ETHANOL, ACETALDEHYDE AND
GASTROINTESTINAL FLORA
Regulatory factors and pathophysiological consequences of microbial
ethanol oxidation and acetaldehyde production in the digestive tract
Jyrki Tillonen
Research Unit of Alcohol Diseases
University of Helsinki
Finland
ACADEMIC DISSERTATION
HELSINKI 2000
ISBN 952-91-2603-4 PDF
2
Supervised by
Professor Mikko Salaspuro, M.D.
Research Unit Of Alcohol Diseases,
Department of Medicine
University of Helsinki
Reviewed by
Docent Onni Niemelä, M.D.
University of Oulu,
Department of Medical Biochemistry
and
Docent Risto Roine, M.D.
Finnish Office for Health Care
Technology Assessment
Opponent
Professor Eero Kivilaakso, M.D.
Helsinki University Central Hospital,
Department of Gastroenterological and
General surgery
3
To my family
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CONTENTS
ABBREVIATIONS 6
ORIGINAL PUBLICATIONS 7
1. INTRODUCTION 8
2. REVIEW OF THE LITERATURE 10
2.1. Biochemical characteristics of ethanol and acetaldehyde 10
2.2. Human oral and gastrointestinal microflora - an overview 11
2.3. Distribution of ethanol in the body 18
2.4. Hepatic ethanol and acetaldehyde metabolism 19
2.5. Metabolism of ethanol and acetaldehyde in the digestive tract
2.6. Microbial ethanol fermentation and oxidation 23
2.7. Alcohol and the alimentary tract 26
2.8. Alcohol and digestive tract cancers 28
2.9. Organ toxicity of acetaldehyde 32
3. AIMS OF THE STUDY 36
4. MATERIALS AND METHODS 37
4.1. Ethical considerations 37
4.2. Acetaldehyde production by human colonic contents in vitro (I) 37
4.3. The effect of ciprofloxacin on ethanol elimination in humans (II) 38
4.4. The effect of ciprofloxacin on human faecal flora and acetaldehyde production (II) 39
4.5. Sustained ethanol and metronidazole treatment in rats (III) 39
4.6. The effect of acetaldehyde on intestinal folate levels in rats (IV) 41
4.7. Human saliva studies (V, VI) 42
4.8. Gas chromatographic measurements of ethanol and acetaldehyde 43
4.9. Statistical analysis 43
5. RESULTS 44
5.1. Enzymatic production of acetaldehyde by human colonic contents in vitro (I) 44
5.2. The effect of ciprofloxacin on ethanol elimination in humans (II) 46
5.3. The effect of ciprofloxacin on human faecal flora and acetaldehyde production (II) 47
5.4. The effect of sustained ethanol and metronidazole treatment on
intracolonic acetaldehyde production in rats (III) 47
5.5. The effect of ethanol and metronidazole treatment on
blood ethanol and acetaldehyde levels in rats (III) 49
5.6. The effect of acetaldehyde on intestinal folate levels in rats (IV) 50
5.7. Factors influencing salivary acetaldehyde production in humans (V) 51
5.8. Microbes associated with acetaldehyde production in the human oral cavity (V, VI) 52
5
6. DISCUSSION 54
6.1. Role of catalase in acetaldehyde production by colonic contents 54
6.2. Role of colonic bacteria in extrahepatic ethanol elimination in humans 55
6.3. The effect of long-term ethanol and metronidazole treatment on
intracolonic acetaldehyde levels 56
6.4. The effect of ethanol and metronidazole treatment on
hepatic ethanol and acetaldehyde metabolism 57
6.5. The effect of acetaldehyde on intestinal folate levels in rats –
a possible carcinogenic action of acetaldehyde 58
6.6. Acetaldehyde in saliva: influencing factors 58
6.7. Microbes associated with acetaldehyde production in the human oral cavity 59
7. SUMMARY AND CONCLUSION 61
REFERENCES 63
6
ABBREVIATIONS
ADH alcohol dehydrogenase
ALDH aldehyde dehydrogenase
ANOVA analysis of variance
3-AT 3-amino-1,2,3-triazole
AUC area under the curve
BMI body mass index
CFU colony forming units
CIPRO ciprofloxacin
CYP cytochrome P450
DMH 1,2-dimethylhydrazine
DNA deoxyribonucleic acid
EER ethanol elimination rate
EGF epidermal growth factor
FPM first pass metabolism
GI gastrointestinal
GOX glucose oxidase
H2O2 hydrogen peroxide
IARC International Agency for Research on Cancer
Ig intragastric
Ip intraperitoneal
kDa kiloDalton
Km Michaelis constant
MAF the mucosa-associated flora
MEOS microsomal ethanol oxidizing system
Mol wt molecular weight
4-MP 4-methylpyrazole
NAD nicotinamide adenine dinucleotide
NADH reduced nicotinamide adenine dinucleotide
NDEA N-nitrosodiethylamine
NDMA nitrosodimethylamine
NDPA N-nitrodi-n-propylamine
N2-Et-dg N2-ethyldeoxyguanosine
O6-MeGT O6 methylguanine transferase
PCA perchloric acid
RER rough endoplasmic reticulum
RR relative risk
SA sodium azide
SCFA short-chain fatty acids
SEM standard error of the mean
SER smooth endoplasmic reticulum
Sp species
Ssp subspecies
Vd volume of distribution
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ORIGINAL PUBLICATIONS
This thesis is based on the following studies which are referred to in the text by their Roman
numerals:
I Tillonen J, Kaihovaara P, Jousimies-Somer H, Heine R, Salaspuro M (1998) Role
of catalase in in vitro acetaldehyde formation by human colonic contents. Alcohol
Clin Exp Res 22:1113-1119.
II Tillonen J, Homann N, Rautio M, Jousimies-Somer H, Salaspuro M (1999)
Ciprofloxacin decreases the rate of ethanol elimination in humans. Gut 44:347-
352.
III Tillonen J, Väkeväinen S, Salaspuro V, Zhang Y, Rautio M, Jousimies-Somer H,
Lindros K, Salaspuro M (2000) Metronidazole increases intracolonic but not
peripheral blood acetaldehyde in chronic ethanol-treated rats. Alcohol Clin Exp
Res 24:570-575.
IV Homann N, Tillonen J, Salaspuro M (2000) Microbially produced acetaldehyde
from ethanol may increase the risk of colon cancer via folate deficiency. Int J
Cancer 86:169-173.
V Homann N, Tillonen J, Meurman JH, Rintamäki H, Lindqvist C, Rautio M,
Jousimies-Somer H, Salaspuro M (2000) Increased salivary acetaldehyde levels
in heavy drinkers and smokers: a microbiological approach to oral cavity cancer.
Carcinogenesis 21:663-668.
VI Tillonen J, Homann N, Rautio M, Jousimies-Somer H, Salaspuro M (1999) Role
of yeasts in the salivary acetaldehyde production from ethanol among risk groups
for ethanol-associated oral cavity cancer. Alcohol Clin Exp Res 23:1409-1415.
8
1. INTRODUCTION
For millenia the consumption of alcoholic beverages has contributed to the pleasure of
eating and drinking in many cultures of the world. In addition to livening up the social
atmosphere, light alcohol drinking may also have beneficial effects on human health. An
example of this is the “J-shaped” relation between the risk of coronary heart disease and
alcohol intake. Overall morbidity is lower among those who drink lightly than those
who drink more heavily or who do not drink at all (Klatsky, 1994). While the optimal or
non-injurious levels of alcohol intake are difficult to estimate, they have been thought to
be quite low, approximately 10-19 g/day for men and less than 10 g/day for women
(Kalant and Poikolainen, 1999). When taken in excess, alcohol has devastating effects
on human health by leading to breakdown of bodily functions and damaging virtually
every organ of the body. Alcohol use may also lead to alcoholism, which can be defined
as “the extreme dependence on excessive amounts of alcohol associated with a
cumulative pattern of deviant behaviours”.
Excessive alcohol consumption is known to increase the risk of developing several
diseases of the liver, which are principally “fatty liver”, hepatitis, and cirrhosis. Also
well-known are cases of alcohol-induced acute or chronic pancreatitis. Ethanol itself has
been thought to be the hepatotoxic agent, but since only a relatively small proportion of
heavy drinkers develop the most severe forms of liver damage, it is probable that other
factors are also involved (Lindros, 1995). The pathogenesis of alcohol-induced
pancreatic injury is still obscure, although major hypotheses so far have emphasized
ethanol-induced changes in the pancreatic ductal system or the toxic effects of ethanol
on pancreatic exocrine metabolism (Singh and Simsek, 1990).
Less is known about other multiple gastrointestinal symptoms and organ toxicities
associated with heavy alcohol use. There is evidence of small intestinal dysfunction
after chronic alcohol consumption, including increased mucosal permeability,
promotion of bacterial overgrowth, altered gut motility, and impaired salt and water
absorption. This can lead to diarrhea, dyspepsia, nausea, and finally to malnutrition,
which are common findings among actively drinking alcoholics (Persson, 1991). The
association between alcohol consumption and certain digestive tract neoplasia has also
been well established. Epidemiological studies have shown that cancers of the mouth,
esophagus, and larynx are associated with alcohol consumption and that the risk
increases in a dose-dependent manner (Doll et al., 1999). Likewise, high alcohol intake
is a suspected risk factor for colorectal cancer. Although this subject has been debated,
two different meta-analyses both reach the conclusion that alcohol leads to a small but
significantly increased cancer risk, especially for the left colon and the rectum (Kune
and Vitetta, 1992; Longnecker et al., 1990).
The mechanism for the increased cancer risk associated with alcohol consumption is not
clear, but has been believed to be at least in part due to the carcinogenic action of the
first metabolite of ethanol, acetaldehyde (IARC, 1985, 1999). This notion is strongly
supported by recent epidemiological studies which show that GI-tract cancer risk is
markedly increased among heavy drinking Asian individuals with a genetically deficient
ability to remove acetaldehyde (Yokoyama et al., 1998). The reactivity of acetaldehyde
may also involve it in promoting organ toxicity other than malignant transformation.
9
During recent years it has become evident that the colonic microbes take part in ethanol
metabolism not only by fermenting sugars to ethanol, but also by oxidizing exogenous
ethanol to acetaldehyde (Jokelainen, 1997; Salaspuro, 1996, 1997). Similarly, oral
microflora have been shown to produce high concentrations of acetaldehyde from
ethanol (Homann et al., 1997a). Ethanol oxidation and consequent acetaldehyde
production by gut microbes occurs at ethanol concentrations that are relevant to those
after moderate alcohol drinking (Homann et al., 1997a; Jokelainen et al., 1994).
Furthermore, since the capacity of intestinal mucosa and flora to metabolise
acetaldehyde further is limited, acetaldehyde accumulates locally in the areas of the
digestive tract covered by microbes (Koivisto and Salaspuro, 1996; Nosova et al.,
1998).
Due to its high reactivity, toxicity and carcinogenicity, acetaldehyde can be expected to
cause organ damage wherever it exists at high concentrations. Therefore, microbial
ethanol oxidation and consequent acetaldehyde production may have important
implications for the pathogenesis of symptoms and organ toxicity associated with
excessive alcohol use. Understanding the mechanisms behind alcohol-induced
gastrointestinal morbidity is helpful in their management and a prerequisite for their
prevention. The present study thus investigates the enzymes involved in microbial
ethanol oxidation, its contribution to total ethanol elimination, and possible regulatory
factors, and is intended to obtain evidence about possible organ toxicity related to
microbial acetaldehyde production.
10
2. REVIEW OF THE LITERATURE
2.1. BIOCHEMICAL CHARACTERISTICS OF ETHANOL AND
ACETALDEHYDE
Ethyl alcohol (CH3CH2OH; Mol wt 46.1) is the accurate term for ethanol. The
synonyms ”alcohol and alcoholic beverages” are also commonly used in the literature
and in colloquial language. The term “alcohol” is, however, slightly misleading, since
several other alcohols, like methanol, also exist. In the context of this thesis the terms
ethanol and alcohol are used as synonyms, and alcoholic beverages means any products
that contain ethanol.
Ethanol is obtained by fermentation of carbohydrates contained in a variety of natural
products. Preparation of absolute anhydrous ethanol (percentage by weight
approximately 99.5%) for research purposes needs special distillation procedures. The
density of absolute ethanol at 20 C compared with water at 4 C is 0.789. The melting
point of absolute ethanol is -114.1 C and the boiling point is 78.5 C. Important
characteristics of ethanol are its small molecular size and miscibility in water in any
proportion. However, ethanol is only slightly soluble in fat; tissue fat takes up about 4%
of the amount of ethanol dissolved in an equal volume of water (IARC, 1988; Wallgren
and Barry III, 1970).
A number of different systems are used to indicate the ethanol content, dosage, solutions
for administration and concentration in body fluids. To indicate ethanol content or
concentration of ethanol in alcoholic beverages, percentage by weight (% w/w) or
volume (% v/v) are used. The dosage for scientific reasons should preferably be
expressed as weight of ethanol given per unit of body weight of the test organism (g/kg
body weight). For solutions to be administered, when the basis is pure ethanol, the most
convenient way is to prepare solutions that contain a known weight of ethanol in a given
volume of the final solution. The clearest expression is thus percentage weight by
volume or % w/v. Percentage by volume (%v/v) can also be employed. There are
several ways of expressing ethanol concentrations in body fluids and in in vitro
incubations. Concentrations are normally stated in millimoles per litre (mM) or as a
mille scale o/oo. One per mille equals one gram ethanol per litre, or 0.1 % w/v, or 21.7
mM. In some scientific publications, blood alcohol is also given as “mg %”, indicating
milligrams ethanol per 100 ml. Table 1 shows the equivalence of units used in
measuring concentrations of ethanol in body fluids.
Units used in this book are as follows: blood and body fluid ethanol concentrations are
generally given as mM. Dosages given are expressed as g/kg body weight, and solutions
for that purpose either in %w/v or %v/v.
11
Table 1. Equivalence of units used in measuring
concentration of ethanol in body fluids
(Modified from Wallgren and Barry III, 1970)
Per cent (%) Per mille (‰) mg/100 ml mM
g/100 ml mg/ml or g/l (“mg%)
0.001 0.01 1 0.217
0.01 0.1 10 2.17
0.1 1.0 100 21.7
0.2 2.0 200 43.4
0.3 3.0 300 65.1
0.4 4.0 400 86.8
0.5 5.0 500 108.5
0.6 6.0 600 130.2
0.7 7.0 700 151.9
0.8 8.0 800 173.6
0.9 9.0 900 195.3
1.0 10.0 1,000 217.0
Acetaldehyde (CH3CHO; Mol wt 44.1) is a byproduct of the organic chemicals
industry. It also occurs in vehicle exhaust and the smoke of tobacco cigarettes. For our
research purposes the main source of acetaldehyde formation is intracellular oxidation
of ethanol. The density of pure acetaldehyde at 20 C is 0.778, and its melting point is -
123.5 C and the boiling point 20.1 C. Like ethanol, acetaldehyde is miscible with water
and most common organic solvents (IARC, 1985, 1999). Since the concentrations of
acetaldehyde in this thesis are given either as μM or μmol/l, which are equivalent, an
acetaldehyde concentration of 100 μM equals 4.4 mg/l.
2.2 HUMAN ORAL AND GASTROINTESTINAL MICROFLORA - AN
OVERVIEW
Salivary microbial flora
The mouth cannot be regarded as a single, uniform environment. The majority of
investigations on the flora of the mouth have been concerned either with saliva or dental
plaque. For the aims and purposes of this thesis, the most important flora in the mouth is
that of the saliva. The reason for this is that ethanol is present in saliva in concentrations
comparable to those of blood ethanol (Jones, 1979), and saliva is in close contact with
the mucosa of the oropharynx and esophagus.
The salivary flora is derived from the dislodgement of microorganisms from various
locations in the oral cavity i.e. the teeth, tongue, cheek and pharyngeal mucous
membranes (Herrera et al., 1988; Nolte, 1977). Adult human saliva contains
approximately 6 x 109 microorganisms per millilitre (Nolte, 1977). Streptococci, the
facultative anaerobic Gram-positive organisms, have been isolated from all sites in the
mouth and comprise a large proportion of the normal oral flora. On average,
Streptococci represent about 45% of the total cultivable flora from saliva and the term
Streptococcus viridans group is often used to generalize these bacteria. It includes,
however, at least 5 different species, of which Streptococcus salivarius appears to
comprise a significant proportion. Anaerobic streptococcus forms part of the anaerobic
12
flora of the oral cavity (Marsh, 1980; Nolte, 1977). Aerobic Gram-positive
Staphylococci, Stomatococcus, and Micrococci have also been isolated from the oral
cavity and saliva, but not in large quantities. Corynebacterium, Lactobacillus and
Actinomyces are Gram-positive rods frequently found in human oral flora, and
consisting of aerobic, facultative anaerobic and strictly anaerobic species. Aerobic
Neisseria and strictly anaerobic Veillonella, which are Gram-negative cocci, have been
isolated in low numbers from most sites in the oral cavity and saliva. The majority of
aerobic or facultatively anaerobic Gram-negative rods fall into the genus Haemophilus
(Marsh, 1980). Yeasts are aerobic microorganisms that can be isolated from
approximately 40% of clinically healthy mouths, and Candida albicans is the most
dominant species (Stenderup, 1990). Most anaerobic oral Gram-negative rods belong to
the genus Bacteroides (Marsh, 1980).
Flora of the stomach and small intestine
All bacteria able to live as commensals in the human body are killed by incubation at
pH values below 3. The pH of the normal resting gastric juice is below 3 and so the
normal resting gastric juice is bacteria free. However, even in young normochlorhydric
persons, the lumen is not bacteria-free for the whole day. During a meal the gastric acid
is buffered, allowing swallowed salivary bacteria to survive or even to proliferate.
However, when the pH returns to less than 3 these swallowed organisms are killed (Hill,
1995). As a consequence, a resident bacterial flora in the stomach can only occur when
gastric acid secretion is impaired to the point that the pH does not fall below 3-4, even
in the resting stomach. Thus, most of the organisms found in the stomach very likely
represent the most acid-resistant components of the oral flora; Lactobacillus,
Streptococcus viridans, Neisseria, Staphylococcus, Bacteroides and Peptostreptococcus
are the genera best represented (Gustafsson, 1982; Hill, 1985). Impairing gastric acid
secretion leads, however, to bacterial overgrowth in the stomach (Drasar et al., 1969).
Conditions resulting in achlorhydria or hypochlorhydria include for example gastric
surgery that includes vagotomy, pernicious anaemia or chronic atrophic gastritis, or the
prolonged use of histamine 2 receptor antagonists or proton pump inhibitors. It has been
shown that treatment with antacids or cimetidine raises bacterial counts 10- to 100-fold
(Snepar et al. 1982). Moreover, it has been shown that gastric and duodenal bacterial
overgrowth is considerably higher in patients treated with omeprazole compared to
cimetidine. This was explained by the more pronounced inhibition of gastric acid
secretion (Thorens et al., 1996).
An apparent exception is infection with Helicobacter pylori. This organism colonizes
the mucosa below the mucin barrier and is able to resist local acid secretion via its
urease activity. Since the mucosal barrier protects the organisms from luminal acid they
are able to proliferate in a locally pH-controlled environment (Marshall et al., 1990).
When the gastric contents enter the small bowel they are mixed with large volumes of
biliary and pancreatic secretions, many of which are bactericidal and help to sterilize
this material. Furthermore, there is extensive fluid secretion from the bowel mucosa,
which serves to flush the crypts and prevent colonization of the mucosal layer. Small
bowel transit time is only two to four hours, an additional barrier to small bowel
colonization. For these reasons the normal small bowel is in general almost sterile or
contains a very sparse flora of transient organisms (Hill, 1985, 1995). Anaerobes only
slightly outnumber facultative organisms, Streptococci, Lactobacillus, Veillonella,
13
yeasts, and Staphylococci being found (Justesen et al., 1984). A resident flora can only
establish itself in areas of stasis, such as diverticulae, surgical blind loops, coeliac
disease or in tropical sprue (Hill, 1995). Additionally, as with the stomach, the use of
drugs to diminish gastric acid secretion has also been shown to lead to bacterial
overgrowth in the jejunal fluid (Shindo et al., 1998).
