed to produce synthesis gas, which is
converted to methanol by low-pressure processes. A small amount of
methanol is obtained as a by-product from the oxidation of butane to
produce acetic acid and from the destructive distillation of wood to
produce charcoal (Grayson, 1981; Elvers et al., 1990).
The composition of methanol obtained directly from synthesis
without any purification or with only partial purification varies
according to the synthesis (e.g., pressure, catalyst, feedstock). The
principal impurities include 5-20% (by volume) water, higher alcohols
(principally ethanol), methyl formate and higher esters, and smaller
amounts of ethers and aldehydes (Grayson, 1981; Elvers et al., 1990).
Methanol is purified by distillation, the complexity required
depending on the desired methanol purity and the purity of the crude
methanol (Grayson, 1981; Elvers et al., 1990).
Natural gas, petroleum residues and naphtha accounted for 90% of
worldwide methanol capacity in 1980, miscellaneous off-gas sources
constituting the remaining 10%. Natural gas alone accounted for 70%,
petroleum residues 15%, and naphtha 5% (Grayson, 1981). Natural gas
feedstock accounted for 75% in the USA and 70% of global capacity in
1980. Methanol produced from residual oil accounted for approximately
15% of USA and worldwide capacity in 1980, while naphtha and coal
feedstocks accounted for approximately 5% and 2%, respectively, of
worldwide methanol capacity in 1980 (Grayson, 1981). About 90% of the
global methanol capacity is currently based on natural gas (SRI,
1992).
The production of methanol from coal, being independent of oil
and natural gas supplies, is noted to be an attractive alternative
feed stock in some quarters (Grayson, 1981; CEC, 1988). Newer
approaches to the production of methanol that have been suggested
include the catalytic conversion from carbon dioxide and hydrogen
avoiding conventional steam reforming (Rotman, 1994a) and the direct
catalytic conversion of methane to methanol (Rotman, 1994b).
3.2.1.2 Production figures
As shown in Table 2, worldwide annual capacity for methanol
production has increased over the past decades from approximately 15 ×
106 tonnes in 1979 (Grayson, 1981) to 21 × 106 tonnes in 1989
(Elvers et al., 1990) and more than 22.1 × 106 tonnes in the
beginning of 1991 (SRI, 1992). Worldwide demand was projected to rise
further to about 25.8 × 106 tonnes in 1994 (Anon., 1991; Nielsen et
al., 1993) and 30.1 × 106 tonnes in 1995 (SRI, 1992). The data
available do not allow capacity and production figures to be compared;
however, it is assumed that approximately 80% of production capacity
is utilized (Fiedler et al., 1990).
The USA and Canada are the largest methanol-producing countries.
About 85% of Canada's production is exported to the USA, Japan and
Europe (Heath, 1991). In Western Europe, Germany, the Netherlands and
the United Kingdom are the major methanol-producing countries,
accounting for 7%, 3% and over 2% of the world capacity, respectively
(SRI, 1992). The production of methanol in Germany in 1991 and 1992
amounted to 715 000 and 770 000 tonnes respectively.
The annual capacity in Eastern Europe was estimated to be 5.8 ×
106 tonnes in 1987. The production in the former USSR was 3.28 × 106
tonnes and 3.21 × 106 tonnes in 1987 and 1988, respectively (Rippen,
1990).
Table 2. Methanol production or production capacity (× 106 tonnes per year) from 1978 to 1995
Year World-wide USA Canada Western Japan Capacity/ Reference
Europe production
1978 12 3.4 3 1 capacity Grayson (1981)
production
1979 15 4.05 3.45 1.35 capacity Grayson (1981)
1980 2.5 production CEC (1988)
1981 8 production CEC (1988)
1983 15.9 5.52 (33%) 1.75 (11%) 2.53 1.27 (8%) capacity SRI (1992)
production CEC (1988)
1988 1.91 production Anderson (1993)
1989 21 capacity Elvers et al. (1990)
19 production
1990 22.3 capacity Anon. (1991);
Nielsen et al. (1993)
1991 22.1 4.42 (20%) 2.21 (10%) 2.65 (12%)a 0.22 (1%) capacity SRI (1992)
1991 2.22 0.077 production Anderson (1993)
1992 2.15 0.034 production Anderson (1993)
1992 3.66 2.15 production Reisch (1994)
1993 4.78 production Reisch (1994)
1995 30.1 capacity SRI (1992)
a Only Germany, the Netherlands and the United Kingdom.
