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Digitally Signed by: Content manager’s Name
DN : CN = Weabmaster’s name
O= University of Nigeria, Nsukka
OU = Innovation Centre
Nwamarah Uche
UNIVERSITY OF NIGERIA, NSUKKA
ALCOHOL DEHYDROGENASE
Onah Donatus
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CHAPTER ONE
INTRODUCTION AND LITERATURE REVIEW
1.0. Introduction
Palm wine is an important alcoholic beverage resulting from the spontaneous fermentation of the
sap of the palm, which has been attributed to yeast and bacteria (Onwuka, 2011; Opara et al.,
2013). It is the fermentad sap of certain varieties of palm trees including raphia palm (Raphia
hookery or R. vinifera) (Ali, 2008). Fresh palm wine is sweat, clear, neutral, colourless juice
containing minimal sugar (less than 0.5%) small amount of protein, gums and minerals (Opara et
al., 2013). According to Oyeku et al. (2009), it consists mainly of water, sugar, vitamins and
many aroma and flavour components in very small amounts. In traditional African societies, the
palm wine play a significant role in customary practices, especially the distilled product from the
palm wine, a potent gin called by various names in West Africa (Amoa-Awua et al., 2006).
Over ten million people consume palm wine in West Africa (Onwuka, 2011)
Traditionally, it is believed that when taken by nursing mothers; palm wine stimulates lactations
and also has diuretic effect. Palm wine has also been used to enhance men’s potency due to yeast
cell concentration. It could also be used for leavening of dough ad used in African medicine
particularly in the treatment of measles and malaria (Onwuka, 2011).
Despite all these good qualities of palm wine, it is a highly perishable sap due to fermentation
which starts soon after the sap is collected and within an hour or two becomes reasonably high
in alcohol (up to 4%). If palm wine is allowed to continue to ferment for more than 24hrs, it
starts to turn into vinegar. This makes it unacceptable to consumers and creates losses to the food
service industries. Fermentation in palm wine is possible because it constitutes a good growth
medium for numerous microorganisms especially for yeast, lactic acid and acetic acid bacteria
(Bechem et al., 2007). Saccharomyces cerevisae constitutes about 70% of the total yeast of palm
wine and the activities of these microbes are believed to be responsible for conversion of sugar
in palm wine to alcohol after a short time while bacteria induces the conversion of alcohol into
vinegar (Onwuka, 2011).
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Authorities who have studied the succession of micro flora in palm wine consistently reported
the emergence of Acetobacter after about 24hrs of fermentation, at which time, alcohol was
present in reasonable quantities (Opara et al., 2013). Earlier researches on the microbiology of
palm wine had isolated Acetobacter from palm wine and these bacteria are believed to be
responsible for souring of palm wine which is not acceptable by many.
Acetic acid bacteria, Acetobacter and Gluconobacter, as well known as vinegar producers are
able to oxidize ethanol to acetic acid by two sequential catalytic reactions of alcohol
dehydrogenase and aldehyde dehydrogenase which are located on the periplasmic side of their
cytoplasmic membrane (Abolhassan et al., 2007). Though these enzymes are important in
industrial production of acetic acid, they are nevertheless spoilage molecules for many types of
food and juices including palm wine (Ameh et al., 2011).
Many attempts have been made to control palm wine spoilage at microbial level (Enwefa et al.,
2004). Locally, the rural people use special leaves such as bitter leave to cover the wine
container which they believe kills or disallows the influx of microorganism into the wine.
Unfortunately, this method do not take care of the organisms already in the wine itself, hence
deterioration continues (Onwuka, 2011). Also, the bark of a tree S. gabonensis was often added
to the fresh palm wine. This impacts an amber colour and bitter taste to the wine. Although it
delays souring of the wine and also lowers the titrable acidity (Ojimelukwe, 2002), the extract
could not inhibit several yeast and bacteria present in the wine. With increasing availability of
modern methods, efforts were directed towards the use of chemicals and pasteurization. Attempt
to preserve palm wine using sulfite failed because at this pH, the concentration of sulfite required
to suppress microfloral activities would be excessive for human consumption. Moreover, the use
of chemical preservatives are discouraged due to the belief of cancer promotion. Convectional
heating methods have been employed to delay spoilage, but the attractive flavor of palm wine is
destroyed, giving room for arguments between wine drinkers and service men on the freshness of
the beverage.