In the distal ileum, mean bacterial counts are much higher than in the proximal small
intestine, and the flora more closely resembles colonic flora with higher counts of
coliforms and Bacteroides. It should be noted that “coliform” is a common name for
those bacteria belonging to the Enterobacteriaceae family that are able to ferment
lactose. It includes Escherichia coli and most other Enterobacteriaceae species
belonging to the normal human flora. Since the terminal ileum appears to be a
“transitional” zone between the relatively sterile upper small intestine and the colon
with its rich bacterial population, its flora being similar to that of the caecum, although
in smaller numbers, and probably results at least in part from reflux through the
ileocaecal valve (Hill, 1995).
Large intestinal microflora
Several problems confront anyone attempting to define the composition of the intestinal
microflora in different parts of the large intestine. More than 400 different bacterial
species and approximately 1014 individual bacteria inhabit a human colon (Goldin,
1990). It has been estimated that a complete bacteriological description of one faecal
sample takes a year of laboratory work (Simon and Gorbach, 1984). The normal colonic
flora is usually inferred from the composition of the faecal flora. However, bacterial
counts vary throughout the large bowel, and the numbers found in faecal specimens may
not accurately represent counts found in other locations. Moreover, faecal flora
represents only the luminal flora and the flora associated with mucosal epithelia differs
markedly (next chapter). Bentley et al. (1972) compared the microflora of the transverse
colon, caecum, and terminal ileum with the microflora of stool samples. The highest
bacterial counts were obtained from stool samples. Microbial counts in the transverse
colon and caecum were on average 2-4 logarithmic values lower than in stool samples,
and counts were even lower in the terminal ileum. Although there were substantial
numerical differences between stool cultures and cultures from various locations in the
large bowel, there did not appear to be marked qualitative differences in the flora.
A characteristic of the luminal flora of the large intestine is that anaerobes outnumber
aerobes by a factor of 100 to 1000 (Cummings, 1983; Simon and Gorbach, 1986).
Several reports indicate that five genera account for the majority of the viable forms of
anaerobic bacteria: Bacteroides, Eubacterium, Bifidobacterium, Peptostreptococcus and
Fusobacterium. Various aerobic, microaerophilic and facultative anaerobic organisms
are also present in the colonic flora, the most common being Enterobacteriaceae,
Enterococcus, Lactobacillus, and yeasts (Salminen et al., 1995). Altogether, it has been
estimated that bacteria account for 35-50% of the volume of the contents of the human
colon. This equals 41-57% of the dry weight of colonic contents (Salminen et al., 1995).
The faecal flora is not the same in any two individuals. The concentration of each
bacterium can vary by as much as 100 000-fold between individuals within a given
group (Moore et al., 1978). It is, however, widely accepted that the faecal flora in one
individual is relatively stable over time (Bornside, 1978). The effect of diet on faecal
14
flora is controversial. It has been shown that there are differences in the faecal flora
between people consuming quite different diets (Aries et al., 1969; Finegold et al.,
1974), but the effect of dietary alterations has turned out to be extremely difficult to
demonstrate (Hill and Drasar, 1975). There is a consensus that dietary alterations change
the composition of the faecal flora only slightly and very slowly or not at all (Bornside,
1978; Hill and Drasar, 1975). Studies of the metabolic activity of the flora based on
measurements of bacterial enzymes have, however, revealed changes in the colonic flora
as a function of diet (Simon and Gorbach, 1984). Although faecal flora in general is
quite stable over prolonged periods of time, hospitalisation, for example, has been
shown to lead to rapid colonization by the specific E. coli serogroups associated with a
particular institute (Simon and Gorbach, 1984). The use of certain antimicrobial agents
is also a potent way to alter colonic microflora.
The colonic mucosa-associated flora (MAF)
Studies on animals have suggested the existence of a specific mucosal-dependent flora
in the colon (Savage, 1970). In contrast to the enormous amount of literature on the
bacteria of faeces, there is little data on the bacteriology of the mucosa-associated flora
(MAF) in humans. Nevertheless, analysis of biopsy material and specimens of surgically
excised tissue have shown that human colon has a flora associated with the mucosa
which is distinct from that of the gut lumen. It is reproducible, stable, and responds to
antibiotic treatment differently from that of the lumen (Bleday et al., 1993; Hill, 1995;
Nelson and Mata, 1970; Peach et al., 1975). One of the most important features of MAF
is that it has approximately equal representation of aerobic and anaerobic organisms
compared with the luminal flora (Marks et al., 1979; Peach et al., 1975). Facultative
anaerobes belonging to the Enterobacteriaceae family are well represented in the MAF
(Marks et al., 1979; Peach et al., 1978), whereas anaerobes have been shown to be
mainly Bacteroides species (Poxton et al., 1997).
The mechanisms by which microbes adhere to the mucosal membrane depend on a
variety of factors. Dietary fibres may influence the composition of the bacterial flora by
providing nutrients or altering the environmental conditions including peristaltic rate or
mucous composition (Savage, 1978). The normal function of the absorptive epithelium
presumably depends upon a suitable oxygen tension, which in turn depends upon blood
flow, arterial oxygen content, and oxygen utilization. One of the earliest studies
measuring oxygen tension in the gut lumen was done by using domestic duck. Oxygen
tension (PO2) was found to be about 25 mm Hg close to the mucosal villi area, while it
was 50 times lower in the centre of the lumen (Crompton et al., 1965). Studies with
dogs have shown that the intestinal mucosa has an oxygen tension of the order of 40 mm
Hg, which is between a quarter and a third of that in air and approximately similar to
that of the venous blood (Hamilton et al., 1968). One study with humans showed about
the same magnitude of oxygen tension in mucosa of the gut (45 mm Hg) (Dawson et al.,
1965). In contrast, flatus usually has a PO2 of less than 15 mm Hg (Askevold, 1956).
Utilization of O2 by colonic bacteria is thought to lower the intraluminal PO2 to the
level present in flatus. Moreover, it has been shown that conventional rats have a lower
intraluminal PO2 and higher pCO2 than germ-free rats. This is probably because of the
consumption of O2 and the production of CO2 by bacterial metabolism (Bornside et al.,
1973). Taken together, the amount of oxygen diffused to the colonic mucosa may be the
predominant factor explaining the proportionally higher counts of aerobic and
facultative organisms in the MAF than in the luminal flora.
15
It has been suggested that the importance of the microbes colonizing the mucosa is that
they modulate the function of the mucosal barrier. Disturbing the balance of MAF has
been shown to lead to alterations in paracellular pathways at the level of the tight
junctions, thereby increasing mucosal permeability (Spitz et al., 1994). Moreover,
intestinal microorganisms are able to oxidize and reduce many types of organic
compound. Consequently, it has been shown that the MAF is of importance in
regulating enzyme levels in the intestinal mucosa of the rat (Hietanen and Hänninen,
1971). This is of great importance, since aerobic and facultative anaerobic microbes are
able to oxidize ethanol to acetaldehyde. Because the ratio of aerobes to anaerobes is
approximately 1:1 in the vicinity of the mucosa, this could be an important site for
bacterial ethanol metabolism in the gut. It has, indeed, been shown that conventional
rats have significantly higher acetaldehyde levels in the mucosa of the rectum and
caecum than germ-free rats after ethanol administration (Seitz et al., 1990).
The effect of antimicrobial agents on human faecal flora with special reference
to ciprofloxacin and metronidazole
The administration of an antibiotic is undoubtedly the most potent way of altering the
markedly stable microflora of the human body. Since many antimicrobial agents cause
changes in the colonic flora, the severity of which depends largely upon the
concentration of the agent in the luminal contents, factors other than the width of the
antibacterial spectrum may be of importance for the ecological consequences of
antibiotic treatment in the colon (Norrby, 1986). Accordingly, the faecal microflora can
be influenced by orally taken antimicrobial agents because of incomplete absorption,
secretion into the bile, or secretion from the intestinal mucosa. Parenteral antimicrobial
agents which are secreted into the bile or from the intestinal mucosa can also cause
significant disturbances in the large intestinal microflora (Nord et al., 1986).
Ciprofloxacin is a broad spectrum fluoroquinolone antimicrobial agent. The primary
mechanism of the action of ciprofloxacin is the inhibition of bacterial DNA gyrase,
which disrupts bacterial DNA replication. After oral administration ciprofloxacin has an
approximate bioavailability of 70% and maximum plasma concentrations are achieved
in 1 to 2 hours. The drug has a large apparent volume of distribution (2.1 to 5 L/kg after
oral or intravenous administration) and becomes concentrated in many body tissues and
fluids, including bile, kidney, liver, gallbladder, prostate and lung tissue. Ciprofloxacin
is excreted largely unmetabolised in the urine and faeces, although small amounts of
metabolites have also been detected (Davis et al., 1996). Furthermore, ciprofloxacin is
partly eliminated through the intestinal wall (Rohwedder et al., 1990), and the
concentrations of ciprofloxacin in the faeces and intestinal mucosa are higher than the
corresponding serum levels (Brismar et al., 1990). This transintestinal elimination
pattern may explain the particular ability of this drug to modify the colonic flora.
The effect of ciprofloxacin in vivo on the composition of faecal flora has been studied
extensively in healthy volunteers. The usual dosage regimens have been from 600 to
1000 mg/day for five or more days. These studies clearly demonstrate the marked
reduction or complete eradication of Enterobacteriaceae. This occurs rapidly, usually
within 3 days of commencing therapy. Following discontinuation of the therapy, these
bacteria return to pretreatment concentrations within 3 to 4 weeks. The effects of
ciprofloxacin on Staphylococci and Enterococci are not as dramatic or consistent as on
16
the Enterobacteriaceae, although many studies report significant reduction in one or
both of these groups. Generally, ciprofloxacin does not affect the levels of total
anaerobic flora. However, where anaerobic rods, Fusobacterium and Bacteroides
species have been analysed separately, some studies have demonstrated decreases in
faecal levels (Campoli-Richards et al., 1988).
Metronidazole was originally introduced to treat Trichomonas vaginalis, but is now
used for the treatment of anaerobic and protozoal infections. Metronidazole is
bactericidal through toxic metabolites which cause DNA strand breakage.
Metronidazole given orally is absorbed almost completely, with bioavailability of
>90%. Metronidazole is distributed widely and has low protein binding (<20%). The
volume of distribution at steady state in adults is 0.5 to 1.1 L/kg. Metronidazole reaches
60 to 100% of plasma concentrations in most tissues studied, and is extensively
metabolised by the liver into 5 metabolites. The majority of metronidazole and its
metabolites are excreted into urine and faeces, with less than 12% excreted unchanged
into urine (Lamp et al., 1999).
Metronidazole is most active in vitro against gram-negative obligately anaerobic bacilli
such as Bacteroides, including the B. fragilis group and Fusobacterium. (Bergan, 1985).
Although metronidazole and its active metabolites are found in the faeces and also in
colonic mucosa, there is normally little suppression of indigenous colonic flora with
metronidazole therapy. This has been thought at least partly because of the degradative
reduction of the drug by bowel flora under the anaerobic conditions in the colon
(Finegold, 1980). However, studies done with mice have shown that high doses of
metronidazole decreases obligate anaerobes in vivo in the large intestinal flora and this
leads to a consequent increase in certain aerobic species. Brook and Ledney (1994)
found that the mean number of facultative anaerobes rose significantly from day 6
(p<0.05), whereas strict anaerobes fell significantly (p<0.05) during the treatment with
metronidazole compared to controls. In another study metronidazole treatment
selectively eliminated strictly anaerobic bacteria with a concomitant 100-fold increase in
aerobic and facultative bacteria (Wells et al., 1987). Similarly, in the human studies, the
number of Bacteroides species has been shown to decrease and the number of E. coli
and faecal Streptococci to increase in the faeces of patients with Crohn’s disease during
treatment with metronidazole (Krook et al., 1981a). Among healthy human volunteers,
however, the count of Bacteroides species was unchanged at the end of metronidazole
treatment, but there was a significant increase in the faecal Streptococci count (p=0.03)
and an almost significant increase in E. coli (p=0.06) (Krook, 1981). These
dissimilarities in metronidazole’s capacity to reduce human anaerobic faecal flora was
speculated to arise from the higher concentrations of the drug in the large intestine of the
Crohn’s disease patients (Krook et al., 1981b). Taken together, it can be concluded that
metronidazole may dose-dependently increase the number of aerobes in the colonic flora
at the expense of the number of strict anaerobes.
The physiological role and metabolic capacity of the intestinal flora
The number of intestinal bacteria equals (Luckey, 1977) or exceeds (Cummings, 1983)
the number of the cells in their human host. Because of the sparseness of the flora in the
proximal GI tract, its metabolic activity is insignificant compared with those of the
colonic flora. The accepted functions of the colon include the conservation of water and
electrolytes and the controlled evacuation of faeces (Moran and Jackson, 1992). The
17
colon is, however, an important organ of its own with an influence on overall
metabolism, and the effect may in large part be attributed to the activity of colonic
microflora (Phillips, 1984). Its metabolic capacity has been estimated to be at least as
great as that of the liver (Bingham, 1988) or even to exceed that of the rest of the human
body (Luckey, 1977). Intestinal bacteria also have a short generation time and can
undergo enzyme induction when exposed to high levels of substrate. This allows the
microflora to adjust itself rapidly to any change in the environment (Gorbach and
Goldin, 1990).
One of the most important features is its protective function against pathogenic
microbes. Antibiotic treatment may select resistant species or strains in the intestinal
flora and lead to superinfections such as colitis caused by Clostridium difficile.
Colonisation resistance by the normal intestinal flora is thus an important host defence
mechanism. Bacteria are also the main source of antigenic materials and the intestinal
flora is the most important stimulant of the body’s defence mechanisms (Gustafsson,
1982).
As stated earlier, the colonic microflora is predominantly anaerobic, and able to ferment
carbohydrates. The main end-products of this bacterial fermentation are short-chain fatty
acids (SCFAs). The carbohydrate available for fermentation in the colon comes from
endogenous sources such as mucus, and exogenous dietary sources that escape digestion
in the small intestine. The main SCFAs are acetate, butyrate and propionate, all of
which have been shown to stimulate the growth and well-being of the colonic mucosa.
Removal of fibre from the diet results in atrophy of the mucosa. This can be reversed by
the infusion of SCFAs into the colon. Production of SCFAs lowers colonic pH and
increases colonic motility. Moreover, since SCFAs stimulate colonic mucosal blood
flow and oxygen uptake, there is evidence suggesting that bacterial fermentation is
directly involved in colonic mucosal function and also more generally in mucosal
metabolism (Cummings and Macfarlane, 1991).
Bile acids are produced in the liver as end-products of cholesterol metabolism and
excreted into the bile as conjugates. In the intestine the primary bile acids are attacked
by microbial enzymes and transformed into a variety of metabolites, which may be
absorbed and further transformed by liver enzymes prior to their re-excretion into the
bile, forming the enterohepatic circulation of bile acids (Cummings, 1975). Intestinal
microbes may also participate to a lesser extent in heme metabolism, the end product of
which, bilirubin, is hydrolysed/deconjugated in the intestine by microbial and mucosal
enzymes. The deconjugated bilirubin is reduced by microbial enzymes into a complex
mixture of urobilinogens, which are excreted with the faeces. Some of these are also
absorbed from the large intestine and reexcreted into the bile and urine (Gustafsson,
1982). The intestinal microflora also metabolise sterols and steroid hormones. The
steroid hormone metabolites reaching the intestine via the bile are usually conjugated
with sulphuric acid or glucuronic acid. These conjugates are split by the intestinal
microflora and the resulting free steroids are further degraded by the gut bacteria
(Gustafsson, 1982).
Bacterial enzymes play important roles in the metabolism of many drugs, often
determining their bioavailability. For example, in 10% of patients given digoxin the
drug is converted to inactive moieties by the gut flora (Lindenbaum et al., 1981).
Bacterial metabolism may also be relevant to the biological effects of some drugs. An
18
example of this is a salicylazosulphapyridine, which is a complex drug containing an
azo link between a sulphonamide and a salicylate. The two moieties, linked by the azo
bond which is resistant to mammalian enzymes, constitute a large molecule which is not
absorbed in the small bowel. This allows the drug to reach the colon, where bacterial
enzymes hydrolyse the azo bond, releasing sulphapyridine and salicylic acid. Since the
components are thought to act therapeutically on the colon and then to be absorbed
(Phillips, 1984), a symbiosis between colonic bacterial enzymes and a therapeutic effect
is clear.
The above examples show that the colonic flora has several physiological functions.
Because any compound taken orally or entering the intestine via the biliary tract or
blood stream is a potential substrate for bacterial transformation, bacterial flora with its
enzymes is also very likely to be involved in many metabolic processes of foreign
compounds and also exogenous and even endogenous ethanol (Goldin, 1990; Salaspuro,
1996, 1997; Simon and Gorbach, 1984).
Species differences in the intestinal flora
Since conventional laboratory rats are widely used for studies on the metabolism of
intestinal flora, in extrapolating the results obtained from the animal experiments to
human subjects, it is important to know possible differences between the human and rat
intestinal flora. The most notable differences lie in the upper regions of the gut. In man
the stomach and duodenum normally harbour only transient flora, but these areas in rats
are colonised by a mixed bacterial populations of about 107 - 108 organisms/g. The
explanation for this is probably the bactericidal action of the strongly acidic gastric juice
of the humans, whereas the pH of the rat stomach is more moderate (ph 4-5). The large
intestine of rats is colonized by 103 to 10 5times higher concentrations of bacteria than
the small intestine, and the colonic concentration of bacteria is equal to humans. Despite
this, some differences have also been found in the activities of the reductive enzymes
associated with the caecal and faecal floras of rats and humans. For example certain
nitro compounds such as nitrobenzenes and dinitrotoluenes which depend on reduction
by the gut flora for their toxic effects, can be much more potent in rats than in humans,
who have much lower bacterial nitroreductase activity than the laboratory animals
(Rowland, 1986).
Taken together, large intestinal microflora in humans and rats are quantitatively
comparable. Because much less is known about enzymatic similarities of the flora,
caution must be taken in extrapolating results in this field. However, it may be possible
to increase the degree of similarity in gut flora metabolism between laboratory rats and
man by modifying the animal diet (Rumney and Rowland, 1992).
2.3. DISTRIBUTION OF ETHANOL IN THE BODY
Ethanol is absorbed by simple diffusion from the gastrointestinal tract because of its
small size, good water solubility, and low solubility in lipids (Wallgren and Barry III,
1970). No transport processes exist for ethanol (Crabb et al., 1987). Most of the ingested
ethanol, 70-80%, is absorbed by the proximal small intestine, duodenum and upper
jejunum. Slow diffusion from the stomach means that only about 20% of the oral dose is
absorbed from the ventricle. After absorption, ethanol reaches the liver via the portal
19
vein (Riveros-Rosas et al., 1997). The rate of absorption is decreased by delayed gastric
emptying (Oneta et al., 1998). Because gastric emptying is slow and prolonged with
food in the stomach, drinking ethanol after eating a meal, regardless of the nutritional
composition, delays its absorption. This produces a slower rise and lower peak value of
the blood alcohol in fed than in fasting subjects (Jones et al., 1997). The concentration
of the ingested ethanol also influences its absorption, at least when taken after a meal. It
has been shown that postprandially taken high concentrations of ethanol result in lower
blood alcohol levels than do dilute solutions, probably because of delayed gastric
emptying (Roine et al., 1991,1993; Roine 2000).
Once ethanol reaches the blood, it is distributed rapidly throughout the body fluids. In
organs with dense vascularization and rich blood supply, such as brain, lungs, and liver,
alcohol rapidly equilibrates with the blood. In contrast, the distribution of alcohol to the
resting skeletal muscle is particularly slow because of the low number of functioning
capillaries (Agarwal and Goedde, 1990; Dundee et al., 1971). Poor lipid solubility
allows tissue lipids to take up only 4% of the amount of alcohol that can be dissolved in
a corresponding volume of water. Women, who have a smaller total water volume in the
body than men, thus reach higher blood ethanol levels even if both sexes ingest identical
quantities of ethanol (Riveros-Rosas et al., 1997). Distribution of alcohol is mainly
related to the water content of various organs and tissues, so that, for instance, ethanol
concentrations in the terminal ileum are approximately equal to those of the blood
(Halsted et al., 1973). Their high water content makes the ethanol concentration in
saliva (Jones, 1979) and urine (Bendtsen et al., 1999) slightly higher than that in the
blood.