The figures in Table 2 indicate a major shift in methanol
production from the developed countries to the developing areas. In
fact, the methanol industry underwent large structural changes during
the 1980s as a result of the discovery of large natural gas fields in
remote regions having little demand for natural gas themselves. Since
methanol production is a very suitable alternative for marketing
natural gases, a number of methanol production plants for export were
built or proposed to be built in Asia (Bahrein, Oman, Qatar, Saudi
Arabia, Indonesia, Malaysia), South America (Chile, Mexico,
Venezuela), the Caribbean (Trinidad) and in New Zealand and Norway
(Fiedler et al., 1990; SRI, 1992). The largest single train plant
based on this concept came on stream in southern Chile in 1988 with an
annual output of 750 000 tonnes (Fiedler et al., 1990).
Future trends in methanol production and demand are being driven
to a large extent by increasing demand for methyl tertiary butyl ether
(MTBE), which is used in gasoline blending as an octane enhancer and
to reduce carbon monoxide emissions (Anon., 1991; Morris, 1993;
Nielsen et al., 1993).
3.2.2 Uses
During the 1890s, the market for methanol (then better known as
wood alcohol) increased as a commercial product and as a solvent for
use in the workplace. It was included in many consumer products such
as witch hazel, Jamaica ginger, vanilla extract and perfumes (Wood &
Buller, 1904). The most notorious use of wood alcohol was and
continues to be as an adulterant in alcoholic beverages, which has led
to large-scale episodes of poisonings since 1900 (Bennett et al.,
1953; Kane et al., 1968).
Historically, in terms of commercial usage, about half of all
methanol produced has been used to produce formaldehyde. Other earlier
large-volume chemicals based on methanol include acetic acid, dimethyl
terephthalate, glycol methyl ethers, methyl halides, methylamines,
methyl acrylate and various solvent uses (Grayson, 1981; CEC, 1988;
Elvers et al., 1990; Nielsen et al., 1993).
3.2.2.1 Use as feedstock for chemical syntheses
Approximately 70% of the methanol produced worldwide is used as
feedstock for chemical syntheses. As shown in Table 3, formaldehyde,
methyl tertiary butyl ether (MTBE), acetic acid, methyl methacrylate,
and dimethyl terephthalate are, in order of importance, the main
chemicals produced from methanol. Methyl halides produced from
methanol include methyl chloride, methylene chloride and chloroform.
Nearly all the formaldehyde manufactured worldwide is produced by
oxidation of methanol with atmospheric oxygen. The annual formaldehyde
production was projected to increase at a rate of 3%, but because
other bulk products have higher growth rates, its relative importance
with respect to methanol use has decreased (Elvers et al., 1990;
Fiedler et al., 1990).
Table 3. Use pattern for methanol (as a percentage of production) according to region and year
Global Global USA USA Japan Western Europe Brazil
1979 1988 1973 1985 n.g. 1985 n.g.
Use for synthesis of:
formaldehyde 52 40 39 30 47 50 60
MTBE 4 20 8 - 5 -
acetic acid 6 9 3.4 12 10 5 -
dimethyl terephthalate 4 6.1 4 1 4 16
methyl methacrylate 4 3.7 4 6 3 2
methyl halides 8a 6.1 9 3 6 -
methyl amines 3.3 4 2 4 9
glycol methyl ethers 1.1
Direct use
solvent 10 6 6 2
fuel 6 - 5 -
Miscellaneous 14 16.9 13 25 12 11
Referenceb [1] [2] [3] [4] [4] [4] [4]
a together with methyl amines production
b Reference: [1] Kennedy & Shanks (1981); [2] Elvers et al. (1990); [3] US EPA (1980a); [4] Rippen (1990)
n.g. = year not given
MTBE has become an important octane-enhancing blending component
in gasoline, particularly in the USA where the Clean Air Act
Amendments of 1990 have prompted further steps toward reducing
emissions from motor vehicles by changing the formulations of
gasoline. This is achieved by using so-called oxygenated fuel, i.e.