Currently, palm wine is bottled on commercial scale with 37.5mg/l of metabisulphite and
pasteurized at 65oC for 35mins, but the search for a more convenient, safer and effective method
must continue.
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1.1. Acetobacter
Scientist have advocated the control of biological activities at molecular level because of its
safety and the purity of the products. This draws our attention to alcohol dehydrogenase, one of
the enzymes in Acetobacter responsible for deterioration of palm wine by converting alcohol, the
most wanted component of palm wine, into acetic acid, with a view to investigating the
possibility of controlling palm wine spoilage at the enzyme level. This entails that the enzyme is
isolated, purified and characterized and that the effect of such parameter as pH, temperature and
ethanol concentration on the activity of alcohol dehydrogenase are investigated.
The knowledge of the effect of these parameters on the activity of alcohol dehydrogenase will be
indispensible in regulating the activity of alcohol dehydrogenase. In this study, alcohol
dehydrogenase was extracted from Acetobacter which was isolated from palm wine, partially
purified, characterized and its thermal and pH stability investigated.
Since their first discovery and reporting as a unique group, the acetic acid bacteria (bacteria that
produce acetic acid) have been labeled with numerous genetic names, which have been the
subject of extensive discussion and revision. The eighth edition of Berger’s Manual of
Determinative Bacteriology (Buchanan and Gibbons, 1974) recognized only two genera,
Acetobacter (motile by petrichous flagella or non-motile) and Gluconobacter (motile by polar
flagella or nonmotile), and placed the genus Gluconobacter with the family Pseudomonadaceae;
however, the genus Acetobacter was not assigned to any particular family and was grouped
within the genera of uncertain affiliation. The Approved List of Bacterial Names, (Skerman et
al., 1980) acknowledged both the genera Acetobacter and Gluconobacter. The ninth edition of
Bergey’s Manual of Systematic Bacteriology (Buchanan and Gibbons, 1984) recognized the fact
that the genera Gluconobacter and Acetobacter were closely related; hence they were placed
within the family Acetobacteraceae. Members of the family are united by their unique ability to
oxidize ethanol to acetic acid. Under this family we have genera Acetobacter, Gluconobacter and
Frateuria. Today, acetic acid bacteria have been classified into 24 different genera. The major
genera involved in vinegar production include: Acetobacter, Gluconobacter, Gluconacetobacter,
Asaia, Neoasaia, Saccharibacter, Frateuria and Kozakia (De Vero and Giudici, 2008).
The microorganisms present in wine-making processes are mainly yeasts, lactic acid
bacteria and acetic acid bacteria, because of the extreme conditions in grape must (juices before
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or during fermentation) such as the low pH (between 3 and 4) or high sugar concentration.
Saccharomyces species (mainly Saccharomyces cerevisiae) are responsible for converting the
sugars in grape must into ethanol and CO2 (Drysdale and Fleet, 1988).
Lactic acid bacteria decrease the acidity of the wine and convert malic acid into lactic
acid and CO2. This is a one-step reaction known as malolactic fermentation, which usually takes
place once the alcoholic fermentation is over (Ribereu-Gayon et al., 2002).
Acetic acid bacteria (AAB) play a negative role in the wine-making process because they
alter the organoleptic characteristics of the wine and, in some cases, can also lead to stuck and
sluggish fermentation. AAB modify wine, mainly because they produce acetic acid, acetaldehyde
and ethyl acetate. They are also involved in other industrial processes of considerable interest for
biotechnology such as the production of cellulose, sorbose and vinegar (Du Toit and Pretorius,
2002).
Acetic acid bacteria can be found in different stages of the wine-making process: for
example, grape ripening, must, alcoholic fermentation, and bottled and stored wine. Although it
has been known that wine can be altered by acetic acid bacteria ever since Pasteur, and they have
a highly undesirable impact on the alcoholic fermentation processes, relatively little is
understood about how they behave. Other microorganisms such as yeasts and lactic acid bacteria
are also present during alcoholic fermentations and have been studied in much greater depth.