Most of the ethanol (90-95%) is metabolised by oxidation and excreted as CO2 and
water. Other routes for elimination are urine, sweat, and breath. Since alcohol is not
concentrated in the urine or sweat, only negligible amounts of ethanol are excreted via
urine (0.3%), and sweat (0.1%). In humans under normal conditions 0.7% can be
eliminated through the lungs (Holford, 1987).
2.4. HEPATIC ETHANOL AND ACETALDEHYDE METABOLISM
It is generally agreed that the liver is the main organ responsible for the oxidation of
ethanol. Estimations of the contribution of the liver to ethanol elimination under normal
conditions vary from 75-90% (Agarwal and Goedde, 1990). However, in severe hepatic
cirrhosis extrahepatic elimination of ethanol has been shown to account for up to 40%
(Utne and Winkler, 1980). There are three metabolic systems capable of carrying out
ethanol oxidation in the liver: cytosolic alcohol dehydrogenase, the microsomal ethanol
oxidizing system located in microsomes, and catalase, located on peroxisomes. All these
hepatic enzymes yield acetaldehyde as an end-product. Acetaldehyde is further
converted to acetate, mainly in the mitochondria catalysed by aldehyde dehydrogenase.
Alcohol dehydrogenase (ADH)
ADH catalyses the reversible oxidation of many alcohols to corresponding aldehydes. In
case of ethanol, the reaction is as follows:
CH3CH2OH + NAD+ ⇔ CH3CHO + NADH + H+
20
Alcohol dehydrogenase is abundant in the liver and its physiological role has been
postulated to be the degradation of the low levels of alcohol produced by microbial
fermentation in the gut. Another possible explanation is its role in the dehydrogenation
of endogenous steroids (Krebs and Perkins, 1970).
ADH is the main enzyme responsible for the oxidation of ingested ethanol in the liver. It
is a dimer composed of approximately 40 kDa subunits and contains 2 zinc atoms per
subunit. Six different classes of alcohol dehydrogenase have been described for
mammalians; for humans, classes I-V and for rats, classes I-IV and class VI have been
described (Jörnvall and Höög, 1995). In humans class I isoenzymes are coded by three
genes (ADH1 to 3), and are explained by the presence of three protein subunits. In rats,
class I ADH is encoded by one gene. Just recently a recommendation for a new
nomenclature for expressing ADHs has been made. Five human ADH classes should be
expressed by an Arabic number as follows: ADH1, ADH2, ADH3, ADH4, and ADH5.
For genes, the italicized root symbol “ADH” for human and “Adh” for rat, followed by
the appropriate Arabic number for the class; i.e. ADH1 or Adh1 for class I ADH genes
has been recommended. Where multiple isoenzymes exist within a class, adding a
capital letter after the Arabic number; i.e. ADH1A, ADH1B, and ADH1C for human
class I ADHs was also suggested (Duester et al., 1999). In the literature the
nomenclature has been confusing, and misconceptions about class distinction have
frequently occurred. Since most papers have used the “older” nomenclature which codes
classes with Roman numbers, this will be followed here.
Class I ADHs, the classic liver alcohol dehydrogenases, are the most important enzymes
in hepatic elimination of ethanol. These enzymes have a low Km ( 1 mM) and high
Vmax for ethanol, and are responsible for the bulk of ethanol oxidation. This means that
ethanol is effectively eliminated from the blood at a constant rate to very low
concentrations, provided that acetaldehyde is also effectively removed. As the Km of
ADH for acetaldehyde is 0.6 mM it could act as a substrate in the reverse reaction (Blair
and Vallee, 1966). However, the rapid transformation of acetaldehyde to acetate keeps
the reaction in the forward direction. When alcohol is oxidized to acetaldehyde via
ADH, nicotinamide adenine dinucleotide (NAD) is reduced to NADH. Normally the
rate of NADH production exceeds its rate of reoxidation, resulting in an increase in the
liver NADH/NAD ratio. This means that the redox state of the liver is markedly
reduced. Most of the acute metabolic effects of ethanol, such as the inhibition of hepatic
gluconeogenesis, the decrease in citric acid cycle activity and the impairment of fatty
acid oxidation, arise from this major effect of ethanol on the intermediary metabolism of
the liver (Lieber, 1994).
The microsomal ethanol oxidizing system (MEOS)
The first indication of a possible interaction between ethanol and the endoplasmic
reticulum or microsomal fraction of the hepatocyte was provided by the observations
that ethanol feeding results in a proliferation of the smooth endoplasmic reticulum
(SER) in rats and human (Iseri et al., 1966; Lane and Lieber, 1966). This led Lieber and
DeCarli (1968) to find the cytochrome P450-dependent system, which oxidizes ethanol
to acetaldehyde as follows:
CH3CH2OH + NADPH+ + H + O2 ⇒ CH3CHO + NADP+ + 2H2O
21
In humans, the cytochrome fraction responsible for ethanol oxidation has been
designated as CYP2E1. This isoenzyme is the major contributor to the MEOS in
humans, although later studies have suggested that other CYP forms may also play a
role (Asai et al., 1996; Niemelä et al., 1999). The term MEOS should thus be
maintained in referring to the overall capacity of the microsomes to oxidize ethanol
rather than to that fraction of the activity specifically catalysed by 2E1 (Lieber, 1997).
Since the Km of MEOS for ethanol is 7-10 mM, it contributes to ethanol elimination at
high blood ethanol levels. This explains the fact that ethanol metabolism increases with
rising ethanol concentrations above the level needed to fully saturate the low Km
hepatic ADH (Lieber, 1997). The contribution of the MEOS to total ethanol elimination
has not yet been fully clarified. Because of the slow turnover it may be limited, and it
has been estimated that only a minor part (1-5%) of all ethanol metabolism in vivo is
carried out by the MEOS (Ingelman-Sundberg, 1997). The most significant role of
CYP2E1 is, however, its adaptive response to constantly high blood ethanol levels. This
accounts for the metabolic adaptation to high concentrations of ethanol and the
acceleration of ethanol metabolism resulting from chronic alcohol consumption. This
metabolic adaptation has to be distinguished from the central nervous system adaptation
to ethanol which results from chronic ethanol consumption, characterized by the
progressive resistance of the brain to the effects of ethanol (Lieber, 1999).
In addition to acetaldehyde production during ethanol oxidation, the MEOS has been
shown to produce reactive oxygen intermediates, such as superoxide radicals. This may
lead to enhanced lipid peroxidation, so that MEOS may contribute to alcoholic liver
disease. Moreover, CYP2E1 has a capacity to activate over 80 toxicologically important
xenobiotics to potentially hepatotoxic or carcinogenic products (Lieber, 1997).
Catalase
Catalase, which is located in peroxisomes, can oxidize ethanol to acetaldehyde when
hydrogen peroxide is available as follows:
CH3CH2OH + H2O2 ⇒ CH3CHO + 2H2O
However, studies using the catalase inhibitor aminotriazole have shown that this
compound does not slow ethanol metabolism in vivo (Teschke et al., 1976), and
examination of the enzymatic reaction has suggested that the activity of catalase is
limited in vivo by the bioavailability of hydrogen peroxide. Since the rate of hydrogen
peroxide production in the liver is rather low (Boveris et al, 1972), there is some notion
that catalase plays only a minor role in hepatic ethanol metabolism (less than 2%).
Aldehyde dehydrogenase (ALDH)
Regardless of the pathway by which ethanol is oxidised, acetaldehyde is the first
metabolic product. Acetaldehyde is far more toxic than its parent compound ethanol.
Fortunately, it is usually quickly further metabolised to acetate in the oxidative reaction
catalysed by aldehyde dehydrogenase. The liver is the key organ for ethanol oxidation
and the bulk of the ALDHs exist there. Human hepatic aldehyde dehydrogenase activity
can be found in the mitochondria and cytosol (Agarwal, 1997). The main isoenzyme
22
responsible for the oxidation of acetaldehyde, both in humans and rats, has been shown
to be the mitochondrial class 2 ALDH (ALDH2). This has a low Km (3 μM or less) and
a high affinity for acetaldehyde (Lands, 1998). At high acetaldehyde concentrations, the
increase in acetaldehyde oxidation is due to the activity of extramitochondrial ALDH,
mainly cytosolic class 1 ALDH (ALDH1), which has a relatively high Km for
acetaldehyde (100 μM) (Crabb et al., 1987). The low steady-state acetaldehyde
concentration in the liver during alcohol metabolism (<10 μM) suggests that the
mitochondrial isoenzyme is the main form responsible for the oxidation of acetaldehyde
(Crabb et al., 1987). This is evidenced by experiments which show that NADH
generated by the ALDH reaction appears almost exclusively in the mitochondria
(Forsander, 1970).
The central role of the low Km mitochondrial ALDH in acetaldehyde metabolism is
strongly indicated by the finding that a mutation in the ALDH2 enzyme in humans
results in impairment of the ability to dispose of acetaldehyde after ethanol ingestion.
ALDH2 isoenzyme has two allelic forms; the active ALDH2*1 and the relatively
inactive ALDH2*2. Patients homozygous for the ALDH2*2 allele lack ALDH2 activity,
while patients heterozygous for this allele exhibit approximately half the activity of
ALDH2*1 homozygotes (Crabb et al., 1989). Deficient ALDH2 isoenzyme has been
found in about 50% of Japanese (Goedde et al., 1979). The homozygous form of
inactive ALDH2 offers full protection against alcoholism. This has been proposed to be
due to the accumulation of acetaldehyde in the blood during alcohol metabolism, which
causes aversive sensations (Peng et al., 1999). Heterozygotic subjects with about half
the normal ALDH2 activity can, however, drink alcohol or develop even alcohol
dependency (Wall et al., 1992). After ethanol intake the heterozygotic subjects show
flushing, palpitations and nausea, which are caused by elevated blood acetaldehyde
levels (Wall et al., 1992). Blood acetaldehyde levels in heterozygotic subjects have been
shown to be between 8 and 24 μM even after a very low dose (0.1 g/kg of body weight)
of ethanol (Enomoto et al., 1991). In contrast, normal healthy subjects have very low
levels (< 0.5 μM) of acetaldehyde in the peripheral blood during ethanol oxidation
(Eriksson and Fukunaga, 1993). This indicates that in normal healthy individuals almost
all the acetaldehyde formed is effectively oxidised in the liver. Heavy drinkers with the
heterozygous ALDH2*2 genotype (ALDH2*1/2*2) can be considered as human
“knock-out models” for deficient acetaldehyde removal. Consequently, the toxicity of
acetaldehyde is strongly corroborated by the fact that individuals with the heterozygous
ALDH2*2 genotype are at higher risk of developing alcohol abuse-related GI-tract
cancer as compared to those with the normal ALDH2 genotype (Yokoyama et al., 1998).
The end-product of hepatic ethanol oxidation, acetate, leaves the liver via hepatic
venous blood, and is almost completely converted to CO2 and H2O in the peripheral
tissues, mainly in the muscles.
2.5. METABOLISM OF ETHANOL AND ACETALDEHYDE IN THE
DIGESTIVE TRACT
Ethanol metabolism
Although the liver is the major organ responsible for ethanol metabolism, such
metabolism also occurs in the digestive tract. Intestinal metabolism of ethanol is of
23
considerable importance, since it may affect the systemic availability of alcohol and lead
to local production of acetaldehyde, possibly resulting in tissue injury.
Immunohistochemical studies have revealed that alcohol dehydrogenase can be detected
in the mucosa of all parts of the gastrointestinal tract. Furthermore, the amount of ADH
observed was higher in epithelial cells exposed to the lumen than in cells at the bottom
of crypts (Pestalozzi et al., 1983). In addition to their localization, the contribution of
the various ADH isoenzymes to ethanol metabolism depends on their kinetic
parameters, particularly their Km values.
Class IV ADH is characteristic of the upper GI tract, including the mouth and the
esophagus (Dong et al., 1996; Yin et al., 1993). The Km values of the gingival ADH
was estimated to be 27 mM (Dong et al., 1996), and of esophageal class IV 12 mM (Yin
et al., 1993) These high Km values suggest that ethanol may be significantly
metabolised in these tissues. Moreover, the esophagus is the organ of highest ADH
activity in the GI tract, with a rate per mg of protein similar to that of the liver and
approximately 4 times that of the stomach enzyme (Parés and Farrés, 1996).
ADH was detected in human gastric mucosa decades ago (Hempel and Pietruszko,
1979; Smith et al., 1972) and has been shown to exhibit multiple isoenzymes. Class IV
ADH and class I ADH coexist in the stomach, with Km values of 40 and approximately
1 mM respectively (Parés et al., 1992; Seitz and Oneta, 1998; Yin et al., 1997). Since
the stomach contains both high Km class IV ADH and low Km class I ADH, this organ
is a transition site for switching expression of class IV ADH to class I ADH, which is
predominant in the rest of the intestinal tract (Yin et al., 1997). The significance of
gastric ADH is its suggested role in the first pass metabolism of ethanol (FPM). The
gastric FPM of ethanol has been used to explain the differences in the areas under the
ethanol concentration-time curves (AUC) obtained after oral and intravenous alcohol
application. However, the relative contribution of gastric and hepatic metabolism to
FPM is still a subject of debate. Some studies suggest that the differences in AUCs may
be due to the differences in ethanol absorption, and therefore speculate that gastric
ethanol metabolism in rats is negligible and that there is no evidence for this
phenomenon in humans (Smith et al., 1992). By contrast, there are studies indicating a
significant role for gastric ethanol metabolism in the FPM (Caballeria et al., 1987; Lim
et al., 1993). The estimations of FPM of ethanol to ethanol metabolism range between
1% and 20% (Seitz and Pöschl, 1997).
The small and large intestinal ADH is mainly composed of class I ADH, with a Km of
1-2 mM for ethanol (Seitz and Oneta, 1998). The activity of rectal ADH was
comparable to gastric ADH activity and, compared to ADH activities in other colonic
segments, was found to be significantly higher (Seitz et al., 1996). This suggests that
ethanol may be effectively metabolised to acetaldehyde in the colonic mucosa and
especially in the rectal mucosa.
In addition to ADH, rat gastric mucosa have been shown to possess catalase activity
(Salmela et al., 1996), but its presence in the rest of the alimentary tract is unknown.
Moreover, immunohistochemistry has revealed that rat duodenal and jejunal villous
cells exhibit CYP2E1 activity, but it is not expressed or induced in the stomach, ileum,
colon and rectum (Shimizu et al., 1990).
24
Acetaldehyde metabolism
Cytosolic high Km ALDH3 is the only ALDH isoenzyme identified in the human mouth
thus far. Both gingival and tongue tissue ALDH exhibit significant amounts of enzyme
activity at 20 mM acetaldehyde (Dong et al., 1996). Regarding human gastric mucosa,
ALDH3 has been estimated to account for more than 80% of the ALDH activity.
Although ALDH3 has a high Km value for acetaldehyde (approximately 88 mM), it has
been suggested that acetaldehyde generated by gastric ADH could be oxidized in the
same tissue (Parés and Farrés, 1996; Yin et al., 1997). A different picture is seen in the
esophagus, where the ALDH3 activity is 5 times below that of the gastric mucosa, while
the rate of acetaldehyde production is higher than in the stomach (Yin et al., 1993).
Accumulation of acetaldehyde could thus occur in esophageal tissue, contributing to the
alcohol-related end-organ damage. ALDHs classes 1 and 2 can be found in the human
duodenum (Liao et al., 1991), but much less is known about the ALDH activity of the
rest of the small intestine. ALDHs 1, 2, and 3 have been observed in human and rat
colonic mucosa, but the expression of low-Km mitochondrial ALDH2 seems to be
particularly low (Koivisto and Salaspuro, 1996; Yin et al., 1994). The activity ALDH in
the colonic mucosa was only slightly lower in ALDH2-deficient subjects than in normal
phenotype carriers, suggesting that mucosal ALDH2 plays only a minor role in the
oxidation of acetaldehyde in the colon (Yin et al., 1994).
2.6. MICROBIAL ETHANOL FERMENTATION AND OXIDATION
Endogenous ethanol
Measurable amounts of ethanol are normally formed in the gastrointestinal tract. In the
caecum of normally fed rats, the mean ethanol concentration has been shown to be 0.9
mM. This endogenous alcohol, which derives from anaerobic degradation of glucose to
ethanol by some microorganisms, is absorbed into the portal circulation and almost
quantitatively removed by the liver (Krebs and Perkins, 1970). Conditions associated
with bacterial overgrowth producing markedly elevated endogenous ethanol levels
detected even in the blood indicate intestinal microbial-derived ethanol production in
humans. For example, in some patients after a jejunoileal bypass operation, detectable
fasting serum concentrations of ethanol up to 1 mM have been noted. In the past this
kind of operation resulted in blind-loop formation and consequent colonisation and
bacterial overgrowth of that segment (Mezey et al., 1975). Additionally, midjejunal
aspirates from the patients with tropical sprue show not only an overgrowth of
coliformic bacteria (Klebsiella pneumoniae, Enterobacter cloacae and Escherichia coli)
but also large quantities of their fermentation end-product ethanol with concentrations
from 2 mM to 31 mM (Klipstein et al., 1973). Small amounts of endogenous ethanol (1
to 27 mM) have also been found in the gastric juice of patients receiving cimetidine or
antacids. Bode et al. (1984a) speculated that this resulted from an increase in the yeast
and/or bacterial population in the stomach due to the reduction of gastric acid induced
by the medication.
The last step in bacterial alcoholic fermentation is the reduction of acetaldehyde to
ethanol, catalysed by bacterial ADH (Reid and Fewson, 1994; Tamm, 1974). This has
been described in full detail in the case of Escherichia coli (Clark, 1989; Dawes and
Foster, 1956; Still, 1940; Wong and Barrett, 1983), group N Streptococci (Lees and
25
Jago, 1976) as well as within the Enterobacteriaceae in general (Salveson and Bergan,
1981). As already stated, this reaction is also reversible in microorganisms (Maconi et
al., 1988). This means that if oxygen is present and given an excess of ethanol, the ADH
mediated reaction can also run in the opposite direction and ethanol may be oxidized to
acetaldehyde in a reaction in which nicotinamide adenine dinucleotide acts as an
electron acceptor.
Microbial acetaldehyde production in vitro
As mentioned earlier, E. coli was shown to possess ADH activity in 1940 by Still.
Jokelainen et al. (1996a) showed that different strains of faecal E. coli and other Gramnegative
rods, mainly belonging to the Enterobacteriaceae family, were able to produce
significant amounts of acetaldehyde when incubated aerobically in vitro with ethanol in
reaction catalysed by NAD-dependent ADH. Later studies have shown that E. coli is
also able to oxidize ethanol in microaerobic conditions, i.e. when 6% O2 is present
(Salaspuro et al., 1999). This is important, since the PO2 in the colonic mucosa is
known to be approximately equal to that of venous blood, and hence the conditions in
the colon are more or less microaerobic (Hamilton et al., 1968). It was shown earlier
that human colonic contents, i.e. the mixture of colonic bacteria, can produce
acetaldehyde from ethanol in a dose-dependent manner in vitro (Jokelainen et al., 1994).
The significant acetaldehyde production occurred at the ethanol concentrations that are
relevant to in vivo conditions. These findings have opened up a new microbiological
approach to researching acetaldehyde production and its pathological consequences in
the gastrointestinal tract.
Regarding the upper digestive tract, in vitro production of acetaldehyde has been
reported when human mouth- and bronchopulmonary washings were incubated
aerobically with ethanol. Since acetaldehyde production could be abolished by
pretreatment with antibiotics, acetaldehyde formation from ethanol was thought to be of
bacterial origin (Jauhonen et al., 1982; Miyakawa et al., 1986; Pikkarainen et al., 1981).
Interestingly, in vitro acetaldehyde production by mouthwashings from oropharyngeal
cancer patients has been shown to be significantly higher than that of the control
patients (Jokelainen et al., 1996b). This suggested that microbially derived acetaldehyde
production could be involved in ethanol-associated organ toxicity.