fuel containing at least 2% oxygen by weight in the form of
oxygenates, but less benzene and other aromatic compounds than
conventional fuel (Health Effects Institute, 1996). MTBE is produced
by reacting methanol with isobutene in acid ion exchangers. In 1987,
MTBE (production of 1.6 × 106 tonnes) ranked 32nd among the top 50
chemicals produced in the USA (Scholz et al., 1990). In 1993, 11 ×
106 tonnes were produced, ranking MTBE ninth of the top 50 chemicals
(Reisch, 1994).
Acetic acid is produced by carbonylation of methanol with carbon
monoxide. Annual growth rates of 6% have been estimated (Fiedler et
al., 1990).
Methanol is present in a broad variety of commercial and consumer
products including shellacs, paints, varnishes, mixed solvents in
duplicating machines (95% concentration or greater), antifreeze and
gasoline deicers (generally containing 35-95% methanol), windshield
washer fluid (contains 35-90% methanol), cleansing solutions
(containing around 5% methanol), model and hobby glues and adhesives,
and Sterno ("canned heat") containing 4% methanol (Posner, 1975; US
EPA, 1980a; CEC, 1988; ATSDR, 1993).
Methanol is also used in the denitrification of wastewater,
sewage treatment application (carbon source for bacteria to aid in the
anaerobic conversion of nitrates to nitrogen and carbon dioxide), as a
substrate for fermentation production of animal feed protein (single
cell protein), as a hydrate inhibitor in natural gas, and in the
methanolysis of polyethylene terephthalate (PET) from recycled plastic
wastes (Posner, 1975; US EPA, 1980a; Kennedy & Shanks, 1981; ATSDR,
1993).
3.2.2.2 Use as fuel
Methanol is a potential substitute for petroleum. It can be
directly used in fuel as a replacement for gasoline in gasoline and
diesel blends. Methanol is in favour over conventional fuels because
of its lower ozone-forming potential, lower emissions of some
pollutants, particularly benzene and polycyclic aromatic hydrocarbons
and sulfur compounds, and low evaporative emissions. On the other
hand, the possibility of higher formaldehyde emissions, its higher
acute toxicity and, at present, lower cost-efficiency favour
conventional fuels (CONCAWE, 1995).
For use in gasoline engines, pure methanol (so-called M100 fuel)
or mixtures of 3, 15 and 85% methanol with conventional petroleum
products (M3, M15, M85) are most common. In diesel engines methanol
cannot be used as an exclusive fuel because of its low cetane number
that would impose proper ignition. Therefore, methanol is injected
into the cylinder after ignition of the conventional diesel fuel
(Fiedler et al., 1990).
3.2.2.3 Other uses
Methanol is used in refrigeration systems, e.g., in ethylene
plants, and as an antifreeze in heating and cooling circuits. However,
its use as an engine antifreeze has been replaced by glycol-based
products. Methanol is added to natural gas at the pumping stations of
pipelines to prevent formation of gas hydrates at low temperature and
can be recycled after removal of water. Methanol is also used as an
absorption agent in gas scrubbers to remove, for example, carbon
dioxide and hydrogen sulfide. According to Table 3, large amounts of
methanol are used as a solvent. Pure methanol is not usually used
alone as a solvent, but is included in solvent mixtures (Fiedler et
al., 1990).
3.2.2.4 Losses into the environment
Given the high production volume, widespread use and physical and
chemical properties of methanol, there is a very high potential for
large amounts of methanol to be released to the environment,
principally to air (US EPA, 1976a,b, 1980a,b, 1994; Nielsen et al.,
1993). Emissions of methanol primarily occur from miscellaneous
solvent usage, methanol production, end-product manufacturing, and
bulk storage and handling losses. The largest source of emissions of
methanol is the miscellaneous solvent use category.