1.2. General Characteristics of acetic acid bacteria
Acetic acid bacteria (AAB) are gram negative, ellipsoidal (regular oval) to rod-shaped,
and can occur singly, in pairs or in chains. They are motile due to the presence of flagella which
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can be both peritrichous (having flagella uniformly distributed over the body surface, as certain
bacteria) or polar (ie when the flagellum is located at one end of the cell). They do not form
endospores as a defensive resistance. They have obligate aerobic metabolism, with oxygen as the
terminal electron acceptor. The optimum pH for the growth of AAB is 5-6.5 (Holt et al., 1994).
However, these bacteria can grow at lower pH values of between 3 to 4. They vary between 0.4-
1µm long. They are catalase positive and oxidase negative. AAB can present pigmentation in
solid cultures and can produce different kinds of polysaccharides (De Ley et al., 1984)
AAB occur in sugar and alcoholised, slightly acid niches such as flowers, fruits, beer,
wine, cider, vinegar, souring fruit juices and honey. On these substrates, they oxidize the sugars
and alcohols, resulting to an accumulation of organic acids as final products. When the substrate
is ethanol, acetic acid is produced, and this is where the name of the bacterial group comes from.
However, these bacteria also oxidize glucose to gluconic acid, galactose to galactonic acid,
arabinose to arabinoic acid. Some of these transformations carried out by AAB are considered
interest for the biotechnological industry. The best known industrial application of AAB is
vinegar production but they are also used to produce sorbose, from sorbitol, and cellulose.
Fig 1. Electron microscope photography of Acetobacter (Gonzalex et al., 2004)
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1.3. Physiological Role of Acetic Acid Bacteria
One of the main characteristics of AAB is their ability to oxidize a wide variety of
substrates and to accumulate the products of their metabolism in the media without toxicity for
the bacteria. This ability is basically due to the dehydrogenase activity in the cell membrane.
These dehydrogenases are closely related to the cytochrome chain (Matsushita et al., 1985).
1.3.1. Ethanol Metabolism
The oxidation of ethanol to acetic acid is the best known characteristics of acetic acid
bacteria. Ethanol oxidation by AAB takes place in two steps. In the first one, ethanol is oxidized
to acetaldehyde and in the second step acetaldehyde is oxidized into acetate. In both reactions,
electrons are transferred and these are later accepted by oxygen.
Two enzymes play a critical role in this oxidation process, both of which are bound to the
cytoplasmic membrane: they are alcohol dehydrogenase and aldehyde dehydrogenase. Both
enzymes have their active site on the outer surface of the cytoplasmic membrane (Adachi et al.,
1978; Saeki et al., 1997).
The bacteria can produce high concentration of acetic acid, up to 150g/l (Sievers et al.,
1996; Lu et al., 1999), which makes them very important to the vinegar industry. Their
resistance is strain dependent (Namba et al., 1984). The enzyme citrate synthase plays a key role
in this resistance, because it detoxicates acetic acid by incorporation into the tricarboxylic or
glyoxylate cycles, but only when ethanol is not present in the media. According to the report of
Menzel and Gottschalk (1985), Acetobacter strain decrease their internal pH in response to a
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lower external pH. However, an adaptation to high acetate concentration seems to be a
prerequisite for high tolerance (Lasko et al., 2000).
1.3.2. Primary and Polyalcohol Metabolism
A considerable number of AAB can oxidize alcohols into sugars; mannitol into fructose;
sorbitol into sorbose or eritritol into eritrulose. An important ability in oenology is to use
glycerol as a carbon source (De Ley et al., 1984), which is converted into dihydroxyacetone, a
small amount of which is used for energy synthesis.
The enzymes that catalyse all these reactions are located in the cell membrane and induce a high
accumulation of substrates in the media, which make AAB suitable microorganisms for the
biotechnological industry (Deppenmeirer et al., 2002)
1.3.3. Carbohydrate Metabolism
AAB can metabolise different carbohydrates as carbon sources. Acetobacter species can
use sugars through the hexose monophosphate pathway (Warburg-Dickens pathway) (De Ley et
al., 1984; Drysdale and Fleet, 1988). And also through the EMbden-Meyerhof-Parnas and
Entner-Doudoroff pathways (Attwood et al., 1991), although such authors as Drysdale and Fleet
(1988) say that this last pathway is not used by AAB to metabolise glucose. From here they are
further metabolized to CO2 and water via the tricarboxylic acid pathway, which is not functional
in Gluconobacter species, although the complete oxidation is only functional when there is no
carbon source in the media.