Helicobacter pylori also exhibits significant cytosolic ADH activity. Consequently, H.
pylori has been shown to produce acetaldehyde when intact cells are incubated with
ethanol, and this has been speculated to contribute to H. pylori-associated
gastroduodenal injury (Roine et al., 1992, 1995; Salmela et al., 1993, 1994).
Microbial acetaldehyde production in vivo
Regarding the human oral cavity, the production of marked amounts of acetaldehyde up
to 140 μM in saliva after the ingestion of a moderate amount of ethanol has been
demonstrated. This acetaldehyde production was significantly reduced by the use of
antiseptic chlorhexidine mouthwash, indicating its microbial origin (Homann et al.,
1997a). Another important finding in the same study was that in vivo salivary
acetaldehyde levels correlated highly significantly with the acetaldehyde levels produced
in vitro. This offers an opportunity to use the in vitro salivary test to reliably
differentiate high acetaldehyde producers from low producers.
26
Intestinal in vivo microbially derived ethanol oxidation and acetaldehyde production
have been shown to occur in rats with a jejunal self-filling diverticulum and
concomitant bacterial overgrowth (Baraona et al., 1986). Moreover, after an acute dose
of ethanol to rats, the rectal mucosal acetaldehyde concentration was significantly higher
in conventional rats than germ-free rats, indicating the role of microbes in acetaldehyde
production (Seitz et al., 1990). Ethanol oxidation to acetaldehyde has also been detected
in the caecum of anaesthetized pigs after intragastric or intravenous ethanol
administration (Jokelainen et al., 1996c). Furthermore, high intracolonic acetaldehyde
levels have been detected in the caecal samples of rats after intraperitoneal ethanol
administration, and these levels were effectively suppressed by pre-treatment with
ciprofloxacin, which reduces the amount of aerobic bacteria in the large intestine
(Visapää et al., 1998). These findings strongly suggest that microbes of the large
intestine are able to oxidize ethanol by a bacteriocolonic pathway for ethanol oxidation.
In a very recent human study, we also showed ethanol-derived acetaldehyde production
in the stomach after treatment with lansoprazole. Medication led to marked bacterial
overgrowth in the neutral stomach, and this was considered to be the most probable
explanation for the enhanced acetaldehyde production detected (Väkeväinen et al.,
2000).
Microbial acetaldehyde metabolism
Aldehyde dehydrogenase activity has been identified in the cytosol of yeasts (Steinman
and Jakoby, 1968) and in anaerobic bacteria (Burdette and Zeikus, 1994). ALDH
activity also exists in facultative anaerobic bacteria, which are able to produce
acetaldehyde via ADH. Escherichia coli has been shown to exhibit aldehyde
dehydrogenase activity (Dawes and Foster, 1956; Wong and Barrett, 1983). Nosova et
al. (1996) studied 27 different bacterial species, mainly belonging to the family
Enterobacteriaceae, and showed them to be capable of cytosolic NAD or NADP-linked
ALDH activity. However, based on kinetic studies, the ability of bacterial ALDHs to
oxidise higher concentrations of acetaldehyde was rather low (Nosova et al., 1998).
Taken together with low ALDH activity in the colonic mucosa (Koivisto and Salaspuro,
1996), this offers an additional explanation for the mechanism of the accumulation of
ethanol-derived acetaldehyde in the large intestine.
2.7. ALCOHOL AND THE ALIMENTARY TRACT
Symptoms and functional changes associated with alcohol use.
Excessive alcohol consumption is frequently associated with gastrointestinal symptoms.
Although most symptoms after both acute and chronic use of alcohol are reversible and
disappear during abstinence, chronic use of alcohol may cause structural or functional
changes that can lead to permanent gastrointestinal diseases. An example of this is the
increased risk of colorectal adenomatous polyps, which are generally regarded as
premalignant lesions (Cope et al., 1991). Alcohol consumption and the risk of colorectal
cancer will be discussed in the next chapter.
Only the upper digestive tract, mouth, esophagus, stomach and upper small intestine are
27
directly affected by the ingested ethanol. However, since ethanol is transported by blood
circulation to other organs ethanol concentrations in the terminal ileum (Halsted et al.,
1973) and colon (Levitt et al., 1982) are approximately equal to those of the blood. A
questionnaire showed that actively drinking alcoholics more frequently report heartburn,
nausea, vomiting, diarrhea, and flatulence than the matched controls. These symptoms
were transient and resolved after two weeks of sobriety. An important note of the study
was that these symptoms were associated with active alcohol use and were not
withdrawal symptoms (Fields et al., 1994).
The etiology for the increased incidence of GI symptoms in alcoholics is probably
multifactorial. Moreover, prolonged use of alcohol may have effects which differ from
the acute effects of a single dose. Some studies suggest that acute alcohol ingestion may
impair the function of the lower esophageal spinchter and reduce the incidence of
primary peristalsis in the distal esophagus (Hogan et al., 1972; Mayer et al., 1978),
although some others have reported the reverse (Keshavarzian et al., 1987; Silver et al.,
1986). However, there is evidence that acute alcohol intake increases esophageal reflux
episodes and impaires the acid clearance of the esophagus in the supine position
(Kaufman and Kaye, 1978; Vitale et al., 1987). This may explain the increased
incidence of heartburn in heavy alcohol users.
Acute ingestion of alcohol has been shown to produce inflammatory changes in the
stomach and duodenum of humans. The most prominent findings included
haemorrhagic erosions, subepithelial blebs and infiltration of inflammatory cells into the
lamina propria (Gottfried at al., 1978). Rubin et al. (1972) showed that after sustained
use of ethanol, the histology of the human small intestine was normal when examined
by normal light microscopy, but morphometric and electron microscopic examinations
demonstrated alterations in the cell organelles of the mucosa. Others have, however,
shown reduction in the villus height and a reduced mucosal surface area of villi in the
small intestine of actively-drinking alcoholics (Bode et al., 1982a; Persson, 1991; Seitz
et al., 1985). Several enzymes known to be located in the absorptive cells may also be
affected by ethanol. Reduced activity in disaccharidases has been noted after sustained
high ethanol administration (Bode et al., 1982b). Decreased lactase activity can
contribute to milk intolerance with diarrhea commonly observed in alcoholics after a
period of heavy drinking (Keshavarzian et al., 1986; Persson, 1991). Moreover, oralcaecal
time has been shown to be decreased in recently drinking alcoholics, suggesting
that the increasing transit of intestinal contents may contribute to the diarrhea commonly
seen during and after an episode of heavy drinking (Keshavarzian et al., 1986). Chronic
alcohol use has also been shown to induce both histological and ultrastructural changes
in the human rectal mucosa. These reversible changes include inflammatory changes, a
decreased number of goblet cells, and alterations in the cell organelles (Brozinsky et al.,
1978).
The nutritional status of advanced alcoholics is often poor, at least among lower-income
and homeless alcoholic populations. The etiology of malnutrition has been generally
thought to be multifactorial (Salaspuro, 1993). On average, ethanol accounts for about a
half of an alcoholic’s caloric intake. It therefore displaces normal nutrients, causing
malnutrition (Lieber, 1995). Secondary malnutrition also occurs through malabsorption
due to the structural and functional changes in the small intestine, but pancreatic
exocrine insufficiency, decreased biliary secretion and impaired hepatic metabolism of
nutrients may also be involved (Lieber, 1995; Salaspuro, 1993).
28
Both acute and chronic exposure to ethanol has been shown to increase intestinal
permeability to macromolecules in experimental animals. Increased small intestinal
permeability has also been observed in human alcoholic patients without liver cirrhosis.
This “leaky gut” phenomenon was shown to persist up to 2 weeks after cessation of
drinking (Bjarnason et al., 1984), and seems to result from a temporary destabilization
of intracellular junctions (Draper et al., 1983). Interestingly, high acetaldehyde levels
have been shown to reversibly increase the paracellular permeability of the Caco-2 cell
monolayer (Rao, 1998). Acetaldehyde production by the intestinal bacteria may thus be
one mechanism by which prolonged alcohol use increases intestinal permeability. The
disruption of the mucosal barrier may promote absorption of normally nonabsorbable
compounds like endotoxins into the portal circulation (Bjarnason et al, 1984; Persson,
1991). Moreover, leaky gut has recently been suggested as a possible mechanism for
alcohol-induced liver damage (Keshavarzian et al., 1999).
Alterations in the oral and intestinal flora caused by chronic alcohol intake
There are no studies available to show whether heavy alcohol consumption directly
alters the oral flora. However, although the overall dental health in subjects misusing
alcohol, investigated by the decayed, missing and filled teeth (DMFT) index, has been
shown to be only slightly poorer than national averages in the United Kingdom, a high
incidence of tooth wear and trauma to the dentition was noticed (Harris et al., 1996,
1997). Tooth wear was presumed to be caused by the regurgitation of gastric acid. This
study suggests that the dental hygiene habits of alcohol misusers may be poorer than that
of abstainers or moderate drinkers. Poor dental hygiene habits may favour the
overgrowth of some microbes in the oral flora. Moreover, poor dietary factors and the
suppressive effects of alcohol intake on various arms of the host immune defence may
predispose to colonization by some bacteria and yeasts in the oral cavity (MacGregor,
1986; Oksala, 1990). Additionally, most heavy alcohol drinkers are also heavy cigarette
smokers (Harris et al., 1996). Heavy smoking may be the strongest factor predisposing
to microbial colonisation in the oral cavity since it has been shown to increase the
presence of yeasts and Gram-positive aerobic bacteria there (Colman et al., 1976;
Macgregor 1988; Rindum et al., 1994; Sakki and Knuuttila, 1996). Moreover, heavy
alcohol consumption in humans has been associated with a marked (70%) reversible
reduction in the flow rate of stimulated parotid saliva. Beside this, chronic alcohol abuse
also reduces protein secretion, like amylase and epidermal growth factor (EGF), in
parotid saliva (Dutta et al., 1992). Ohmura and coworkers (1987) suggested that low
EGF levels in saliva may decrease the resistance of the mucosal barrier to chemical
stress. Therefore, low salivary levels of EGF may be one possible mechanism by which
ethanol or acetaldehyde may influence the development of oral lesions associated with
alcohol use.
Marked qualitative and quantitative changes have been observed in the flora of the small
intestine of chronic alcoholics. The most significant finding was the increase in the
number of bacteria per unit volume of jejunal juice, especially in the number of aerobic
and facultative anaerobic Gram-negative rods and obligate anaerobic bacteria (Bode et
al., 1984b). Evidence for an increased incidence of bacterial overgrowth in alcoholics
also stems from studies using the hydrogen breath test after ingestion of lactulose (Bode
et al., 1989). Mucosal bacterial overgrowth has also been detected more often and with
higher microbial counts in the biopsy samples of the stomach and duodenum among
29
alcoholics (Hauge et al., 1997). This upper GI tract bacterial overgrowth has been
speculated to be due to the increased pH of the gastric juice.
Small intestinal bacterial overgrowth may lead to malabsorption or morphological
changes in the small intestinal mucosa (Persson, 1991). Moreover, bacterial overgrowth
might contribute to the increased release of endotoxin and lead to endotoxemia, which is
often detected in patients with alcoholic liver disease (Bode, 1987). Endotoxin, a cellwall
constituent of Gram-negative bacteria, may play a role in alcoholic liver injury by
activating macrophages to release cytokines (McClain et al., 1999). The small intestinal
bacterial overgrowth may thus also contribute to the pathogenesis of extraintestinal
disorders associated with alcohol use. Bacterial overgrowth probably also leads to
increased microbially-derived acetaldehyde production from ethanol in the small
intestine.
2.8. ALCOHOL AND DIGESTIVE TRACT CANCERS
Cancer of the oropharynx and esophagus
There is extensive epidemiological evidence, mainly from case-control studies although
also from cohort studies, showing that tobacco smoking and heavy alcohol consumption
increase the risk of cancers of the mouth, pharynx, esophagus and larynx (Doll et al.,
1999; Franceschi et al., 1990; Grønbæk et al., 1998; Mashberg et al., 1993; Merletti et
al., 1989). In Europe these factors have been estimated to account for about threequarters
of all cases (La Vecchia et al., 1997). Both smoking and alcohol consumption
have been shown to be independent risk factors for upper digestive tract cancers. The
cancer risk increases proportionally with the quantity of cigarettes smoked or alcoholic
beverages drunk. When combined, there is epidemiological evidence indicating that
alcohol and tobacco act together in a multiplicative rather than in an additive manner,
having synergistic tumour-promoting effects (Blot et al., 1988; Brugere et al., 1986; La
Vecchia et al., 1997). This means that the combined effect of both of these agents is
greater than simply adding the effects of each together. Many attempts have been made
to separate the effects of different types of alcoholic beverages in their potential for
carcinogenicity. The consensus, however, is that the main component of alcoholic
beverages determining the risk of cancer is ethanol (Doll et al., 1999). This is further
supported by the fact that the regular use of mouthwashes with a high ethanol content
increases the risk of oropharyngeal cancer (Winn et al., 1991).
Other possible risk factors for upper gastrointestinal tract cancer are poor nutritional
status and intake of micronutrients, hereditary factors, certain papilloma viruses, and
occupational hazards (Bundgaard et al., 1995; Graham et al., 1977; La Vecchia et al.,
1997; Marshall et al., 1992; Marshall and Boyle, 1996). Moreover, dentition, tooth loss,
poor dental status and oral hygiene habits are associated with higher risks, especially for
oral cavity cancer (Bundgaard et al., 1995; Graham et al., 1977; Maier et al., 1993;
Marshall et al., 1992). It is generally agreed that the influence of poor oral hygiene as a
risk factor is much less compelling than alcohol and smoking, but there is some
experimental evidence that its influence might become clinically more important among
alcoholics, whose poor dental status and hygiene may be a common problem (Maier et
al.,1993). The reason for this finding is unclear.
30
Although alcohol and tobacco smoke are well-known independent and strong risk
factors for upper gastrointestinal tract cancer, the exact mechanism by which they exert
an influence upon the oral mucosa is poorly understood. Consequently, the only things
known to reduce the risk of upper digestive tract cancers is to limit alcohol consumption
and to stop or never start using tobacco and, perhaps, to have a regular diet rich in fruit
and green vegetables (Harris, 1997). Many compounds in tobacco smoke are hazardous
to health and some are undoubtedly carcinogenic (IARC, 1986). In contrast, the tumourpromoting
effects of alcohol consumption are less well defined. Moreover, ethanol is
not a carcinogen by standard laboratory tests. As alcohol is involved synergistically in
the attributable risk of cigarette smoking, it has been suggested that unifying
pathogenetic mechanism may underlie these epidemiological findings. The possible
mechanisms by which alcohol may influence the development of cancers are discussed
separately, with special reference to acetaldehyde in chapter 2.9.
Cancer of the stomach
Most epidemiological retrospective cohort studies show no increase in the stomach
cancer rate among alcoholics or heavy drinkers. In these studies cancer incidence in
groups with high alcohol intake was compared with that of the general population. In
follow-up studies of populations with known consumption most studies showed no
elevated risk, with RR from 0.9 to 1.2 (Doll et al., 1999). However, two studies (Gordon
and Kannel, 1984; Kato et al., 1992) reported a significant increase with heavy drinking,
with a relative risk of 3.05 in the study by Kato et al. In that study stomach cancer
mortality was prospectively studied among 9753 Japanese, who frequently show the
presence of partially inactive ALDH 2*2 isoenzyme (Goedde et al., 1979). It has since
been shown that the mutant allele is associated with higher frequency of GI
malignancies, including stomach cancer, if alcohol is consumed regularly (Yokoyama et
al., 1998). This limits the extrapolation of this study to Caucasians with no such enzyme
defect, but indicates the role of acetaldehyde in carcinogenesis associated with alcohol
use. Some case-control studies have shown a significant positive relationship between
alcohol consumption and stomach cancer (RR 1.5 - 1.7), whereas others showed no
significant relationship (Doll et al., 1999). Taken together, epidemiological evidence
between alcohol consumption and cancer of the stomach is far from clear. Moreover,
there has been a dramatic worldwide decline in the incidence of stomach cancer, a
finding in contrast to the general increase in alcohol consumption and alcohol-related
diseases such as cirrhosis of the liver. IARC stated in 1988 that there is little aggregate
data to suggest a causal link between drinking alcoholic beverages and stomach cancer.
However, Doll et al. (1999) later concluded that alcohol may have an etiological role in
the stomach cancer, albeit minor and unproven.
An exception to this may be the cancer of the gastric cardia, the incidence rates of which
have been recently increasing. The cardia region of the stomach is the uppermost area
where the stomach adjoins the esophagus. It has been suggested that this cancer
resembles a specific type of cancer of the lower oesophagus and may share common risk
factors, notably tobacco and alcohol exposure (Vaughan et al., 1995).
Cancer of the large intestine
As with gastric cancer, there is considerable epidemiological data concerning the
possible association between cancer of the large bowel and consumption of alcoholic
31
beverages. Some studies showed no evidence of such an association, while others
showed a statistically significant association. The rest of the studies were somewhere
“in-between” showing no consistent and significant overall association, but the RRs
were elevated in some subgroups (Doll et al., 1999). However, two different metaanalyses
of more than 60 studies between 1951 and 1991 both conclude that alcohol
leads to a small but significantly increased cancer risk, especially for the left colon and
the rectum with an estimated overall RR of 1.1 (95% CI 1.05-1.14) (Kune and Vitetta,
1992; Longnecker et al., 1990). Moreover, just recently a panel of European experts at a
World Health Organization (WHO) Consensus Conference on Nutrition and Colorectal
Cancer declare that alcohol had a causal effect in colorectal carcinogenesis (Scheppach
et al., 1999).
Possible mechanisms of ethanol-induced carcinogenesis
It has been concluded that there is sufficient evidence for the carcinogenicity of
alcoholic beverages in humans (IARC, 1988), but there is no experimental evidence to
indicate that ethanol itself is a carcinogen (Doll et al., 1999). This means that pure
ethanol has not been shown to be carcinogenic in laboratory experiments. In in vitro
studies with human and other mammalian cells it has been shown that ethanol does not
induce DNA damage, sister chromatid exchanges or chromosomal aberration. Most of
the in vivo studies with mice or rats described in the literature cannot be used to evaluate
the carcinogenicity of alcohol due to limitations in experimental design (IARC, 1988).
Animal experiments suggest, however, that ethanol may act as a co-carcinogen in the
production of cancers. This means that it modifies or enhances the carcinogenic
potential of known carcinogens. Grici t et al. (1982, 1984) exposed C57B mice by
gastric intubation to N-nitrosodiethylamine (NDEA) or N-nitrodi-n-propylamine
(NDPA), either in tap water or in a 40% ethanol solution, twice a week for 50 weeks. A
significant increase in the incidence of squamous cell carcinomas of the
esophagus/forestomach was observed in the group given the carcinogens in ethanol as
compared to those given the nitrosamines in the tap water. Seitz et al. (1984) studied the
effect of chronic ethanol administration on 1,2-dimethylhydrazine (DMH) induced rectal
carcinogenesis in male rats fed a nutritionally adequate liquid diet containing 36% of
total energy as ethanol or isocaloric carbohydrate. Sustained ethanol ingestion increased
the number of rectal tumours significantly (17 vs. 6; p<0.02).
Although many hypotheses exist, experimental work has so far failed to elucidate the
underlying mechanisms by which excessive consumption of alcoholic beverages may
act as a co-carcinogen under certain conditions. Some of these hypotheses are listed
below.
Alcohol may contain congeners and other contaminants that may be carcinogenic.
Several substances known or thought to cause cancer in humans have been detected in
alcoholic beverages. Special attention has been paid to N-nitroso compounds, which
have been related to colorectal cancer in humans (Knekt et al., 1999). Several N-nitroso
compounds, e.g. nitrosodimethylamine (NDMA), have been found in higher
concentrations in some beers than in other beverages (Walker et al., 1979). This is a
possible mechanism for a specific carcinogenic effect of beer drinking in relation to
rectal cancer. While some studies have found an elevated risk (Riboli et al., 1991), these
were not confirmed by others (Potter and McMichael, 1986). The consensus is that there
32
is no appreciable and consistent difference in the risk among different types of alcoholic
beverages (Doll et al., 1999).