US EPA (1980b) estimated emission factors for the release of
methanol and volatile organic compounds (VOC) from the low-pressure
synthesis of methanol from natural gas in a model plant with a
capacity of 450 000 tonnes/year. The process and capacity were typical
of those built in the late 1970s. The overall emission factors were
estimated to be: uncontrolled emissions, 1.56 kg methanol/tonne
produced; controlled emissions, 0.14 kg methanol/tonne produced
(Nielsen et al., 1993).
It was estimated that about 1% of the methanol used in the
production of formaldehyde would be released to the environment during
the production process by which formaldehyde is produced by either a
metallic silver-catalyst process or a metal oxide-catalyst process (US
EPA, 1976a; 1980b). In the oxidation-dehydrogenation process with
metallic silver catalyst, 0.89 kg methanol/tonne of 39% (by weight)
formaldehyde solution was released principally from the product
absorber vents, and 1.24 kg methanol/tonne from the fractionator
vents. The production of formaldehyde using the catalytic oxidation,
metal oxide catalyst process resulted in the release of 1.93 kg
methanol/tonne of 37% formaldehyde solution with emissions from the
absorber vent (US EPA, 1980b).
US EPA (1994) reported that methanol was the most released
chemical to the environment (air, water and land) based on the 1992
Toxic Release Inventory which utilized 81 016 individual chemical
reports from a total of 23 630 facilities (approximately 65% of
facilities reporting). The air, water and land releases of methanol
totalled 1.09 × 105 tonnes, consisting of 1.53 × 104 tonnes of
fugitive or non-point air emissions, 72 956 tonnes of stack or point
air emissions, 7444 tonnes of surface water discharges and 15 095
tonnes released to land. Additionally, 1.283 × 104 tonnes were
transferred via underground injection.
Methanol had the largest off-site transfers (51 672 tonnes) to
publicly owned treatment works (POTWs) in 1992. During the same
period, methanol ranked third largest of the Toxic Release Inventory
Chemicals with off-site transfers for treatment. The total transfers
to treatment were 18 098 tonnes, consisting of 4 tonnes for
solidification, 10 295 tonnes for incineration/thermal treatment, 1971
tonnes of incineration/insignificant fuel value; 5311 tonnes for
wastewater treatment and 147 tonnes to waste broker-waste treatment. A
total of 493 980 tonnes of methanol was treated, consisting of 260 875
tonnes treated on-site and 197 400 tonnes off-site. A total of 1510
tonnes of methanol was released to land, primarily to on-site
landfills (US EPA, 1994).
The total amount of methanol release in Canada in 1993 was
306 222 tonnes distributed as follows: air, 15 326; water, 14 248;
underground, 819 and land, 205 (Ministry of Supply & Services Canada,
1993).
Tail pipe emissions as well as evaporative emissions are
monitored by a number of agencies. Emissions and air quality modelling
results have been reported from methanol/gasoline blends in prototype
flexible/variable fuelled vehicles (US EPA, 1991; Auto/Oil Air Quality
Research Program, 1992, 1994). Motor vehicle emissions are affected in
various ways by the use of methanol fuels in production flexible/
variable fuel vehicles. Higher molecular weight hydrocarbons are
reduced and carbon monoxide is reduced under some circumstances, while
increases in methanol and formaldehyde can occur (US EPA, 1991).
Methanol has been found in significant amounts in the exhaust
from gasoline-powered vehicles as well as in diesel exhausts. Methanol
was measured at levels of 100-226 mg/kg in the exhaust emissions from
non-catalyst vehicles fuelled with isobutane/methanol/gasoline
(2/15/83; M-15). Methanol emissions from a light-duty diesel vehicle
fuelled with 95% methanol were one order of magnitude higher
(3.4 g/kg) (Jonsson et al., 1985).
Chang & Rudy (1990) reported methanol emission factors for
vehicles fuelled by M-85 (85% methanol + 15% gasoline) and M-100 (100%
methanol) in the USA. For M-85-fuelled vehicles, factors were 0.156-
0.7 g methanol/mile driven in exhaust emissions and 0.055-0.25 g
methanol/mile driven in evaporative emissions. For M-100 fuelled
vehicles, they were 0.5 g methanol/mile driven in exhaust emissions
and 0.072-0.134 g methanol/mile driven in evaporative emissions.