Glycerol dihydroxyactetone
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Sugar is more preferred as a carbon source by Gluconobacter than by Acetobacter because the
species of this genus can obtain energy more efficiently by the metabolisation of the sugar via
pentose phosphate pathway (De Ley et al., 1984).
Glucose metabolism by these species produces a considerable number of industrially important
metabolites (Olijve and Kok, 1979; Weenk et al., 1984; Qazi et al., 1991; Qazi et al., 1993;
Velizarov and Beschkov, 1994). Some of these metabolites are 2-ketogluconic, 5-ketogluconic
and 2,5-diketogluconic acids.
The most characteristic reaction is the direct oxidation of glucose into Glucono-δ-lactone, which
is oxidized into gluconic acid. This last reaction is particularly active in Gluconobacter species in
media with high concentration of sugars such as grapes and must. This metabolite can be used as
an indicator of the presence of these bacteria.
Acetic acid bacteria can also use other carbohydrate, such as arabinose, fructose, galactose,
mannitol, mannose, ribose, sorbitol and oxylose (De Ley et al., 1984) (Fig. 1)
1.3.4. Organic Acid Metabolism
AAB are able to metabolise a variety of organic acids. They do so through the tricarboxylic acid
cycle which oxidizes these acids to CO2 and water. Gloconobacer, which lacks a functional
tricarboxylic acid cycle, is unable to oxidize most organic acids (Holt et al., 1994). Acetic, citric,
fumaric, lactic, malic, pyruvic, and succinic acids are completely oxidized to CO2 and water.
These changes are very important in wine making, because they mean that the quality of the
wines decrease.
Another important by product of lactate metabolism is acetoin (important in the world of
oenology (the scientific study of all aspect of wine and wine making) (De Ley, 1959). The
buttery aroma of this compound is considered to be an unwanted flavor in wine, in which its
detection limit is 150g/l (Romano and Suzzi, 1996; Du Toit and Pretorius, 2000).
1.3.5. Nitrogen Metabolism
Although some AAB species (Gluconacetobacter diazotrophicus) (Gillis et al., 1989) can fix
atmospheric nitrogen, most of them use ammonium as a carbon source (De Ley et al., 1984). So
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these bacteria can synthesize all the amino acids and nitrogenated compounds from ammonium.
Depending on the amino acid in the media, their growth can either be stimulated or inhibited. So,
glutamate, glutamine, proline, and histidine stimulate the growth of AAB, whereas valine for
Gluconobacter oxidans and threonine and homoserine for Acetobacter aceti seem to have an
inhibitory effect (Belly and Claus, 1972). However, no studies have been made on the nutritional
needs of AAB nitrogen in wine. It has been observed that AAB selectively prefer some amino
acids during vinegar production (Valero et al., 2003), and leave significant amount of
ammonium in the media.
1.4. History of Acetobacter
The first taxonomist of AAB is the French scientist, Pasteur. Studying the Orleans method of
vinegar production, he demonstrated that the acetic acid came from ethanol oxidation and that
long-term oxidation of acetic acid converted it into CO2 and water. His results led him to
formulate the involvement of the microorganisms in the process of transforming alcohol into
vinegar, and confirmed the existence of Mycoderma aceti which Persoon had already described
in 1882. Subsequently, in the year 1879 Hansen observed that the microbial flora which
converted alcohol into acetic acid was not pure and consisted of various bacterial species. It was
through the work of Beijerinck (1899) that the genera Acetobacter was proposed.
1.5. Taxonomy of Acetobacter and other acetic acid bacteria
The first classification was proposed by Hansen in 1894, based on the occurrence of a film in
the liquid media, and its reaction with iodine. Asai (1934) formulated the proposal of classifying
AAB into two genera: Acetobacter and Gluconobacter. The main differences between these two
genera were both cytological (based on the cell bacterial cell structure, function and formation)
and physiological (the scientific study of an organism’s vital functions, including growth and
development, the absorption and processing of nutrient, the synthesis and distribution of proteins
and organic molecules, and the functioning of different tissues, organs and other anatomic
structures). The main physiological difference was that Acetobacter oxidized ethanol into acetic
acid and, subsequently, completed the oxidation of acetic acid into water and CO2.