Alcohol intake generates metabolites which may be carcinogenic to humans, or ethanol
itself may act as a solvent, increasing penetration of other carcinogens into the target
tissue (Wight and Ogden, 1998). The major metabolite of ethanol is acetaldehyde. There
is sufficient evidence for the carcinogenicity of acetaldehyde to experimental animals
(IARC 1985, 1999), and indirect strong epidemiological evidence in humans
(Yokoyama et al., 1998). Because of the central role of acetaldehyde in the studies of
this thesis, this topic will be discussed separately in the next chapter.
Either alcohol intake may reduce the intake and/or bioavailability of nutrients which
could inhibit cancer, or alcohol consumption may enhance nutritional deficiencies that
increase the risk of cancer. In some cases, nutritional deprivation may lead to nutritional
deficiencies that may alter epithelial cell chemistry and function, increasing
susceptibility to carcinogens (Blot, 1992), an example being folate deficiency. In
epidemiological studies, decreased folate status has been associated with an increased
risk of neoplastic transformation in the colon (Giovannucci et al., 1995). The
pathogenetic mechanism of the increased cancer risk in folate deficiency has been partly
elucidated. Folate is a crucial methyl group donor for many transmethylation reactions
in the human body. Diminished folate leads to hypomethylation of the DNA, which has
been observed in several experimental cancer models and in human cancers (Goelz et
al., 1985; Kim et al., 1997). Since high levels of acetaldehyde have been shown to be
able to catabolize folate in vitro (Shaw et al., 1989), there is conjecture that alcohol
intake and low folate together could play a role in colorectal carcinogenesis
(Anonymous, 1994; Boutron-Ruault et al., 1996; Collins et al., 1992; Giovannucci et al.,
1995).
Prolonged alcohol use may also inhibit the detoxification of carcinogenic compounds.
Experimental studies have suggested that the effects of alcohol on the liver may block
hepatic inactivation of carcinogens, thus increasing exposure to these compounds (Blot,
1992). Moreover, heavy alcohol use may catalyse the metabolic activation of some
compounds into carcinogens. For example, alcohol intake induces hepatic CYP2E1,
which also has a unique capacity to activate over 80 toxicologically important
xenobiotics to potentially carcinogenic products (Lieber, 1997). Since oxygen free
radicals are generated during alcohol metabolism, target cells may be exposed to
oxidants and this may lead to activation of carcinogens at the cellular level. Increased
oxidative stress may also increase the risk of DNA damage and malignant
transformation. However, the tumour promotion associated with alcohol and with the
generation of free radicals is in general not very clear and the role played by ethanol in
this process is even less so (Mufti et al., 1993).
Alcohol is also known to cause immunosuppression, a possible contributing factor in
the increased cancer rate among alcoholics. The role of immunosuppression in ethanolassociated
cancers is, however, questionable, since increased incidences of lymphoma,
the tumour most closely associated with depressed immune function, have not been
reported among heavy drinkers (Blot, 1992).
33
2.9. ORGAN TOXICITY OF ACETALDEHYDE
Nearly every organ system in the human body can be affected by heavy and prolonged
use of alcohol. In addition to GI-tract cancers, there is no doubt that the consumption of
alcoholic beverages increases the risk of alcoholic liver diseases. As already mentioned,
ethanol itself is not carcinogenic. Similarly, the fact that only a minor proportion of
alcohol abusers develop the most severe forms of liver damage suggests that other
mechanisms than the direct effect of ethanol in creating tissue toxicity have to exist.
Acetaldehyde, the first metabolite of ethanol, possesses toxic properties that markedly
exceed those of ethanol, and there is increasing evidence suggesting its part in the
detrimental action of alcohol on the digestive tract.
Acetaldehyde-protein adducts
Acetaldehyde has properties that make it very suitable for potential nucleophilic attacks.
Since many nucleophilic groups are present in the proteins, they are natural targets in
various tissues for reactive acetaldehyde. The binding of acetaldehyde with proteins
results in the formation of two major types of product, which have been classified as
unstable and stable acetaldehyde-protein adducts. The unstable adducts may serve as
intermediates in stable adduct formation and can be stabilized by reducing agents such
as NADH, which is found in large amounts in the liver during ethanol metabolism. The
stable adducts appear to be the most likely causes of toxic effects (Nicholls et al., 1992).
In vitro, acetaldehyde has been shown to bind covalently to many cellular and
extracellular proteins (Nicholls et al., 1992). In vivo, it is well established that
acetaldehyde adduct formation occurs in the liver during ethanol oxidation in both
experimental animals and humans (Lin et al., 1988; Niemelä et al., 1991). To localize
acetaldehyde-protein adducts, immunohistochemical studies have been performed with
specific antibodies. Adducts have been demonstrated mainly in the cytoplasm of the
perivenular hepatocytes, where acetaldehyde production is believed to be the highest
(Niemelä et al., 1991, 1994). Moreover, acetaldehyde adducts have been detected in the
areas of active fibrogenesis in the liver biopsy specimens from alcoholic patients
(Holstege et al., 1994). More recently, acetaldehyde adducts have also been
demonstrated in the rough endoplasmic reticulum (RER), and in some peroxisomes of
hepatocytes, as well as in myofibroblasts and Ito cells (Paradis et al., 1996).
Although the ability of acetaldehyde to bind to hepatic proteins during ethanol
metabolism has been well established, the precise role of acetaldehyde-protein adducts
in the pathogenesis of alcoholic liver disease has not been clarified. One mechanism
through which such adducts may be involved in alcoholic liver disease is that
acetaldehyde-protein adducts may be recognized as neoantigens by the immune system.
This may trigger potentially harmful immune responses directed against liver cells
(Nicholls et al., 1992). The presence of circulating antibodies against acetaldehydeprotein
adducts have indeed been described in humans (Israel et al., 1986; Niemelä et
al., 1987). As several observations indicate that immunological mechanisms are
involved in alcoholic liver disease, the appearance of such antibodies against
acetaldehyde-protein adducts may be involved in the development and progression of
liver injury (Tuma and Klassen, 1992).
Acetaldehyde-protein adducts may also alter the biological properties of the modified
34
proteins. In the case of hepatic enzymes, the covalent binding of acetaldehyde has been
shown to lead to the inhibition of the activities of enzymes which contain lysine at the
catalytic site (Mauch et al., 1986). Moreover, long-term ethanol consumption has been
shown to produce a decrease in O6 -methylguanine transferase (O6 -MeGT) activity in
rats in vivo (Garro et al., 1986). This has been shown to be due to the acetaldehydecysteine
adduct in the active site of the enzyme, and it occurs even at nanomolar
acetaldehyde concentrations (Espina et al., 1988). O6 -MeGT is a DNA repair enzyme,
which removes alkyl groups from the O6 position of guanine. Since alkylation at this
position is associated with both mutagenesis and carcinogenesis (Kleihues et al., 1979;
Lewis and Swenberg, 1980; Newbold et al., 1980), the inhibition of O6 -MeGT offers
one explanation of how ethanol may act as a cocarcinogen, enhancing the carcinogenic
potential of other agents.
There is not much evidence about acetaldehyde-protein adducts in the mucosa of the
gastrointestinal tract. One study shows that both exogenous acetaldehyde and that
produced locally from ethanol binds to gastric mucosa. This adduct formation has been
suggested to be a pathogenetic factor behind ethanol-associated gastric injury (Salmela
et al., 1997). No evidence indicates whether acetaldehyde adducts are formed in the
mucosal proteins in the rest of the digestive tract or not. Since microbes produce high
levels of acetaldehyde from ethanol in the oral cavity and the colon, similar adduct
formation at these anatomical sites as in the gastric mucosa might also occur.
Acetaldehyde as a carcinogenic agent
Acetaldehyde has been shown to be a highly mutagenic agent. Specifically, it may
induce chromosomal aberrations, and micronuclei and/or sister chromatid exchanges in
cultured mammalian cells (Dellarco, 1988; IARC, 1985). Moreover, it has been shown
to induce gene mutations in human lymphocytes (He and Lambert, 1990). The induction
of cytogenetic effects has been postulated to be related to the ability of acetaldehyde to
form DNA-DNA and/or DNA-protein cross-links. The mechanisms of these DNA
cross-links caused by acetaldehyde may involve direct attack by acetaldehyde on DNA.
A series of studies has shown that acetaldehyde can react with DNA bases to produce
specific types of base adduct, which is a critical initiating event in the multistage process
of chemical carcinogenesis (Hemminki and Suni, 1984; Vaca et al., 1995). Under
biologically relevant conditions, the most prevalent of these is N2 -ethyldeoxyguanosine
(N2 -Et-dG) (Fang and Vaca, 1995; Vaca et al., 1995).
Fang and Vaca demonstrated that N2 -Et-dG becomes detectable in the liver DNA of
mice treated with 10% ethanol in their drinking water for 5 weeks. This adduct was
undetectable in control mice not given alcohol (Fang and Vaca, 1995). In humans such
adducts have been detected in peripheral white blood cells of alcohol abusers (Fang and
Vaca, 1997). DNA adducts have also been identified in nontumoral colon mucosa of
human patients with colorectal cancer (Pfohl-Leszkowicz et al., 1995). Although these
DNA adducts were unspecified and not necessarily related to acetaldehyde, this study
indicates that covalent modification of DNA by xenobiotics may be involved in
chemical carcinogenesis. Moreover, acetaldehyde-DNA adducts have also been detected
in human buccal cells exposed to acetaldehyde in vitro (Vaca et al., 1998).
There is sufficient evidence for the carcinogenicity of acetaldehyde to experimental
animals (IARC, 1985). Acetaldehyde has been tested for carcinogenicity in rats and
35
hamsters by inhalation exposure. In such experiments, an increased incidence of
carcinomas was observed in the nasal mucosa of rats (Woutersen et al., 1984), and
laryngeal carcinomas were induced in hamsters (Feron et al., 1982). Moreover, in rats
given either tap water or water containing acetaldehyde at a concentration of 120 mM,
marked histopathological hyperplastic and hyperproliferative changes in the tongue,
epiglottis, and forestomach were noted in the animals receiving acetaldehyde. These
changes mimic those known to occur after treatment with alcohol (Homann et al.,
1997b).
Lipid peroxidation
A free radical is generally defined as a molecule that contains one or more unpaired
electrons. As the presence of unpaired electrons usually confers a large degree of
chemical reactivity on the molecule, most free radicals (superoxide and hydroxyl
radicals) may lead to cell injury by abstracting a hydrogen atom from a polyunsaturated
fatty acid, and thus initiating the degradative process known as lipid peroxidation. Since
lipids are major components of biological membranes, peroxidative loss of membrane
integrity has been thought to lead to tissue injury (Mufti et al., 1993). Glutathione is
present in all animal cells in high concentrations, and one of its functions is the
protection of cells against free radicals. A severe reduction in the levels of glutathione
has been shown to increase lipid peroxidation in vivo (Lieber, 1988; Wendel et at.,
1979).
Enhanced lipid peroxidation has been suggested as a mechanism for ethanol-associated
liver injury (Situnayake et al., 1990). During ethanol oxidation reactive oxygen
intermediates are produced by the MEOS. Acetaldehyde can also induce lipid
peroxidation, as demonstrated in isolated perfused livers (Müller and Sies, 1982). The
mechanism underlying this may be acetaldehyde’s capacity to reduce hepatic glutathione
levels (Shaw et al., 1981). In addition, the incubation of rat liver supernatant with
acetaldehyde results in the conversion of xanthine dehydrogenase to xanthine oxidase,
an enzyme known to be able to generate superoxide radicals (Sultatos, 1988). Whether
ethanol administration enhances in vivo lipid peroxidation has long been debated, but
studies done on laboratory animals (Niemelä et al., 1995; Shaw et al., 1981) and humans
(Niemelä et al., 1994; Shaw et al., 1983) point to this ability. Moreover, high
acetaldehyde concentrations administered to rats have been shown to result in the
formation of free radical reactions in vivo (Reinke et al., 1987). Taken together, ethanolderived
acetaldehyde may lead to a severe reduction in glutathione, which favours lipid
peroxidation, and the damage is possibly further enhanced by the increased generation
of active radicals through induced MEOS following sustained ethanol consumption.
Lipid peroxidation products, which result from the attack of reactive oxygen species on
lipids, also react with DNA, forming ethenobases and adducts with known
carcinogenicity and miscoding potential. Thus, during chronic alcohol abuse, where
levels of reactive oxygen species and lipid peroxidation products are elevated and
antioxidant levels are reduced, there is potential for significant damage to DNA and the
production of DNA adducts that can compromise cellular function and may lead to
oncogenic transformation (Brooks, 1997).
36
Evidence in humans
Acetaldehyde exposition studies with humans are nowadays naturally forbidden. In the
past, however, these have been done. Human volunteers exposed for 15 min to
acetaldehyde vapour (90 mg/m3) experienced mild eye irritation. Men exposed to a
higher concentration of acetaldehyde (360 mg/m3) for the same time developed
transient conjunctivitis. Moreover, all 14 men exposed to 241 mg/m3 of acetaldehyde
for 30 min developed mild upper-respiratory tract irritation (IARC, 1985).
Indirect but stronger evidence for acetaldehyde as the major factor behind ethanolassociated
carcinogenesis is derived from recent studies linking the genotypes of
ethanol- and acetaldehyde-metabolizing enzymes with enhanced tumour risk. Rapid
metabolizing alcohol dehydrogenases (ADH3), leading to higher and quicker production
of cellular acetaldehyde (Harty et al., 1997), and the lack of a low-km aldehyde
dehydrogenase (ALDH2), leading to a longer and delayed exposure to acetaldehyde
(Yokoyama et al., 1996a, b, c), have both been shown to be associated with an increased
cancer risk in the oropharynx and esophagus. In a recent study among Orientals, the
association between ALDH2 genotype mutation and cancer risk in alcoholics has been
expanded to all possible alcohol-related cancers. In this study, the frequencies of the
mutant ALDH2*2 allele were significantly higher in alcoholics with oropharyngeal,
laryngeal, esophageal, stomach, colon and lung cancer, but not with liver or other
cancers (Yokoyama et al., 1998). The relationship between alcohol consumption,
ALDH2 heterozygocity, and the risk of colon cancer has also been confirmed very
recently by others (Murata et al., 1999) as well as the lack of risk of alcohol-associated
hepatocellular carcinoma in ALDH2 heterozygotes (Takeshita et al., 2000).
So far this phenomenon has been thought to arise from systemic effects of elevated
blood acetaldehyde (Yokoyama et al., 1996a, b, c). Interestingly, however, all the organs
with enhanced cancer risk are covered by microbes. It is therefore possible that the
impaired detoxification of acetaldehyde in ALDH2 deficient subjects might become
clinically relevant only in cases of marked acetaldehyde production by microbes. This
hypothesis is strongly supported by our very recent finding demonstrating that Asians
with mutant ALDH2*2 allele had 2-3 times higher in vivo salivary acetaldehyde levels
after a moderate dose of ethanol than those Asians with a normal ALDH 2*1 genotype
throughout the whole follow-up period of 240 minutes. The in vitro capacity of saliva to
produce acetaldehyde from ethanol was equal in these two groups, which suggests that
there were no obvious difference in the capacity of the oral flora to produce
acetaldehyde from ethanol between the groups. The subjects with the mutant ALDH2*2
allele appeared to be able to produce higher in vivo salivary levels of acetaldehyde
because their parotid glands also contributed to acetaldehyde production, a phenomenon
that did not occur in subjects with the normal genotype. Possible differences in the
capacity of the oral mucosa to metabolise acetaldehyde further to acetate might also
explain this (Väkeväinen et al., 2000). This study, together with earlier epidemiological
findings, provides strong evidence for the local carcinogenic potential of acetaldehyde in
humans
37
3. AIMS OF THE STUDY
Alcohol consumption has increased in Finland from the late 1960's. Alcohol-related
gastrointestinal disorders and organ injuries are consequently an increasing health
problem. Relatively little is, however, known about their pathogenesis. Ethanol per se
appears to be unable to cause most of them.
During recent years it has become evident that microbes representing the normal flora of
the alimentary tract participate in the metabolism of exogenous ethanol. Consequently,
high levels of acetaldehyde are produced in those parts of the GI tract that hold the
largest number of microbes - the oral cavity and colon. Because of its high toxicity and
carcinogenic potential, acetaldehyde can be expected to cause organ damage wherever it
exists at high concentrations.
Understanding of the mechanisms behind alcohol-associated gastrointestinal morbidity
could be helpful in their management and a prerequisite for their prevention. Research
aimed at exploring those mechanisms is therefore justified.
The specific aims of the study were:
1. To study the enzymatic mechanisms underlying acetaldehyde production from
ethanol by human colonic contents.
2. To evaluate the role of microbial ethanol oxidation in ethanol metabolism in
humans.
3. To explore the effect of antibiotic treatment on the formation of acetaldehyde in the
gastrointestinal tract and to clarify the bacterial species responsible for the
intracolonic acetaldehyde production from ethanol.
4. To study the effect of chronic ethanol treatment on intracolonic acetaldehyde levels
in rats.
5. To investigate the effect of folate depletion on the carcinogenic action of acetaldehyde.
6. To elucidate the factors and microbial species which regulate the microbial
production of acetaldehyde from ethanol in the oral cavity of humans.
38
4. MATERIALS AND METHODS
4.1. ETHICAL CONSIDERATIONS
In the studies with humans volunteers (I, II, V, VI) informed consent was given, after approval
by the Ethical Committee at the Helsinki University Central Hospital. The rat studies were
approved either by the ethical committee of the Helsinki University Central Hospital (III) or by
the Committee on Animal Experimentation of the County Council (IV). Animal studies were
performed according to the institutional guidelines and principles of the Animal Care Unit of
the University of Helsinki.
4.2. ACETALDEHYDE PRODUCTION BY HUMAN COLONIC CONTENTS IN
VITRO (I)
Collection of colonic contents
Colonic contents were collected from 14 Finnish patients undergoing colonoscopy for lower
GI symptoms. The age of the patients ranged from 29 to 74 years (mean 51 years). The
exclusion criterion was the use of antibiotics during the period of 4 weeks preceding the
colonoscopy. During the colonoscopy, approximately 10 ml of colonic content was aspirated
through a fiberoscope from the caecum and transverse colon. The samples were frozen
immediately at -80 C pending analysis.
Determination of acetaldehyde production in vitro
Samples were first lyophilized for 24 hr, whereafter the dry mass was dissolved in 100 mM of
potassium phosphate buffer (pH 7.4) at a concentration of 10 mg/ml, except when studying the
effect of different quantities of the colonic contents. 400 μl of resuspended colonic content was
transferred into a gas chromatographic vial, and 50 μl of potassium phosphate buffer (final
concentration 100 mM, pH 7.4) containing ethanol (final concentration 22 mM) was added and
the vials were immediately tightly closed. The vials were incubated for 60 min at 37 C. The
effect of different incubation times, increasing ethanol concentrations and different amounts of
colonic contents on acetaldehyde production were also tested.
To study the effect of different enzyme inhibitors, Sodium azide (SA, a catalase inhibitor), 3-
amino-1,2,3-triazole (3-AT, a catalase inhibitor), 4-methylpyrazole (4-MP, an ADH inhibitor),
or metyrapone (a MEOS inhibitor) were added at increasing concentrations 15 min prior to
ethanol/buffer mixture to the vials containing colonic contents.
To confirm catalase and ADH as the enzymes responsible for acetaldehyde production,
increasing concentrations of the hydrogen peroxide generating system (final glucose
concentration 10 mM, glucose oxidase 0.003 - 0.3 μmol/min) or exogenous NAD (final
concentration 1 - 10 mM) were added to the incubation mixture.
After incubations reactions were stopped by injecting 50 μl of 6 mol/l perchloric acid (PCA)
through the rubber septum of the vial. Acetaldehyde was analysed by head space gas
chromatography as described in chapter 4.8.
39
Analyses of catalase activity
To prepare supernatant for measurement of enzyme activity, the colonic contents were
dissolved in 100 mM potassium phosphate buffer and first sonicated. This was followed by
centrifugation of the sonicate at 100,000 g at 5 C for 60 minutes. Catalase activity was
determined spectrophotometrically at 240 nm after the addition of 10 mM of H2O2 at 37 C.
Catalase activity was related to the protein concentration of the supernatant (Lowry et al.,
1951).