Methanol was found at levels of 130-800 µg/m3 (0.1 to 0.6 ppm)
in the exhaust from nine hydrocarbon test fuels, e.g., iso-octane,
iso-octene, benzene, 2-methyl-2-butene, toluene, o-xylene,
benzene/ n-pentane, toluene/ n-pentane and iso-octane/toluene/
iso-octene (Seizinger & Dimitriades, 1972).
Methanol, formaldehyde and hydrocarbon emissions from methanol-
fuelled cars were reported by Williams et al. (1990). The variable
methanol-fuelled vehicles using fuel mixtures of 100, 85, 50, 15 and
0% methanol and a dedicated methanol vehicle all gave similar emission
patterns. The organic composition of the exhaust was 85-90% methanol,
5-7% formaldehyde and 3-9% hydrocarbons.
4. ENVIRONMENTAL TRANSPORT, DISTRIBUTION AND TRANSFORMATION
4.1 Transport and distribution between media
Methanol is released into the environment from both natural and
man-made sources, the latter being the most significant. Methanol
is released predominantly from its production and use as a solvent
in industrial processes (in extraction, washing, drying and
recrystallization operations), and to a lesser degree from a variety
of industrial processes and domestic uses (US EPA, 1980a,b; Graedel et
al., 1986; CEC, 1988; Howard, 1990; Nielsen et al., 1993).
Methanol volatilization half-lives of 5.3 and 2.6 days have been
estimated for a model river (1 m deep) and an environmental pond,
respectively (Howard, 1990).
Methanol is expected to exist almost entirely in the vapour phase
in the ambient atmosphere, based on its vapour pressure (Eisenreich
et al., 1981; Graedel et al., 1986). Because of methanol's water
solubility, rain would be expected to physically remove some methanol
from the air (US EPA, 1980a,b; Snider & Dawson, 1985).
Methanol has been found in the atmosphere (Graedel et al.,
1986). It can be the product of atmospheric alkane chemistry with
concentrations as high as 131 µg/m3 (100 ppb) being found. Methanol
is expected to become an important additional trace gas in the
atmosphere due to its projected increased use as an alternative fuel
to gasoline or in a gasoline blend (CEC, 1988; Chang & Rudy, 1990).
The miscibility of methanol in water and its low octanol/water
partition coefficient suggest high mobility in soil. Lœkke (1984)
studied the adsorption of methanol onto three soil types at 6°C. The
soils tested comprised two sandy soils (organic matter contents of
0.09 and 0.1%), and a clay soil (organic matter content of 0.22%).
Methanol solutions with concentrations of 0.1, 1.0, 9 and 90 mg/litre
were used in 1-h exposure studies. Adsorption coefficients for all
soil methanol concentrations and soil types ranged from 0.13 to 0.61,
indicating methanol has a low adsorptive capacity on these soils.
However Nielsen et al. (1993) suggested that the soils used in the
Lœkke (1984) study had low organic matter contents compared to typical
agricultural surface soil which can have organic matter contents of 1
to 2%, and up to 5% in some soils. A soil containing a typical amount
of organic matter might therefore be expected to retain methanol and
prevent it from reaching the subsoil.
Additionally, the relatively high vapour pressure and low
adsorptive capacity suggests significant evaporation from dry
surfaces.
4.2 Transformation
4.2.1 Biodegradation
Methanol is readily biodegradable in soil and sediments, both
under aerobic and anaerobic conditions. A large number of strains/
genera of microorganisms have been identified as capable of using
methanol as a growth substrate (Hanson, 1980; Braun & Stolp,
1985; Nielsen et al., 1993). These include Pseudomonas sp.,
Methylobacterium organophilium; Hyphomicrobium sp., Desulfovibrio;
Streptomyces sp., Rhodopseudomonas acidophilia; Paracoccus
denitrificans; Microcyclus aquaticus; Thiobacillus novellus;
Micrococcus denitrificans; Achromobacter 1L (isolated from activated
sewage sludge) and Mycobacterium 50 (isolated from activated sewage
sludge). Most microorganisms possess the enzyme alcohol dehydrogenase
which is necessary for methanol oxidation. The methanogen,
Methanosarcine barkeri can grow on and produce methane from methanol
(Hippe et al., 1979).