Gluconobacter species, on the other hand, were unable to complete this oxidation of acetic acid.
It was Frateur (1950) who formulated a classification based mainly on five physiological criteria:
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1. Presence of catalase
2. Gluconic acid production from glucose
3. Oxidation of acetic acid into CO2 and water
4. The oxidation of lactate into CO2 and water and
5. The oxidation of glycerol into hydroxyacetone
On the bases of these criteria, he divided Acetobacter genera into four groups
1. Peroxydans
2. Oxydans
3. Mesoxydans and
4. Suboxydans
Those AAB that had peritrichous flagella and were able to completely oxidize ethanol into
CO2 and water, are grouped into the genus Acetobacter while those that had polar flagella and
unable to perform the complete oxidation are grouped into the genera Gluconobacter. The
taxonomical keys for bacteria taxonomy have been historically collected in Bergey’s Manual of
Systematic Bacteriology. In its last edition (De Ley et al., 1984), some molecular techniques
were included as fatty acid composition, soluble protein electrophoresis, percentage of G + C
content, and DNA-DNA hybridization. Gluconobacter and Acetobacter genera were included in
the family of Acetobacteraceae. Acetobacter genus was composed by 4 species: A. aceti, A.
pasteurianus, A. liquefaciens and A. hansenii. The Gluconobacter genus only consisted of G.
oxydans.
1.5.1. Taxonomy Based on Molecular Techniques
Classification of AAB based initially on morphological and physiological criteria has been
submitted to continuous variation and reorientations. These variations are due, basically, to the
application of molecular techniques to the taxonomic study. DNA-DNA hybridization,
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percentage base ratio determination, and 16S rDNA sequence analysis are the most common
techniques used for this purpose.
1.5.2. DNA-DNA Hybridization
From taxonomical point of view, this is the most widely used for describing new
species within bacterial groups. The technique measures the degree of similarity between the
genomes of different species. When several species are compared in this way, the similarity
values make it possible to arrange the species in a phylogenetic tree, which shows the degree of
intraspecific and interspecific similarity.
1.5.3. Percentage Base Ratio Determination
This was one of the first molecular tools to be used in bacterial taxonomy. It calculates
the percentage of G + C in a bacterial genome (G for guanine and C for cytosine. Guanine and
adenine are nitrogenous bases in DNA). Although this percentage must be taken into
consideration, by itself it cannot identify a given microorganism. In AAB, the % value of G + C
vary between 55.5 and 64.5%.
1.5.4. 16S rDNS Sequence Analysis
The 16S rDNA gene is a highly preserved region with small changes that can be characteristic
of different species. Ribosomal genes are compared in most taxonomical studies of bacteria.
Acetobacteraceae family is no exception in this reorganization of species and genera. Six
new AAB genera have been added to both the Acetobacter and Gluconobacter genera mentioned
above. These include
1. Acidomonas (Urakami et al., 1989)
2. Gluconacetobacter (Yamada et al., 1997)
3. Asaia (Yamada et al., 2000)
4. Saccharibacter (Jojima et al., 2004)
5. Swminathania (Loganathan and Nair, 2004) and
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6. Kozakia (Lisdiyanti et al., 2002)
At present, the Acetobacteriaceae family consists of 8 genera and 38 species. It has been
proposed that the following species should be added to what was previously established by
Bergey’s (De Ley et al., 1984). These are Acetobacter cerevisiae, A. malorum (Ceenwerck et al.,
2002), A. tropicalis, A. orleaniensis, A. lovaniensis, and A. estuniensis, A. syzgii, A.
cibinongenesis and A. oreintalis (Lisdiyanti et al., 2001), A. pomorum and A. oboediens
(Sokollek et al., 1998), A. intermedians (Boesch et al., 1998), Kozakia baliensis (Lisdiyanti et
al., 2002), Gluconobacter johannae and Ga. azotocaptuans (Fuentes-Ramirez et al., 2001), Ga
swigsii and Ga. rhaeticus (Cleenwerck et al., 2005) and Ga. sacchari (Franke et al., 1999), Asaia
krungthepensis (Yukuphan et al., 2004), As. siamensis (Katsura et al., 2001), As. bogorensis
(Yamada et al., 2000), Saccaribacter floricola (Jojima et al., 2001), Swaminathania salitolerans
(Loganathan and Nair, 2004). A. oboediens and A. intermedius were subsequently reclassified as
Glucoacetobacter by Yamada (2000).