4.3. THE EFFECT OF CIPROFLOXACIN ON ETHANOL ELIMINATION IN
HUMANS (II)
Volunteers
Eight healthy Caucasian males (age range of 21-31 years, mean BMI 23.8 ± 0.4 kg/m2)
participated in the study. None of the subjects had received any antibiotics for four weeks
preceding the study. The volunteers’ weekly average consumption of alcohol was about 70
grams of ethanol. All participants were told to refrain from ethanol for at least 36 hours before
the study.
Study design
The design was open, non-randomized, and non-placebo controlled, each subject serving as his
own control. The two study days were separated by a 1-week interval, and the protocol was
exactly the same on both occasions. Two intravenous lines were placed in the antecubital
veins, one for the administration of ethanol and one for obtaining blood samples. Ethanol (0.63
g/kg body weight) was mixed in 5% glucose solution at 7% v/v concentration and was
administered intravenously at a constant rate over a 30-min period. Repeated blood samples (3
ml) were taken into vacutainer tubes for measurements of blood alcohol level by head space
gas chromatography. Baseline samples were taken before ethanol administration had started
(time 0) and at five minute intervals during the first hour, at 15 minute intervals during the
second hour, and at 20 minute intervals until the breath ethanol analyser showed no detectable
blood ethanol levels. For seven days between the experiments, the volunteers received 750 mg
ciprofloxacin orally twice a day.
Pharmacokinetic calculations of ethanol in blood
The concentration-time profiles of ethanol were evaluated according to zero-order kinetics.
The y-intercept of the regression line (C0) is the concentration of ethanol in blood if the dose
of 0.63 g/kg was distributed into total body water immediately after the infusion started. The
ratio of the dose of ethanol (g/kg) divided by the parameter C0 is the apparent volume of
distribution of ethanol (Vd). The ethanol elimination rate (EER) from the body was obtained
by dividing the dose given by the estimated time of reaching zero concentration of ethanol in
blood (time0). The time0 parameter corresponds to the x-intercept of the concentration-time
regression equation. The areas under the concentration-time profiles (AUCs) were determined
by the trapezoidal method (Rangno et al., 1981).
40
The effect of ciprofloxacin on human hepatic ADH in vitro
A piece of human liver tissue was obtained from a patient undergoing surgery. The tissue was
first homogenised 1:4 with 100 mM potassium phosphate buffer, pH 7.4. The homogenate was
centrifuged at 1000 g at 4 C for 10 min, followed by centrifugation at 100,000 g at 5 C for 60
min to obtain the cytosolic fraction. Hepatic ADH activity was determined
spectrophotometrically by monitoring the formation of NADH at 340 nm at 37 C. The ADH
activity of cytosolic fraction was measured in 100 mM potassium phosphate buffer containing
3 mM of NAD and 25 mM of ethanol. The effect of ciprofloxacin on ADH activity was tested
by adding increasing drug concentrations to the buffer (final concentrations 0, 1, 10, 20 μg/L).
4.4. THE EFFECT OF CIPROFLOXACIN ON HUMAN FAECAL FLORA AND
ACETALDEHYDE PRODUCTION (II)
Microbial analysis
The faecal samples were thawed, and 1 gram of each specimen was suspended and serially
diluted (10-fold) in peptone yeast extract broth. The undiluted sample and a 10 μl aliquot of
the appropriate dilutions were inoculated and spread on several selective and non-selective
agar media for the enumeration and isolation of total counts and main groups of aerobic and
anaerobic bacteria and yeasts. The aerobic plates were incubated at 35 C in an atmosphere
containing 5% CO2 for up to 5 days; anaerobic plates were incubated in anaerobic jars filled
by the evacuation replacement method with mixed gas (90% N2, 5% CO2, 5% H 2) up to 14
days for the final inspection. The bacteria were enumerated and identified by established
methods (Murray et al., 1999; Summanen et al., 1993).
Determination of faecal ADH and catalase activity
Faecal samples were collected before and after the ciprofloxacin treatment from the volunteers
described in 4.3. Samples were frozen at -80 for further analysis. Supernatant from lyophilized
faecal samples was prepared, and the ADH activity of the supernatant was determined as
described in 4.3. Catalase activity was determined as described in 4.2. Enzyme activities were
related to the protein concentrations of the supernatant (Lowry et al. 1951).
Faecal acetaldehyde production in vitro
250 μl of supernatant was transferred into a gas chromatography vial, and 200 μl of potassium
phosphate buffer (final concentration 100 mM, pH 7.4) containing ethanol (final concentration
22 mM) and different coenzymes (NAD or glucose + glucose oxidase or both) was added and
the vials were closed. The vials were incubated for 60 minutes and the reaction was stopped by
PCA. Acetaldehyde production was related to the protein concentration of the supernatant
(Lowry et al., 1951).
4.5. SUSTAINED ETHANOL AND METRONIDAZOLE TREATMENT IN RATS (III)
Animals and study protocol
41
Male Wistar rats (strain Hsd/wi barrier) were used and kept under conventional conditions.
To study the effect of long-term ethanol administration and metronidazole treatment on
ethanol and acetaldehyde metabolism in rats, 32 rats were housed individually in stainlesssteel
wire-bottomed cages. When the rats reached an average weight of 190 ± 1 g they were
switched to a liquid diet (Lindros and Järveläinen, 1998) This was the control diet. After two
days adaptation, the rats were divided into 4 groups as follows.
Group 1 (C, control, n=6) Control diet, pair-fed to 3
Group 2 (CM, n=6) Control diet + metronidazole 50 mg/kg b.w./day, pair-fed to 4
Group 3 (E, ethanol, n= 10) Ethanol diet ad libitum
Group 4 (EM, ethanol metronidazole, n=10) Ethanol diet ad libitum + metronidazole 50 mg/kg b.w./day
The ethanol content of the diet was increased gradually up to the final 5% w/v. The diets were
renewed every 24 h at 9.00. Diet intake was recorded daily and weight gain twice a week. The
rats were maintained on pair-feeding for 6 weeks.
Blood ethanol and acetaldehyde analysis
To measure diurnal blood ethanol concentrations, tail-vein blood samples (50μl) were taken
from rats receiving ethanol on four different days at 6.00, 12.00, 18.00, and 24.00 and
haemolysed in 450 μl of ice-cold water. Terminal blood was taken by heart puncture. Blood
ethanol and acetaldehyde levels were determined by head-space gas chromatography.
Colonic ethanol and acetaldehyde levels
Caecal samples were diluted 1:6 with distilled ice-cold water and mixed carefully. An aliquot
of 450 μl was pipetted into gas chromatographic vials and immediately mixed with 50 μl of 6
mol/l PCA. Intestinal ethanol and acetaldehyde levels were analysed by head space gas
chromatography.
Preparation of the tissues for enzyme determination
Before enzyme analyses, livers were perfused in situ with saline, and samples were transferred
to - 80 C pending analysis. Ice-cold medium containing 0.25 M of sucrose, 5 mM of Tris, and
0.5 mM of EDTA (pH 7.2) was then added to the tissues in an amount making the tissue
constitute 15 to 20% of the total. This was followed by homogenization and sonication for 9 x
5 sec at +4 C. The homogenate was centrifuged at 700 g for 15 min to remove unbroken cells
and nuclei and the supernatant was used for all enzyme assays.
The colons were washed with cold saline and the mucosal layers collected by gentle scraping.
To analyse enzyme activity, mucosa of the large intestine were handled as described above.
Determination of ALDH and ADH activity
Hepatic and colonic mucosal ALDH and ADH activities were determined
spectrophotometrically by measuring the formation of NADH at 340 nm at 25 C. The ALDH
activity of the supernatant was assayed in 60 mM of sodium pyrophosphate buffer (pH 8.8)
containing 0.5 mM of NAD, 0.1 mM of 4-MP, 2 μM of rotenone, and either 100 μM (low Km
activity) or 5 mM (total activity) of acetaldehyde. The ADH activity of cytosolic fractions was
42
measured in 100 mM glycine buffer (pH 9.6) containing 1 mM of NAD, 2 μM of rotenone,
and 25 mM of ethanol. The protein concentration was determined by the Bio-Rad method
(Bio-Rad protein assay, Hercules, USA).
Microbial analysis
Caecal content microbial analysis was carried out as described in chapter 4.4.
4.6. THE EFFECT OF ACETALDEHYDE ON INTESTINAL FOLATE LEVELS IN
RATS (IV)
Animals and study protocol
The animals were male Wistar rats (strain Hsd/wi barrier).
To study the effect of ethanol-derived acetaldehyde on intestinal folate levels, 40 rats were
randomly divided into 4 groups as follows.
Group 1 (saline, n=10) Intragastric (ig incubation 2x/day with saline
Group 2 (Saline/Cipro, n=10) Ig incubation 2 x/day with saline and ciprofloxacin 50 mg/kg
Group 3 (Ethanol, n=10) Ig incubation 2 x/day with ethanol 3g/kg b.w. as 38% v/v
Group 4 (Ethanol/Cipro, n=10) Ig incubation 2 x/day with ethanol and ciprofloxacin as above
Intubations were carried out with the same volume in every group for 14 days. Rats had free
access to tap water and standard chow (Altromin Nr 1324 Pellets, Lage, Germany) with a
folate content generally accepted as meeting the basal dietary requirement for a rat (2mg folate/
kg diet) (Reeves et al., 1993). The general condition, body weight, and food intake were
recorded daily. Rats were killed after anasthesia with 1 mg/kg phenobarbital on day 14 of the
study, 45 minutes after the last intubation.
Blood ethanol analysis
Terminal blood ethanol levels were determined by head-space gas as described in 4.5.
Colonic and small intestinal ethanol and acetaldehyde levels
The small intestinal content, obtained 40 cm caudal from the stomach, and caecal contents
(about 1.5 ml) were transferred into Eppendorf tubes containing 50 μl of PCA. Thereafter
contents were spun down by centrifugation at 2400 g for 20 sec. An aliquot of 475 μl of the
supernatant was immediately transferred into a gas chromatographic vial containing 25 μl of
PCA. Intestinal ethanol and acetaldehyde levels were analysed by head space gas
chromatography.
Preparation of the tissues for folate analysis
For folate measurements small intestinal and colonic mucosa were transferred to pre-weighed
Eppendorf tubes. Thereafter, the wet weight of the scraped mucosa was overlaid with ten
volumes of folate-lysis buffer (Kim et al., 1996). Folate glutamates were cleaved and
43
transferred by incubation with folate-free chicken pancreas conjugase, then stored at –70 C
until analysed (Kim et al., 1996).
Folate analysis
Serum and erythrocyte folate levels were measured by conventional enzymatic methods
following the manufacturer’s instructions (Simultrac Radioassay Kit, ICN Pharmaceuticals,
NY, USA), and taking the current hematocrit value into account. The folate content of the gut
mucosa was measured by enzymatic methods using the same kit as for the serum folate. The
folate levels of the intestine was related to the mucosal protein content (Bio-Rad protein assay,
Hercules, USA).
4.7. HUMAN SALIVA STUDIES (V, VI)
Subjects
A total of 326 Caucasian volunteers participated in the studies. This cohort consisted of 114
healthy volunteers, 122 patients seeking dental examination or treatment, 26 patients with a
malignant tumour of the oral cavity (11 were untreated and 15 were in the follow-up stages),
and 64 alcoholics recruited from a municipal alcohol detoxification clinic.
Questionnaire
A questionnaire was answered by each volunteer. Information concerning age, gender, tobacco
use, alcohol use, diet, oral health status, oral hygiene habits and other characteristics were
elicited. Tobacco use indicators included the average number of cigarettes smoked per day
within the past 30 days, duration of smoking in years, and the date when a possible smoking
cessation may have occurred. The daily tobacco consumption was calculated as cigarettes
smoked per day. Alcohol consumption was estimated as the average number of drinks ingested
(about 12g of pure alcohol) for every drinking day during the past 30 days and as the frequency
of alcohol intake per week. The average amount of alcohol consumed as grams of pure ethanol
per day was calculated from this data. Volunteers were ranked as non-drinkers (less than 1
gr/day), moderate drinkers (1-30 gr/day for females, 1-40 gr/day for males) or heavy drinkers
(>30 gr/day for females, >40 gr/day for males).
Salivary samples
Stimulated whole saliva was collected between 9 and 12 a.m. after one minute’s use of a
paraffin chewing gum (Orion Diagnostics, Espoo, Finland), and was immediately frozen at -70
C. Exclusion criteria were as follows: treatment with oral antiseptic or antibiotics in the past
month, food or fluid intake, smoking or toothbrushing in the previous 90 minutes, recent
alcohol intake or a measurable amount of alcohol in the saliva by head space gas
chromatography.
Determination of salivary acetaldehyde production capacity
Saliva was thawed and preheated to 37 C before analysis. 400 μl of saliva was transferred into
a gas chromatography vial, and 50 μl of potassium phosphate buffer (final concentration 100
mM, pH 7.4) containing ethanol (final concentration 22mM) was added and the vials were
44
immediately tightly closed. The vials were incubated for 90 minutes and the reaction was
stopped by injecting 50 μl of PCA, whereafter acetaldehyde was analysed using head space gas
chromatography.
Salivary microbiological analysis
Among all 326 volunteers, the 10 saliva samples with the lowest and the highest acetaldehyde
production were chosen for microbial analysis (V). Since this analysis revealed that yeasts
were found in higher concentrations and more frequently among the subjects with higher
acetaldehyde production, all the saliva samples with acetaldehyde production of more than 250
μM (23 samples) or less than 40 μM (32 samples) were used to assess the prevalence of yeasts
(VI).
The saliva samples were thawed, and serially diluted in peptone yeast extract broth. A 10 μl
quantity of undiluted sample and the appropriate dilutions were inoculated on several selective
and nonselective agar media for the enumeration and isolation of aerobic and anaerobic
bacteria and yeasts. The aerobic plates were incubated at 36 C in an atmosphere containing 5%
CO2 for a total of 5-7 days, and anaerobic plates in anaerobic jars filled with the evacuation
replacement method with mixed gas (85% N2, 10% CO 2, 5% H 2) were incubated up to 14
days for the final inspection. Bacterial counts were determined by multiplying the number of
colonies by the dilution factor, adjusted for inoculation volume (V). Yeast colonies (VI) were
enumerated and identified by germ tube test and by API ID 32 C (bioMérieux, Marcy l’Etoile,
France). The yeast cultures were then harvested three times before yeast suspensions were
prepared. The actual number of viable yeasts, expressed as colony forming units (CFU/ml), in
the vials was determined by quantitative viable count (VI).
Acetaldehyde production by isolated yeast strains
Acetaldehyde analysis was carried out as with the saliva samples, except the incubations were
carried out for 60 minutes.
4.8. GAS CHROMATOGRAPHIC MEASUREMENTS OF ETHANOL AND
ACETALDEHYDE
Acetaldehyde production from ethanol by colonic contents in vitro (I), faecal samples in vitro
(II), caecal samples in vivo (III, IV), salivary samples in vitro (V), and isolated oral yeasts (VI)
was analysed using head space gas chromatography by heating the vials to a temperature of 37
C as reported earlier (Pikkarainen et al., 1979). The conditions for analysis were: Column
60/80 Carbopack B/5% Carbowax 20M, 2 m x 1/8" (Supelco Inc, Bellefonte, PA, USA); oven
temperature, 85 C; transfer line and detector temperature, 200 C; carrier gas flow rate (N2), 20
ml/min. Acetaldehyde production after certain incubation times in vitro (I, II, V, VI) or in in
vivo studies immediately after collection was stopped with PCA.
An artifactual formation of acetaldehyde from ethanol prior to the headspace analysis
(Eriksson and Fukunaga, 1993) is a problem associated with analysis of acetaldehyde in
biological fluids. Precipitation of proteins results in the non-enzymatic production of
artifactual acetaldehyde, an effect that cannot be completely eliminated, even by centrifugation
and removal of the protein precipitates (Sippel, 1972; Stowell et al., 1977). Several analytical
modifications have been described in order to minimize artifactual acetaldehyde formation;
45
nevertheless, parallel analyses with control samples using corresponding ethanol
concentrations should be carried out (Eriksson and Fukunaga, 1993).
To control for non-enzymatic artifactual acetaldehyde formation from ethanol during the
protein precipitation, perchloric acid was added simultaneously with ethanol into additional
incubation vials (incubation time 0). The acetaldehyde concentrations of these control samples
were subtracted from acetaldehyde values obtained after longer incubation periods (I-VI).
4.9. STATISTICAL ANALYSIS
All results are expressed as means ± SEM (I-VI). To test the effect of drugs or conditions on
acetaldehyde production ANOVA for repeated measures, followed by Bonferroni’s t test was
used (I). The possible statistical significance of the differences before and after the
ciprofloxacin intake was analysed by Student’s t test (II). In the animal studies the differences
between the groups were analysed by ordinary ANOVA, followed by the Tukey-Kramer
Multiple Comparison Test. Logarithmic transformation was performed when appropriate (III,
IV). To analyse the effect of various factors on salivary acetaldehyde production, a Spearman
correlation matrix was computed for the entire study population as a preliminary analysis. This
was followed by multivariate regression analyses. As co-linearity was obvious for smoking and
heavy alcohol intake, the multivariate analyses were re-run for non-smokers and for moderate
and non-drinkers in order to adjust for this confounding factor. A multiple linear regression
analysis, a forward stepwise regression analysis (r2 as the best criterion) was run with the best
descriptor for all variables, setting acetaldehyde production as the dependent variable (V).
When the statistical significance of the differences between “high” and “low” acetaldehyde
producers was compared, the nonparametric unpaired Mann Whitney U test was used (V, VI).
Fisher’s exact test was used to identify possible differences in the presence of yeasts between
two groups (VI).
46
5. RESULTS
5.1. ENZYMATIC PRODUCTION OF ACETALDEHYDE BY HUMAN COLONIC
CONTENTS IN VITRO (I)
The amount of acetaldehyde produced by the human colonic content was proportional to the
ethanol concentration, the amount of colonic content, and the length of the incubation time.
Both catalase inhibitors, SA and 3-AT, reduced the amount of acetaldehyde produced from
ethanol in a concentration dependent manner as compared to the control samples (Fig. 1). SA
decreased the acetaldehyde production (by 26.4 ± 6.9%) significantly at a concentration of 0.1
mM and 3-AT (by 32.6 ± 13.0%) at a concentration of 10 mM. Metyrapone and 4-MP had no
significant effect on acetaldehyde production under these conditions.
Col 1
Acetaldehyde production (% of control)
0
20
40
60
80
100
*
*
*
*
* *
*
*
*
*
Sodium azide (mM) 3-amino-1,2,3-triazole (mM)
0 .1 .5 1 5 10 50 100 0 1 5 10 50 100
Fig. 1. The effect of catalase inhibitors on acetaldehyde production by colonic contents during a 90 min
incubation with 22 mM ethanol. * p <0.05 compared to control. (Reproduced with permission from
Lippincott Williams & Wilkins).
Exogenous hydrogen peroxide produced by the glucose + glucose oxidase system significantly
increased acetaldehyde production, in proportion to higher glucose oxidase activity. The
highest value was reached with 0.3 μmol/min glucose oxidase (677% ± 156 of the control)
(Fig. 2). The acetaldehyde production was also increased after the addition of NAD. With 3
mM NAD the acetaldehyde production was increased up to 5 times as compared to the
controls.
47
Glucose oxidase (μmol/min)
Acetaldehyde production (% of control)
0
100
200
300
400
500
600
700
800
*
*
0 0.003 0.03 0.3
Fig. 2. The effect of a H2O2 generating system (glucose + glucose oxidase on acetaldehyde
formation during a 60 min incubation with 22 mM ethanol. * p < 0.05 compared to control. (Reproduced
with permission).
The mean catalase activity of the supernatant was 0.53 ± 0.1 μmol/min/mg protein with 10mM
H2O2. There was a significant correlation between catalase activity and acetaldehyde
production by the supernatant in the presence of glucose + glucose oxidase (r=0.96, p<0.05)
indicating the role of catalase in acetaldehyde production under these circumstances.
5.2. THE EFFECT OF CIPROFLOXACIN ON ETHANOL ELIMINATION IN
HUMANS (II)
Table 2 summarizes the pharmacokinetic parameters of ethanol derived from blood
concentration time data. The time to reaching zero ethanol concentration in blood (time0)
increased after ciprofloxacin medication, and accordingly there was a highly significant 9.4%
decrease (range 5.1% to 17.6%; p=0.001) in the ethanol elimination rate (EER).