The following genera of methanol-oxidizing yeasts have been
reported: Pichia; Saccharomyces; Hansenula; Rhodotorula; Kloechera;
Candida; Torulopsis (Stensel et al., 1973; Hanson, 1980; Nielsen et
al., 1993). Okpokwasili & Amanchukwu (1988) isolated Candida sp.
from Niger Delta sediment which utilized methanol as a growth
substrate.
4.2.1.1 Water and sewage sludge
In a closed bottle test, according to OECD guideline 301D,
methanol was found to be readily biodegradable with 99% COD removal
after the test period of 30 days (Hüls AG, 1978). In another closed
bottle test using unadapted inoculum from domestic sewage the
degradation of methanol at concentrations of 3, 7 or 10 mg/litre in
both freshwater (settled domestic wastewater) and synthetic seawater
incubated for a maximum of 20 days under aerobic conditions was
studied by Price et al. (1974). Methanol was readily degraded in both
inocula at all concentrations with average disappearance of methanol
as follows: a) after 5 days, 76% bio-oxidation in fresh water and 69%
in salt water; b) after 10 days, 88% bio-oxidation in fresh water and
84% in salt water; c) after 15 days, 91% bio-oxidation in fresh water
and 85% in salt water and d) after 20 days, 95% bio-oxidation in fresh
water and 97% in salt water.
Matsui et al. (1988) studied the biodegradability of methanol in
a batch reactor using activated sludge from an industrial wastewater
treatment plant which was acclimatized to the wastewater originating
from a petrochemical complex in Japan. Methanol at an initial
concentration of 100 mg/litre and an acclimation period of 1 day was
found to be highly biodegradable with 91% COD removal and 92% TOC
removal achieved.
Incubation of 0.05 mg methanol/litre for 5 days in activated
sludge from a municipal sewage plant resulted in the degradation of
37% of the methanol (Freitag et al., 1985). Hatfield (1957) found that
at a feed rate of 333 or 500 mg/litre, methanol was virtually
completely oxidized (with a major portion of the BOD and COD removed)
by acclimated microorganisms within 6 h in a settled domestic sewage
inoculum.
The microbial metabolism of methanol in a model activated sludge
system monitored by Swain & Somerville (1978) revealed that methanol
was not broken down when added transiently (0.23% by volume) to the
system operating with a retention time of 11 h. However adaptation of
the sludge in such a system to 0.1% by volume occurred over a period
of several days. After 2 days acclimation, about 50% of the methanol
was utilized, and after 6 days acclimation more than 80% of the
methanol had been metabolized. There were no apparent toxic effects
caused by the addition of methanol (0.1% by volume) to the sludge
prior to and after adaptation to methanol.
The anaerobic treatment of wastes containing methanol and higher
alcohols (approximately 50:50 mix) was studied by Lettinga et al.
(1981). In batch and continuous experiments using an inoculum
consisting of sugar beet waste and active anaerobic sludge, the
breakdown of methanol began within a few days while the breakdown of
higher alcohols occurred immediately depending on the initial load of
waste applied.
Denitrification is facilitated by heterotrophic and autotrophic
bacteria. Heterotrophic bacteria require a carbon source for their
growth and cell metabolism which can be supplied by methanol (Stensel
et al., 1973; Nyberg et al., 1992; Jansen et al., 1993; Upton, 1993).
Bacteria such as the organisms of the genera Pseudomonas,
Micrococcus, Achromobacter, Spirillum, and Bacillus reduce
nitrate, nitrogen oxide and nitrous oxide under anaerobic conditions.
The addition of methanol to promote denitrification has been suggested
in situations where nitrate accumulates, and methanol has been
added as an economic exogenous organic carbon source to increase
denitrification (Stensel et al., 1973; Nyberg et al., 1992; Jansen et
al., 1993; Upton, 1993).