These are summarized in table 1.
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Table 1. Species of acetic acid bacteria
Acetobacter Gluconacetobacter Gluconobacter
A. cerevisiae,
A. malorum
A. tropicalis,
A. orleaniensis,
A. lovaniensis
A. estuniensis,
A. syzgii,
A. cibinongenesis
A. oreintalis
A. pomorum
A.aceti
A. pasteurianus
A.indonosiensis
A. peroxydans
Ga. johannae
Ga. azotocaptans
Ga swigsii
Ga. rhaeticus
Ga. Sacchari
Ga. hansenii
Ga. entanii
Ga. xylinus
Ga. liquefaciens
Ga. diazotrophicus
Ga. europaeus
Ga. Oboediens
Ga. intermedius
G. oxydans
G. frateurii
G. assaii
Asaia
As. Bogorensis
As. Siamensis
As. Indonesiensis
As. rugthepensis
Swaminathania
S. salitolerans
Acidomonas Kozakia
Ac. methanolica K. baliensis
Saccharibacter
Sa. floricola
Gonzalex et al., 2004
1.6. Isolation of Acetobacter
These physiological difference among genera made it possible to develop differential
culture media. Various media have been reported for isolating AAB whose carbon source is
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glucose, mannitol, ethanol etc. some of these media can also incorporate CaCO3 or bromocresol-
green as acid indicators (Swings and De Ley, 1981; De Ley et al., 1984; Drysdale and Fleet,
1988). Culture media are usually supplemented with pimaricin in the agar plates to prevent the
yeast from growing and with penicillin to eliminate lactic acid bacteria.
Most of the widely used culture media are GYC (5% D-glucose, 1% yeast extract, 0.5%
CaCO3 and 2% agar (w/v), described by Carr and Passmore (1979), and, YPM ( 2.5% mannitol,
0.5% yeast extract, 0.3% peptone and 2% agar (w/v)). Plates must be incubated for between 2 to
4 days at 28oC under aerobic conditions. These culture media are suitable for wine samples (Du
Toit and Lamberchts, 2002; Bartowsky et al., 2003), and no problems have been detected
culturing and isolating AAB from wine samples.
Nevertheless, some works indicate the difficulty of culturing this bacterial groups from
vinegar samples (Sokollek et al., 1998). This problem has been partially solved by introducing a
double agar layer (0.5% agar in the lower layer and 1% agar in the upper layer (w/v) into the
cultures and media containing ethanol and acetic acid in an attempt to stimulate the atmosphere
of the acetification tanks, such as AE medium (Entani et al., 1985).
1.7. Identification of acetic acid bacteria
Identification of acetic acid bacteria is done using either classical method or molecular
techniques.
1.7.1. Classical Methods
Classical microbiological taxonomy has traditionally used morphological and
physiological differences among the species to discriminate between them. The tests could
only discriminate at the species level, although the physiological methods would not be able
to distinguish the currently described species. At the genus level, several characteristics can
contribute to the differentiation. The Gluconobacter genus cannot completely oxidize acetic
acid into CO2 and water. The main characteristic of Acidomonas is that it can grow in
methanol, and Asaia is characterized by its inability to grow in a media with an acetic acid
concentration higher than 0.35%. The other two genera, Gluconacetobacter and Acetobacter,
can be differentiated on the bases of their ubiquinone content. Ubiquinone Q9 is present in
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Acetobacter, and ubiquinone Q10 in Gluconacetobacter (Trcek and Teuber, 2002). Kozakia
have low similarity values of % G + C content among the other genera (7 – 25% lower than
the other species), the major ubiquinone is Q10 and have a weak activity in oxidation of
lactate and acetate into carbon dioxide and water. The genus Saccharibacter has a negligible
or very weak productivity of acetic acid from ethanol and the osmophilic growth properties
(its adaptation to environment with high osmotic pressure, such as high sugar concentration)
distinguished this genus from other AAB. Swaminathania genus is able to fix nitrogen and
solubilized phosphate in the presence of NaCl. Some of the phenotypic characteristics of the
former species described in Bergey’s Manual are shown in Table 2.