Table 2. Effects of ciprofloxacin on ethanol pharmacokinetics (mean±SEM)
Before Cipro After Cipro p
Peak EtOH (mM) 22.5 ± 1.0 23.3 ± 1.3 n.s.
Vd (l/kg) 0.70 ± 0.02 0.70 ± 0.02 n.s.
C0 (mM) 19.7 ± 0.5 19.7 ± 0.6 n.s.
EER (mg/kg/h) 107.0 ± 5.3 96.9 ± 4.8 0.001
AUC (mM·h) 58.9 ± 2.9 65.5 ± 3.3 0.0004
Time0 (h) 6.0 ± 0.3 6.6 ± 0.3 0.0003
48
5.3. THE EFFECT OF CIPROFLOXACIN ON HUMAN FAECAL FLORA AND
ACETALDEHYDE PRODUCTION (II)
The effect on microbial flora
Ciprofloxacin treatment for seven days produced a 17-fold decline in the total number of
faecal aerobic bacteria. Enterobacteriaceae, the predominant aerobic flora present in every
volunteer at the beginning of the study, disappeared from the faeces entirely after ciprofloxacin
treatment. Enterococcus sp., which were found in five of the eight subjects before the
ciprofloxacin intake, were also eradicated after the treatment. Other aerobic species responded
variably. Yeasts appeared at low levels in two subjects after ciprofloxacin administration. The
count of anaerobic bacteria declined slightly. This was mainly due to a drop in the number of
Bifidobacterium species. The bacteriological data is seen in detail in the original article.
The effect on faecal enzymes and acetaldehyde production
The mean ADH activity of the faecal samples measured after the ciprofloxacin treatment was
significantly lower at both 25 mM (p=0.013) and 1.5 M (p=0.006) ethanol concentrations than
of the samples taken before the treatment. The ADH activity at both ethanol concentrations
decreased approximately 60% after the treatment. The catalase activity, however, remained
unchanged after ciprofloxacin dosing. The acetaldehyde production capacity of the faecal
samples also decreased significantly (by 60%, p=0.007) after ciprofloxacin treatment when
NAD was used as a cofactor to activate ADH, but remained unaltered when glucose + glucose
oxidase was used to activate catalase. There was also a statistically significant correlation
(r=0.75, p<0.001) between faecal ADH activity at 1.5 M ethanol and acetaldehyde production
from ethanol.
5.4. THE EFFECT OF SUSTAINED ETHANOL AND METRONIDAZOLE
TREATMENT ON INTRACOLONIC ACETALDEHYDE PRODUCTION IN RATS
(III).
Animals
All animals tolerated the six-week treatment period. However, the body weight decreased most
in the rats receiving ethanol and metronidazole. The weight of the rats (initials-finals) were as
follows: group 1 (C, control) 185±3 g - 214±3 g, group 2 (CM, control metronidazole) 188±3
g - 206±3 g, group 3 (E, ethanol) 194±2 g - 177±7 g, group 4 (EM, ethanol metronidazole)
192±3 - 146±5 g.
Intracolonic ethanol and acetaldehyde concentrations
Intracolonic terminal ethanol concentrations were comparable in groups E and EM (34.0±2.0
mM and 36.8±0.6 mM respectively, n.s.). However, the rats in group EM had five times higher
intracolonic acetaldehyde levels than those receiving only ethanol (431.4±163.5 μM and
84.7±14.4 μM respectively, p<0.01, Fig. 3).
49
0
100
200
300
400
500
600
Col 3
C CM E EM
p<0.001 vs CM
p<0.01 vs E
p<0.01 vs C
Fig. 3. Intracolonic acetaldehyde levels in the study
groups (mean±SEM)
C= Controls
CM = Controls receiving metronidazole
E = Ethanol group
EM = Ethanol and metronidazole group
(Reproduced with permission from Lippincott Williams & Wilkins).
Colonic mucosal ADH and ALDH activities
Metronidazole treatment did not inhibit colonic mucosal ADH or ALDH activity. Indeed,
colonic mucosal ALDH activity was found to be higher in the rats which received ethanol and
metronidazole than the ethanol group (Table 3).
Table 3. Colonic mucosal ADH and ALDH activities in the study groups (mU/mg)
Group (n) ADH ALDH ALDH
(High Km) (Low Km)
C (6) 4.8 ± 0.4 4.0 ± 0.4 1.3 ± 0.2
CM (6) 6.7 ± 1.8 3.5 ± 0.4 1.9 ± 0.3
E (10) 6.9 ± 1.0 4.2 ± 0.3 0.9 ± 0.2
EM (10) 6.7 ± 1.5 6.6 ± 0.7*,** 2.2 ± 0.2***
* p<0.05 compared to E
** p<0.01 compared to CM
*** p<0.001 compared to E
50
Bacteriological analysis of caecal contents
Ethanol and metronidazole treatment led to 3-fold reduction in the total anaerobic caecum
flora compared to the rats receiving only ethanol (p<0.05). Particularly the number of
Bacteroides fragilis were reduced (p<0.01). By comparison, ethanol and metronidazole
treatment produced a 18-fold increase in the number of total aerobes (p<0.01) compared to the
ethanol group, and especially the number of Enterobacteriaceae (p<0.05) and -haemolytical
Streptococci (p<0.001) increased. It should also be noted that ethanol treatment alone
increased significantly the number of total aerobes and Enterobacteriaceae as compared to the
controls. The exact bacteriological counts are seen in the original article.
5.5. THE EFFECT OF ETHANOL AND METRONIDAZOLE TREATMENT ON
BLOOD ETHANOL AND ACETALDEHYDE LEVELS IN RATS (III)
Ethanol intake, blood ethanol and acetaldehyde concentrations
Ethanol intakes were equal in groups E and EM: 11.8±0.2 vs. 11.4±0.2 g/kg body wt. per day
respectively (n.s.). Animals in groups E and EM were found to be ethanol-intoxicated
throughout the 24 h period, and no significant differences in the diurnal blood ethanol levels
between these groups could be detected during the trial. The terminal blood ethanol levels in
groups E and EM were 38.0±1.8 mM and 42.1±2.0 mM respectively (n.s.). Terminal blood
acetaldehyde levels were 7.0±0.3 μM in group E and 7.4±0.2 μM in group EM (n.s.).
Liver ADH and ALDH activity
Metronidazole treatment did not inhibit hepatic ADH or ALDH enzymes in a statistically
significant way (Table 4.).
Table 4. Hepatic ADH and ALDH activities in the study groups (mU/mg)
Group (n) ADH ALDH ALDH
(High Km) (Low Km)
C (6) 17.7 ± 1.3 9.2 ± 1.7 8.5 ± 1.3
CM (6) 16.7 ± 1.6 7.6 ± 1.0 7.6 ± 0.7
E (10) 16.2 ± 0.8 11.5 ± 1.0 6.5 ± 0.8
EM (10) 15.5 ± 0.5 13.2 ± 0.9* 6.3 ± 0.6
* p<0.05 compared to CM
5.6. THE EFFECT OF ACETALDEHYDE ON INTESTINAL FOLATE LEVELS IN
RATS (IV)
Animals
All animals tolerated the two weeks’ experiment, except of one rat from group 4 that died due
51
to the regurgitation of intubated solution. During the study, the rats in group 1 kept their
weights, whereas there were significant weight losses (4 to 10 %, p<0.01) in the other three
groups. However, the differences in the total body weight at the end of the experiment were
not significant between the groups.
Ethanol and acetaldehyde levels
Mean blood ethanol concentration in the ethanol-treated rats was 34±5 mM in group 3 and
33±5 mM in group 4 (n.s.). Intracolonic ethanol levels were also similar in groups 3 and 4
(28±5 and 33±7 mM respectively, n.s.). There was a highly significant correlation between the
individual blood ethanol and intracolonic ethanol levels (r=0.84, p<0.0001, data not shown).
Small amounts of endogenous ethanol (1.4±0.3 mM) were detectable in the colon in the nonethanol-
treated rats, and this was reduced by ciprofloxacin treatment to the level of 0.9±0.1
mM (n.s.). Small intestinal ethanol levels in group 3 and 4 were similar.
The local acetaldehyde levels in the colon after ethanol intake were strikingly high (387±185
μM in group 3), which was significantly reduced (p<0.001) by concomitant ciprofloxacin
treatment (Fig. 4). Also, substantial endogenous colonic acetaldehyde levels were detectable in
control group 1 (59±14 μM), these being significantly reduced (p<0.001) by ciprofloxacin
treatment (Fig. 4). Only traces of acetaldehyde were detectable in the small intestine in any of
the groups.
Folate levels
Rats receiving ethanol had a significantly lower folate intake when assessed by average daily
food intake than the rats receiving saline or ciprofloxacin. There were no significant
differences in serum folate, erythrocyte folate and small intestinal mucosal folate levels among
the four animal groups (data shown in detail in the original article). However, folate levels of
colonic mucosa were significantly reduced in the ethanol-treated group 3, by approximately
50% in comparison to the other three groups (Fig. 4).
0
10
20
30
40
50
60
70
80
Colonic acetaldehyde levels
Colonic folate levels
Group 1
(saline)
Group 2
(saline/cipro)
Group 3
(ethanol)
Group 4
(ethanol/cipro)
p<0.05
Folate level (pM/mg prot)
Acetaldehyde level (x10 μM)
Fig. 4. Colonic acetaldehyde and folate levels in the study groups (mean ± SEM). P value refers to the
corresponding bars of groups 1, 2, and 4. (Reproduced with permisson from Int J Cancer).
52
5.7. FACTORS INFLUENCING SALIVARY ACETALDEHYDE PRODUCTION IN
HUMANS (V)
The various regression analyses clearly showed that smoking and heavy alcohol intake are
powerful predictors of microbial acetaldehyde production. As these factors themselves showed
colinearity, analysis was repeated for all non-smokers (n=189) to estimate the influence
attributable to alcohol intake alone, and for subjects with only moderate or no alcohol
consumption to estimate the influence of smoking alone (n=259). This analysis method
demonstrated that both were independent risk factors of higher salivary acetaldehyde
production. Smoking and heavy alcohol consumption increased the salivary acetaldehyde
production in comparison to the control groups by 60-75%, and combined misuse further
increased it (Fig. 5).
Acetaldehyde production (μmol/l)
0
20
40
60
80
100
120
140
160
180
200
220
Nonsmoker,
moderate
alcohol
(n=173)
Smoker,
moderate
alcohol
(n=82)
Nonsmoker,
heavy
drinker
(n=14)
Smoker,
heavy
drinker
(n=53)
* * *
Fig. 5. Salivary acetaldehyde production in correlation with smoking and drinking. * p < 0.05
versus non-smokers, moderate alcohol consumption. (Reproduced with permission from
Carcinogenesis).
Age (inverse correlation, r=-0.13, p=0.02) and the reported frequency of dry mouth (positive
correlation, r=0.1, p=0.06) were other, less compelling factors possibly associated with
increased acetaldehyde production. Dry mouth also showed co-linearity with smoking and
alcohol intake. After adjustment for confounders, the reported frequency of a dry mouth
showed a slight but significant contribution to salivary acetaldehyde production, at least in
non-smokers. It can be estimated that a subject with very frequent complaints of a dry mouth
has approximately 20% higher salivary acetaldehyde levels than subjects without such
symptoms.
None of the other variables contributed significantly to salivary acetaldehyde production.
Patients with cancer of the oral cavity did not have salivary acetaldehyde production which
differed significantly from the rest of the cohort. This was true both for patients with fresh,
untreated tumours as for patients in follow-up after surgical operation. Patient characteristics
and statistical analysis are shown in detail in the original article.
53
5.8. MICROBES ASSOCIATED WITH ACETALDEHYDE PRODUCTION IN THE
HUMAN ORAL CAVITY (V, VI)
Microbial analysis of the ten saliva samples with the highest and ten with the lowest
acetaldehyde production capacity showed a clear increase in the total count of aerobes among
"high" producers. Aerobic species that were significantly associated with higher acetaldehyde
production were Streptococcus salivarius, the hemolytic viridans group Streptococci,
Corynebacterium sp., Stomatococcus sp. and yeasts. No bacterial species were found to be
significantly more frequent in the saliva of "low" acetaldehyde producers. Yeasts were not only
found at higher concentrations but also more frequently among the ten subjects with high
acetaldehyde production (8 of 10 versus 2 of 10, p=0.02).
Since this analyses revealed that the presence of yeasts could be an important microbiological
factor that may determine an individual’s capacity to produce acetaldehyde from ethanol in
saliva, yeasts were isolated from all the saliva samples that produced acetaldehyde of more
than 250 μM and less than 40 μM. Yeasts colonization was found in 78% of the high
acetaldehyde-producing salivas, compared with 47% in the low acetaldehyde-producing
salivas (p = 0.026). Among the yeast carriers, the number of isolated strains per subject were
similar in both groups. However, the density of yeasts was higher in high than in low
producers (p = 0.025).
Of the 55 subjects, 8 were heavy users of both tobacco (15 to 40 cigarettes/day, mean 23) and
alcohol (78 to 150 g/day, mean 112 g). All of these saliva samples belonged to the high
acetaldehyde producing group, and the presence of yeasts in this group was statistically
significant compared to the group of non-smokers and moderate (<30 grams per day) or nondrinkers
(n=20) (100% in smokers/drinkers vs 45% in controls, p=0.0097). Moreover, the
yeast counts were higher in the group of smokers/drinkers than the controls (1.8 ± 1.3 x105
CFU/ml vs. 3.1 ± 2.6 x 104 CFU/ml, p=0.0081).
Each of the yeast species isolated were able to produce acetaldehyde from ethanol in vitro, and
the acetaldehyde production capacity was in proportion to the length of the incubation, the
quantity of yeasts, and the concentration of ethanol. However, there was a 180-fold difference
in the acetaldehyde production capacity between the strains over 60 min of incubation, ranging
from 1.3 nmol ach/106 CFU to 236.4 nmol ach/106 CFU (Strain dependent data shown in
detail in the original article).
Candida albicans was the dominating yeast species in both groups, constituting 88% of all oral
isolates (87% and 90% among the high and low producing salivas respectively). Moreover, C.
albicans strains isolated from the high acetaldehyde-producing salivas produced significantly
higher acetaldehyde levels from ethanol than C. albicans strains from low acetaldehyde
producing salivas (73.1 nmol ach/106 CFU vs. 43.2 nmol ach/106 CFU, p = 0.035).
54
6. DISCUSSION
6.1. ROLE OF CATALASE IN ACETALDEHYDE PRODUCTION BY COLONIC
CONTENTS
Under microaerophilic or aerobic conditions and in the presence of excess ethanol, some
microorganisms can oxidize ethanol to acetaldehyde. In the past this bacteriocolonic pathway
for ethanol oxidation was primarily thought to be mediated via reversed microbial ADH
reaction (Jokelainen et al., 1996a). However, some members of the Enterobacteriaceae family
appear to produce considerable amounts of acetaldehyde from ethanol with minimal ADH
activity (Jokelainen et al., 1996a). This raised the question of whether enzymes other than
ADH also participate in bacterial acetaldehyde production. A good explanation of this may be
the existence of catalase in these bacteria.
The enzyme catalase is, indeed, present in most cytochrome-containing aerobic and facultative
anaerobic bacteria, and only strict anaerobes lack this enzyme (Deisseroth and Dounce, 1970).
For example, members of the Enterobacteriaceae family like Escherichia coli, Klebsiella
pneumoniae, Klebsiella oxytoca, Proteus mirabilis, Hafnia alvei (Chester and Moskowits,
1987), as well as Helicobacter pylori (Hazell et al., 1991) and a number of yeasts (Fukui et al.,
1975; Tosado-Acevedo et al., 1992) have been shown to possess catalase activity. The
physiological role of catalase in bacteria is to protect aerobes and aerotolerant anaerobes from
the toxicity of oxygen, resulting from its reduction to hydrogen peroxide (H2O2). Catalase
protects bacteria by catalysing the reaction: 2H2O2 2H2O + O2 (Fukui et al., 1975). In
addition to catalysing the breakdown of H2O2 (catalatic activity) the enzyme also has another
function. In the presence of low concentrations of H2O2, it is able to oxidize electron donors
(peroxidatic activity) (Percy, 1984). In the case of ethanol, this reaction yields acetaldehyde
(Oshino et al., 1973). Because catalase has two enzymatic functions it is possible that catalase
may react either catalatically or peroxidatically depending on the microenvironment of the
bacterial cells.
In this study acetaldehyde production by human colonic contents was inhibited by catalase
inhibitors, sodium azide and 3 -amino-1,2,4-triazole, but not with cytochrome P-450 inhibitor,
metyrapone, or the alcohol dehydrogenase inhibitor, 4-methylpyrazole. The poor inhibitory
effect of 4-MP can be explained by its low efficacy on microbial ADHs. Another possibility is
that the amount of NAD in lyophilizised contents or inside colonic bacteria is so slight that the
microbial ADHs are not able to take part in the bacterial acetaldehyde production from ethanol
under these in vitro conditions. The fact that acetaldehyde production was markedly increased
after the addition of exogenous NAD is in favour of the latter possibility.
The amount of acetaldehyde produced increased in the presence of both exogenous NAD or
the H2O2 producing system. Thus in colonic contents there are at least two enzyme systems
that can produce acetaldehyde from ethanol. The aspirated colonic contents are a mixture of
electrolyte solution, sloughed mucosal cells, and mainly colonic bacteria. Moreover, since
lyophilization has been used as a gentle method for the preservation of biological material,
particularly proteins, the enzymes that participate in acetaldehyde production are probably of
bacterial origin. Our findings indicate that in addition to microbial alcohol dehydrogenase,
microbial catalase may also participate in bacterial ethanol metabolism and acetaldehyde
production in the colon.
55
6.2. ROLE OF COLONIC BACTERIA IN EXTRAHEPATIC ETHANOL
ELIMINATION IN HUMANS
The most important route of ethanol elimination is its metabolism in the liver, and only about
one percent is excreted unmetabolised (Holford, 1987). However, extrahepatic elimination of
ethanol occurs as well. In rats, the extrahepatic gastrointestinal metabolism of circulating
ethanol has been shown to be up to 30% of that in the liver (Huang et al., 1993) and in patients
with cirrhosis of the liver, extrahepatic elimination has been estimated to constitute about 40%
of the total ethanol elimination (Utne and Winkler, 1980). Our previous studies show that
bacterial ethanol metabolism in the large intestine may be one of the major extrahepatic
pathways for ethanol oxidation (Jokelainen et al., 1997).
This study showed that one week’s treatment with ciprofloxacin reduces the ethanol
elimination rate in humans by 9.4%. No change in peak ethanol concentration or volume of
distribution was detected. The intravenous administration of ethanol excludes the possible
effects of altered gastric emptying and gastric first-pass metabolism on ethanol
pharmacokinetics (Fraser, 1997). Ciprofloxacin did not inhibit hepatic ADH activity in vitro in
concentrations known to exist in the liver tissue during treatment (Dan et al., 1987). It has,
however, been reported to reduce the hepatic metabolism of coadministered xanthines, such as
theophylline and caffeine, leading to increased serum concentrations and reduced elimination
of these substances (Radandt et al., 1992). The mechanism behind this effect is the inhibition
of CYP1A2 activity (Mizuki et al., 1996). CYP1A2 has been shown to be able to metabolise
ethanol in MEOS, although the major contributor to the MEOS in humans is the isoenzyme
CYP2E1 (Asai et al., 1996). Thus the inhibitory effect of ciprofloxacin on the EER could at
least partly be the consequence of the drug’s interference with cytochrome mediated ethanol
oxidation. Since the contribution of the MEOS to ethanol metabolism, however, is at most 5%
(Ingelman-Sundberg, 1997), the interactions between ciprofloxacin and ethanol oxidizing
enzymes in the liver probably explain only a small part of the reduction in the ethanol
elimination rate. Hepatic metabolism of ethanol may also be reduced because of the changes in
hepatic blood flow. Ciprofloxacin, however, has no effect on the clearance of indocyanine
green, a dye largely extracted by the liver, which indicates the lack of any effect on hepatic
blood flow (Nix et al., 1987). Thus the decrease in ethanol elimination found in this study is
unlikely to be the result of decreased hepatic blood flow.