At a wastewater treatment plant in Malmo, Sweden, complete
denitrification was obtained after approximately one month at 10°C
after methanol was added for denitrification. Microscopic examination
revealed a growing population of budding and/or appendaged bacteria,
presumably Hyphomicrobrium spp. reaching a stable maximum at the
time when optimal nitrate removal occurred (Nyberg et al., 1992)
Upton (1993) described a pilot study in the United Kingdom
indicating that denitrification in deep-bed sand filters is a feasible
technology utilizing methanol addition. Nitrogen removals greater than
70% were possible at winter sewage temperatures.
Several other laboratory studies using a variety of methodologies
have demonstrated the rapid biodegradation of methanol by sewage
organisms. These show degradation of between 66 and 95%, and usually
greater than 80%, within five days (Kempa, 1976; Hüls AG, 1978; Matsui
et al., 1988).
4.2.1.2 Soils and sediments
Methanol is biodegradable in soils and sediments, both under
aerobic and anaerobic conditions. Methanol is a normal growth
substrate for many soil microorganisms, which are capable of
completely mineralizing methanol to carbon monoxide and water (CEC,
1988; Howard, 1990; Howard et al., 1991; Nielsen et al., 1993).
Methanol at concentrations of up to 1000 mg/litre was degraded to
non-measurable amounts within a year or less in subsurface soil and
ground water sites in Pennsylvania, New York and Virginia (USA)
believed to be previously uncontaminated. Complete degradation of
100 g methanol/litre occurred in less than 30 days in one aerobic soil
sample from a Pennsylvania site (Novak et al., 1985).
Scheunert et al. (1987) monitored the formation of 14CO2 from
labelled methanol in aerobic and anaerobic suspended soil and found
methanol to be readily degradable after 5 days incubation at 35°C.
Rates and patterns of biodegradation of methanol in surface and
subsurface soils from eight sites in New York, Pennsylvania and
Virginia in static microcosms under anaerobic conditions were
evaluated by Hickman & Novak (1989) and Hickman et al. (1989). The
rates of methanol degradation varied considerably between sites, but
the soils could be characterized into two general types, namely fast
soils, in which degradation rates were high and rates were increased
by addition of nitrate and sulfate, and slow soils, in which
biodegradation rates were low and decreased by the addition of nitrate
or sulfate and inhibition of sulfate increased degradation rates.
Biodegradation rates in subsurface soils were generally within the
range of 0.5-1.1 mg/litre per day and indicated that no acclimation
period was required. Biodegradation rates were calculated and used to
estimate a half-life of between 58 and 263 days for methanol in these
soils (Hickman et al., 1989).
Compared to other substrates studied, e.g., acetate,
trimethylamine and methylamine, methanol (at concentrations less than
3 µM) was degraded relatively slowly mainly to carbon dioxide,
principally via sulfite-reducing organisms, and could not be
considered a significant in situ precursor in surface sediments of
an intertidal zone in Maine, USA (King et al., 1983).
Methanol was found to be an important substrate for methanogenic
bacteria in anaerobic sediments (highly reduced and containing methane
and hydrogen sulfide), collected from a salt marsh located in
San Francisco Bay, California. The sediments were homogenized
anaerobically with San Francisco Bay water and 310-340 µmol methanol/
flask, resulting in 83-91% conversion to methane, carbon dioxide and
water after 3 days (Oremland et al., 1982).
A sulfate-reducing bacterium of the genus Desulfovibrio, which
is capable of degrading methanol after growth on pyruvate, malate or
fumarate, completely converted anaerobic samples of 14C-methanol to
carbon dioxide. However the 14C-label was not used as a carbon source
by the bacterium and was not assimilated into cellular material (Braun
& Stolp, 1985).
4.2.2 Abiotic degradation
4.2.2.1 Water
In a 5-day experiment, 14C-labelled methanol applied to
soil-water suspensions under both aerobic and anaerobic conditions
yielded 53.4 and 46.3% 14CO2, respectively (Sche