Table 2. Phenotypic characteristics of the species belonging to the Acetobacter and
Gluconobacter
Characteristics A. aceti A. liquefaciens A. pasteurianus A. hansenii G. oxydans
Ethanol overoxidation + + + + _
Growth in:
ethanol + + V _ _
Sodium acetate + V V _ _
Dulcitol _ _ _ _ _
Glycerol Cetogenesis + + _ V V
Lactate oxidation + + + + _
Pigment production _ + _ _ +
Source: Gonzalex et al., 2004
1.7.2. Molecular Techniques
There are many molecular techniques of identifying AAB both on species level and on
strain level. One of them is PCR-RFLP of the rDNA 16S method. This technique was used by
Ruiz et al. (2000) to identify AAB and is appropriate for differentiating and characterizing
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microorganisms on the basis of their phylogenetic relationships (phylogenetic analysis exploits
the changes in DNA sequence that arise through mutations during evolution to reconstruct the
evolutionary history of different groups of organisms) (Carlotti and Funke, 1994). In eubacterial
DNA, the RNA loci include 16S, 23S and 5S rRNA gene, which are separated by internally
transcribed spacer (ITS) regions. The techniques consist on the amplification of the 16S rDNA
regions followed by the digestion of the amplified fragment with a restriction enzyme. The DNA
fragments obtained are separated by electrophoresis. The resulting patterns are characteristic of
every species and make it possible to characterize almost all the AAB species.
One of the techniques used to identify AAB on strain level is Random amplified
polymorphic DNA-PCR (RAPD-PC). RAPD fingerprint based on the amplification of the
genomic DNA with a single primer of arbitrary sequence, of 9 or 10 bases of length, which
hybridise with sufficient affinity to chromosomal DNA sequences at low annealing temperatures
so that they can be used to initiate the amplification of bacterial genome regions. The
amplification is followed by agarose gel electrophoresis, which yields a band pattern that should
be characteristic of the particular bacterial strain (Caetano-Anolles et al., 1991; Meunier and
Grimont, 1993). The technique has already been used to characterize rice vinegar AAB. They
managed to discriminate among AAB strains and the patterns yielded between 7 and 8 DNA
fragments).
1.8. Ecology of Acetobacter
Ecology is the science of the study of the relationship between living organism and its
environment. AAB can grow in different environment and the components of the environment
affect its growth and activities
1.8.1. Acetobacter in palm wine
The sap of the oil palm tree (Elaeis guinneesis) serves as a rich substrate for various types
of micro-organisms to grow. However, it is as a source for producing palm wine that the
substrate is mainly known for (Amoa-Awua et al., 2006). In various African countries and
beyond, the sap of the palm tree is tapped and allowed to undergo spontaneous fermentation,
which allows the proliferation of yeasts species to convert the sweet substrate into an alcoholic
beverage. Fresh palm wine is sweat, clear, neutral, colourless juice containing minimal sugar
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(less than 0.5%) small amount of protein, gums and minerals (Opara et al., 2013). According to
Oyeku et al. (2009), it consists mainly of water, sugar, vitamins and many aroma and flavour
components in very small amounts.
In various traditional African societies, the palm wine plays a significant role in
customary practices, especially the distilled product from the palm wine, a potent gin called by
various names in West Africa.
Traditionally, it is believed that when drank by nursing mothers; palm wine stimulates
lactations and also has diuretic effect. Palm wine has also been used to enhance men’s potency
due to yeast cell concentration. It could also be used for leavening of dough and used in African
medicine particularly in the treatment of measles and malaria (Onwuka, 2011).
Despite all these good qualities of palm wine, it is a highly perishable sap due to
fermentation which starts soon after the sap is collected and within an hour or two becomes
reasonably high in alcohol (up to 4%). If palm wine is allowed to continue to ferment for more
than 24 hours, it starts to turn into vinegar. This makes it unacceptable to consumers and creates
losses to the food service industries. Fermentation in palm wine is possible because it constitutes
a good growth medium for numerous microorganisms especially for yeast, lactic acid and acetic
acid bacteria (Bechem et al., 2007). According to the research of Amoa-Awua et al. (2006),
concurrent alcoholic, lactic acid and acetic acid fermentation occurs during the tapping of palm
wine from oil palm trees. Yeast growth dominated by S. cerevisiae starts immediately after
tapping begins and alcohol concentrations become substantial in the product after the third day.