This study is also in line with our previous findings with rats. Jokelainen et al. (1997) reported
a 9% (p<0.02) reduction in ethanol elimination in rats after four days’ high dose ciprofloxacin
treatment, with a concomitant decrease in faecal aerobic bacteria and ADH activity. The
results of this study suggest that the decrease in the ethanol elimination produced by
ciprofloxacin is at least partly due to the reduction of gut aerobic flora and the consequent
inhibition of ethanol oxidation via colonic bacteria.
The findings from the faecal analysis support this hypothesis. In this study ciprofloxacin
treatment showed a tendency to decrease aerobic flora, with only small changes in the
anaerobic bacteria. Anaerobes that were only slightly affected were Bifidobacterium sp. The
marked change was the complete suppression of the Enterobacteriaceae, which was the
predomint species before the medication. Enterococcus sp., initially present in 63% of
volunteers, also disappeared. The other aerobic species responded variably, and slight
overgrowth of yeasts occurred only in two volunteers of eight. The bacteriological changes in
faecal flora induced by ciprofloxacin are well in line with the previous studies (Campoli-
Richards et al., 1988).
56
This is the first study to show that human stool samples possess ADH and catalase activity and
to produce acetaldehyde from ethanol in vitro. The faecal ADH activity and acetaldehyde
production capacity decreased significantly after one week’s ciprofloxacin treatment, whereas
catalase activity remained unaltered. Furthermore, there was a significant correlation between
faecal ADH activity and acetaldehyde production. Since Enterobacteriaceae has been shown
to produce acetaldehyde from ethanol in vitro (Jokelainen et al., 1996a), these findings suggest
that ethanol oxidation by colonic bacteria in man is probably mediated by Enterobacteriaceae
and ADH-associated reactions.
6.3. THE EFFECT OF LONG-TERM ETHANOL AND METRONIDAZOLE
TREATMENT ON INTRACOLONIC ACETALDEHYDE LEVELS
In the present study, alcohol treatment for 6 weeks led to elevated intracolonic acetaldehyde
levels. Acetaldehyde production was further increased 5-fold after metronidazole treatment.
Since metronidazole has been shown to increase the number of aerobic bacteria in the large
intestine at the expense of the number of strict anaerobes both in experimental animals and
humans (Brook and Ledney, 1994; Krook, 1981), it is an effective drug against anaerobic
bacteria. This was also confirmed by our study, the total count of aerobes and especially that of
Enterobacteriaceae and Staphylococcus species being significantly higher in the caecal
contents of the metronidazole-treated rats than among their corresponding controls. Since
metronidazole treatment did not inhibit the ALDH activity of the colonic mucosa, the increase
in intracolonic acetaldehyde production from ethanol can be explained by the replacement of
intestinal anaerobes by ADH-containing facultative Enterobacteriaceae. These results indicate
the significance of these Gram-negative bacteria in ethanol oxidation and acetaldehyde
production in the large intestine.
We have previously shown marked intracolonic acetaldehyde production from ethanol in
experimental animals after a single dose of ethanol (Jokelainen et al., 1996c; Visapää et al.,
1998). This is the first study to show that intracolonic acetaldehyde levels also remain high
during long-term ethanol intake, indicating that colonic mucosa or bacteria do not adapt, at
least not completely, to removing toxic acetaldehyde during chronic ethanol intoxication.
However, adaptation might occur to a certain extent since colonic mucosal ALDH activity was
significantly higher in the rats receiving ethanol and metronidazole than in the ethanol treated
rats. On the other hand, the quantitative and qualitative changes in the intestinal microflora
after long-term ethanol intake, i.e. the increase in the number of Enterobacteriaceae as shown
in this study, may increase microbial acetaldehyde production. Acetaldehyde accumulation
inside the large intestine has been related to the low ALDH activity of the colonic mucosa and
intestinal bacteria (Koivisto and Salaspuro, 1996; Nosova et al., 1996). In this study colonic
mucosal ALDH activities were four times lower than in the liver, supporting the previous
concept.
Acetaldehyde concentrations in the colon are much higher than in the liver, but the
pathogenetic role of intracolonic acetaldehyde remains still, at least partly, uncertain. Since
ciprofloxacin treatment has been shown to decrease intracolonic microbially-derived
acetaldehyde production from ethanol (Visapää et al., 1998) and metronidazole increases it,
these treatment models can be used as a tool to investigate gastrointestinal morbidity
associated with alcohol and high intracolonic acetaldehyde production.
57
6.4. THE EFFECT OF ETHANOL AND METRONIDAZOLE TREATMENT ON
HEPATIC ETHANOL AND ACETALDEHYDE METABOLISM
Metronidazole did not inhibit hepatic ALDH or ADH activity. Furthermore, blood ethanol and
acetaldehyde levels were similar in ethanol and ethanol and metronidazole receiving rat
groups. These findings suggest that metronidazole treatment has no significant effect on
ethanol metabolism in the rats. This is interesting, since some case-reports and uncontrolled
studies with metronidazole identify disulfiram-like symptoms when taken with ethanol
(Alexander, 1985; Goodwin and Reinhard, 1972; Lehmann et al., 1966; Strassman et al.,
1970). Disulfiram itself blocks hepatic low-Km aldehyde dehydrogenase, which leads to
increased blood acetaldehyde levels if used with ethanol. This causes unpleasant symptoms,
flushing, palpitations, headache, nausea, and sometimes vomiting (Peachey and Sellers, 1981).
The mechanism behind metronidazole related disulfiram-like reaction has been thought to be
similar to that of disulfiram (Lau et al., 1992). However, others have shown that metronidazole
does not increase blood acetaldehyde levels or inhibit ALDH in rats (Kalant et al., 1972;
Vasiliou et al., 1986). Since these findings were confirmed in the present study, there is no
proper explanation for the reported disulfiram-like effects of metronidazole after alcohol
intake. The existence of metronidazole and alcohol interaction is also called into question in
two very recent review articles (Garey and Rodvold, 1999; Williams and Woodcock, 2000).
Despite this, patients should be advised not to consume these two agents together.
Other antimicrobial agents which may have disulfiram-like effects include beta-lactams, e.g.
cephamandole, cefoperazone and moxalactam, with N-methyltetrazolylthiomethyl groups at
the 3-position of the cephalosporin nucleus (Buening et al., 1981; Matsubara et al., 1987; Uri
and Parks, 1983). These drugs have been shown to inhibit liver ALDH activity and to increase
the levels of acetaldehyde in the blood after ethanol intake, leading to a real disulfiram-like
reaction. Our study implies that reported disulfiram-like effects caused by metronidazole and
alcohol intake might develop in a different way. One explanation could be the high
intracolonic acetaldehyde formation from ethanol after metronidazole treatment. At the
periphery, acetaldehyde levels comparable to those found in this study have been shown to
induce histamine release from purified Mast cells (Koivisto et al., 1999) and to depress
histamine elimination, resulting in elevated histamine levels in tissues (Zimatkin and
Anichtchik, 1999). Thus, it is possible that acetaldehyde-induced histamine release may
contribute to various hypersensitivity reactions caused by alcohol ingestion. This is supported
by findings that histamine can mediate the acetaldehyde-induced flushing reaction. Elevated
plasma histamine levels following the administration of histamine correlate well with clinical
symptoms resembling the alcohol-induced flushing in orientals (Zimatkin and Anichtchik,
1999). Thus, it may hypothetically be that the mechanism producing the metronidazole related
disulfiram-like effects associated with alcohol intake are rather located in the large intestine
microflora’s capacity to produce acetaldehyde than in the liver.
There is no data suggesting whether metronidazole inhibits human hepatic ALDH activity. In
humans and rats the isoenzyme mainly responsible for the oxidation of ethanol-derived
acetaldehyde is hepatic mitochondrial ALDH2. Noteworthy, however, is nitrefazole’s ability to
cause a disulfiram-like reaction with elevated blood acetaldehyde level, flushing, tachycardia,
and hypotension after ethanol ingestion (Suokas et al., 1985). Nitrefazole is structurally related
to metronidazole, except that the nitro group in nitrefazole is in position 4 of the imidazole
ring, whereas metronidazole is mainly substituted in position 5 (Klink et al., 1985). It has been
shown to cause a strong and long-lasting inhibition of hepatic mitochondrial ALDH (Klink et
al., 1985). The structural differences between metronidazole and nitrefazole may generate
58
dissimilarities in their ALDH inhibitory potential as with various beta-lactams. There is
evidence that hepatic ALDH2 in rats differs in its sensitivity to disulfiram and nitrefazole
compared to the human one, although both drugs inhibit it in both species (Zorzano and
Herrera, 1990); thus it is possible that human and rat liver ALDHs differ in their sensitivity to
the inhibitory effect of metronidazole as well. One must therefore be cautious in extrapolating
the past and present results obtained with rats to human subjects.
6.5. THE EFFECT OF ACETALDEHYDE ON INTESTINAL FOLATE LEVELS IN
RATS - A POSSIBLE CARCINOGENIC ACTION OF ACETALDEHYDE
High alcohol and low folate intake are independent risk factors for colorectal carcinogenesis
(Giovannucci et al., 1995; Kune and Vitetta, 1992; Longnecker et al., 1990). It has been
suggested that folate deficiency enhances colorectal neoplasia by depleting labile methyl
groups and by inducing DNA hypomethylation (Cravo et al., 1992). Acetaldehyde, a known
carcinogen, has been postulated to be a factor possibly responsible for ethanol-associated
carcinogenesis, since high levels accumulate in the large intestine through the microbial
oxidation of alcohol (Salaspuro, 1996, 1997). Moreover, if high alcohol intake and low-folate
diet are observed, the attributable relative risk of colon cancer increases synergistically in a
multiplicative manner, while in individuals with supraphysiological folate repletion, no
elevated cancer risk from alcohol is observed (Boutron-Ruault et al., 1996; Giovannucci et al.,
1995). Increased alcohol intake and a low-folate diet thus exert a synergistic effect on
colorectal carcinogenesis, and are influenced by each other (Anonymous, 1994).
One explanation for this could be the high intracolonic acetaldehyde levels formed after
ethanol administration, since it has been shown that high acetaldehyde levels can cleave folate
to inactive forms via acetaldehyde/xanthine oxidase-generated superoxide (Shaw et al., 1989).
This interesting in vitro mechanism, by which alcohol leads to folate deficiency, has often
been considered insignificant in vivo as the concentrations of acetaldehyde have been thought
to be too low. However, as shown in this study, the local intracolonic acetaldehyde levels may
reach the required level. Low vapour pressure and high water solubility (Matysiak-Budnik et
al., 1996) enable most acetaldehyde in the colonic lumen to reach the colonic mucosa (Seitz et
al., 1990). The observed decreased folate levels of colonic mucosa in ethanol-treated rats are
thus probably caused by the cleavage of folate through high intracolonic levels of acetaldehyde
produced from ethanol by gut microbes.
The involvement of high colonic acetaldehyde in local folate decrease is also supported by the
fact that the effect of alcohol on mucosal folate levels was effectively attenuated by a
concomitant treatment with ciprofloxacin, which reduced the intracolonic acetaldehyde levels
to 5% of that found after treatment with alcohol alone. Moreover, acetaldehyde levels in the
small intestine were very low, and only the folate level of the colonic mucosa, not small
intestinal mucosa, was decreased by alcohol. Low acetaldehyde in the small intestine can be
explained by the fact that small intestine is colonized by fewer bacteria than the large intestine
(Rowland, 1986). As indicated by normal serum folate levels and by the sufficient folate
uptake of the rats treated with ethanol decreased nutritional intake cannot explain our findings.
Mechanisms other than this antimicrobial effect of ciprofloxacin on acetaldehyde production
from ethanol are unlikely to play a significant role, as there were no differences in the colonic
folate levels between the control-saline group and the control-ciprofloxacin group.
59
6.6. ACETALDEHYDE IN SALIVA: INFLUENCING FACTORS
Although alcohol and tobacco smoke are well-known independent risk factors for upper
gastrointestinal tract cancer, their combined action on these epithelia has remained poorly
understood. There is epidemiological evidence indicating that alcohol and tobacco act together
in a multiplicative rather than additive manner and, accordingly, seem to have synergistic
tumor-promoting effects (La Vecchia et al., 1997). As alcohol is involved synergistically in the
attributable risk of both smoking and poor oral hygiene, it is conceivable that there may be a
unifying pathogenetic mechanism behind these epidemiological findings. This may be the
local production of carcinogenic acetaldehyde from ethanol by oral microbes. Salivary
acetaldehyde may reach all target tissues of the upper aerodigestive tract, including the larynx,
pharynx, oral cavity, and esophagus, via normal distribution and evaporation.
In the present study, we were able to demonstrate that smoking and heavy alcohol
consumption significantly increase salivary acetaldehyde production. Smoking showed a
positive linear correlation and it can be estimated that a smoker with a daily consumption of
approximately twenty cigarettes has an increased salivary acetaldehyde production of about
50-60%. This implies that smokers, even after moderate alcohol intake, produce much higher
levels of carcinogenic acetaldehyde in the oral cavity than non-smokers. The evidence for
increased microbial salivary acetaldehyde production in smokers, together with the
epidemiological description of the multiplicative carcinogenic action of alcohol and smoking,
suggests that the salivary acetaldehyde production mediated by microbes could be the
biologically plausible pathogenetic mechanism for these findings. Alcohol seems to interact
and increase salivary acetaldehyde production only if consumed heavily; when an increase is
observed it is dose dependent. Smoking and alcohol together increase the salivary
acetaldehyde production by about 100% as compared to non-smokers and moderate alcohol
consumers.
6.7. MICROBES ASSOCIATED WITH ACETALDEHYDE PRODUCTION IN THE
HUMAN ORAL CAVITY
As stated above, one possible pathogenetic explanation underlying the joint effect of tobacco
smoking and alcohol drinking on oropharyngeal carcinogenesis could be the increased local
production of acetaldehyde from ethanol by oral microbes. This however could also be due to
an altered oral microflora producing more acetaldehyde.
This theory is supported by the literature and our results. Smoking has been shown to increase
the number of yeasts in the oral flora (Sakki and Knuuttila, 1996). In general, a microbial
"switch" with a significant increase in the proportion of Gram-positive versus Gram-negative
bacteria has been described in smokers (Colman et al., 1976; Macgregor, 1988), whereas
Neisseriae spp. have been reported to occur less frequently in the oral cavity of smokers
(Colman et al., 1976). Although, microbial changes in the oral microflora of alcoholics have
not been described, slight immunodeficiency associated with alcoholism together with poor
oral hygiene and nutritional defects may favour the growth of bacteria and Candida albicans in
the oral cavity (MacGregor, 1986; Oksala, 1990). These are well in line with our observation
that almost all aerobic Gram-positive bacteria were significantly increased in "high"
acetaldehyde producers (the facultative commensals Staphylococcus sp. and Streptococcus
mutans being the only exceptions), whereas the Gram-negative aerobic bacteria, Haemophilus
60
sp. and the already-mentioned Neisseria sp. were not associated with higher acetaldehyde
production. Thus, there is in general a good link between our microbial observations that some
species are associated with higher acetaldehyde production and the well-known effect of
smoking on the oral microflora.
Moreover, our microbial analyses revealed that yeasts (and possibly Corynebacterium sp.)
were found in higher loads and more frequently in the high acetaldehyde-producing saliva
group than the low group. Since Candida albicans was the dominating yeast species isolated
from the saliva and C. albicans produced on average higher acetaldehyde levels than other
yeast species detected, this can be regarded as the most common and important yeasts species
with respect to acetaldehyde production capacity from ethanol. This is further supported by the
finding that C. albicans strains isolated from the high acetaldehyde-producing salivas formed
significantly higher acetaldehyde levels than those isolated from the low acetaldehydeproducing
salivas.
C. albicans is an aerobic microorganism exhibiting ADH activity and it is closely
associated with mucosal membranes. C. albicans colonization is also often present in the
case of oral cancer, but whether these yeasts are causally involved in the development of
this cancer is still not clear (Krogh et al., 1987; Sciubba, 1995). Oral leukoplakia is a
premalignant transformation often invaded by yeasts, and if untreated 5-10% of the cases will
develop into carcinoma. C. albicans infection together with simultaneous existence of several
etiological factors seem to play a role in the malignant transformation (Banoczy, 1977; Krogh
et al., 1987). It has been hypothesized that certain Candida types from oral leukoplakia have
higher nitrosation potential than others, which might indicate the involvement of specific yeast
types in the transformation of leukoplakia into carcinoma (Krogh et al., 1987). In the light of
our hypothesis, an additional plausible etiological explanation could be alcohol drinking and
consequent high acetaldehyde production via reversed ADH-mediated reaction by certain
Candida albicans strains.
61
7. SUMMARY AND CONCLUSION
The key findings of the present study were:
1. These in vitro results demonstrate that in addition to alcohol dehydrogenase (ADH), colonic
contents and faeces exhibit catalase activity, which probably is of bacterial origin. This
indicates that part of the acetaldehyde produced in the large intestine via the bacteriocolonic
pathway for ethanol oxidation may be catalase dependent. However, it is probable that the role
of catalase-dependent acetaldehyde production by large intestinal microflora is limited, and the
main source of acetaldehyde is via the reversed ADH reaction.
2. Ciprofloxacin treatment decreases the ethanol elimination rate by a mean of 9.4% in man,
with a concomitant decrease in faecal ADH activity and acetaldehyde production in vitro.
Since there is no evidence that ciprofloxacin interferes with hepatic ethanol metabolism, our
findings can be explained by the reduction of aerobic and facultative anaerobic bacteria in the
lumen and mucosal surfaces of the human large intestine. These findings support the evidence
for colonic bacteria having a significant, and at least approximately 10% role in the
extrahepatic ethanol elimination in humans.
3. Ciprofloxacin treatment effectively reduced the intracolonic acetaldehyde production from
ethanol in rats to 5% of that found after treatment with ethanol alone. In the human study,
ciprofloxacin effectively eradicated Enterobacteriaceae from the stool and reduced faecal
ADH activity. Before medication these bacteria were the main aerobic species in the faeces,
and it has previously been shown that isolated Enterobacteriaceae are able to produce
acetaldehyde from ethanol in a reaction catalysed by bacterial ADH in vitro. Moreover,
metronidazole and ethanol treatment of rats increased acetaldehyde production in the large
intestine to 5 times that of rats treated with ethanol only. Since this was associated with an
increase in caecal aerobic flora, especially Enterobacteriaceae and Streptococcus species, it
can be concluded that facultative Gram-negative Enterobacteriaceae are probably responsible
for acetaldehyde production in the large intestine.
4. We have already demonstrated intracolonic acetaldehyde production from ethanol in
experimental animals after a single dose of ethanol. Our previous results show that
intracolonic acetaldehyde levels also remain high during long-term ethanol intake. This
indicates that colonic mucosa or bacteria do not adapt to remove acetaldehyde during longterm
ethanol intoxication. Since acetaldehyde is toxic and carcinogenic, the knowledge that
acetaldehyde levels remain high even after long-term alcohol use implies that microbially
produced acetaldehyde may play a role in the gastrointestinal symptoms and morbidity
associated with chronic alcohol use.
5. High alcohol intake leads to local folate deficiency in rat colonic mucosa, probably via the
high levels of microbially produced acetaldehyde. Our preliminary results suggest that
microbial production of acetaldehyde from ethanol also occurs in vivo in the human colon.
Accordingly, these results suggest a possible mechanism by which the unique and synergistic
effects of high alcohol and low folate intake on colorectal carcinogenesis might be explained.
6. Tobacco smoking and alcohol consumption are the most potent external risk factors for
upper digestive tract cancer. In this study we demonstrated an increased microbial salivary
acetaldehyde production associated with these conditions. Moreover, our results demonstrate
62
that almost all aerobic Gram-positive bacteria and Candida albicans were found in
significantly higher numbers in the saliva among high acetaldehyde producers. Numerous
studies support the hypothesis that acetaldehyde is the substance behind the tumor-promoting
effect of alcohol on the mucosa of the oral cavity. Our findings thus provide a biologically
plausible mechanism for the synergistic and multiplicative manner in which the attributable
cancer risks of alcohol and smoking act.
63
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