Lactic acid bacteria dominated by L. plantarum and L. mesenteriodes are responsible for a rapid
acidification of the product during the first 24 h of tapping whilst the growth of acetic acid
bacteria involving both Acetobacter and Gluconobacter species become pronounced after the
buildup in alcohol concentrations on the third day. Increases in the alcohol level of palm wine
are faster in the container into which the palm wine accumulates during the tapping than in the
receptacle cut out in the tree trunk, and samples which accumulate overnight have alcohol
contents of over 3%. During the holding/marketing of palm wine, the concentration of alcohol
increases from 3% to over 7% in 24 h, remains high for the next 3 days and begins to drop. The
concentration of acetic acid increases slowly from a concentration of about 0.42–0.48% and after
4 days had exceeded the acceptable level of 0.6% (Amoa-Awua et al., 2006). Saccharomyces
cerevisae constitutes about 70% of the total yeast of palm wine and the activities of these
19
microbes are believed to be responsible for conversion of sugar in palm wine to alcohol after a
short time while bacteria induces the conversion of alcohol into vinegar (Onwuka, 2011).
Because of the central role that the alcoholic beverage has played in the traditional society, it is
important that the microbiology and biochemistry of the fermentation process are well
understood.
Authorities who have studied the succession of micro flora in palm wine consistently reported
the emergence of Acetobacter after about 24 hours of fermentation, at which time, alcohol was
present in reasonable quantities (Opara et al., 2013). Earlier researches on the microbiology of
palm wine had isolated Acetobacter from palm wine and these bacteria are believed to be
responsible for souring of palm wine which is not acceptable by many.
Acetic acid bacteria, Acetobacter and Gluconobacter, as well known as vinegar producers are
able to oxidize ethanol to acetic acid by two sequential catalytic reactions of alcohol
dehydrogenase and aldehyde dehydrogenase which are located on the periplasmic side of their
cytoplasmic membrane (Abolhassan et al., 2007). Though these enzymes are important in
industrial production of acetic acid, they are nevertheless spoilage molecules for many types of
food and juices including palm wine (Ameh et al., 2011).
Previous studies on the microbiology of oil palm tree (E. guineensis) and R. hookeri have
incriminated several bacterial and yeast flora to be involved in the fermentation process (Okafor,
1975). Acetobacter species were earlier isolated from oil palm wine (Faparusi, 1973; Okafar,
1975).
1.8.2. Acetobacter in other wines
Alcohol fermentations are carried out by yeast (mainly Saccharomyces cervisiae), which
are responsible for transforming the sugars present in the musts (glucose and fructose) into
ethanol. The second group of microorganisms involved in the wine production is lactic acid
bacteria. These bacteria are responsible for the malolactic fermentation, the process by which the
malic acid is transformed into lactic acid, thus deacidifying and softening the wine. The third
group of wine microorganisms are the acetic acid bacteria. Unlike the other microorganisms
involved in fermentation processes, they have received very little attention, and little is known
about their behavior and dynamics in wine making processes or their contribution to the spoilage
of must and wines. According to Margalith (1981), acetic acid in wine becomes objectionable at
20
concentration exceeding 0.7–1.2g/l. Acetic acid is the main volatile acid in wines and its
presence is frequently described as volatile acidity (Margalith 1981). An excess of acetic acid in
wine is the main problems found nowadays in wineries. Another consequences of high volatile
acidity in wines is the presence of ethyl acetate, which also gives the wines a vinegary taint
(traces of undesirable quality) and makes the wine smell like glue.
The wine making process begins in the vineyard. The grapes acquire and harbor the right sugar
and physiological composition of their juice so that, once they have been crushed, it can be
transformed into wine by yeast. The growth of AAB has been reported during various steps of
the wine-making process, including some conditions in which they would not be expected to
grow.
1.8.3. Acetobacter in Grapes and Musts
As the grapes become mature, the amount of sugars (glucose and fructose) increases.
Those sugars are an optimum growing media for AAB, and in particular for G. oxydans, because
this species clearly prefer ethanol as the carbon source. In these conditions the predominant
species in grapes is usually G. oxydans, and the most common populations are around 102-
105cfu/g (Joyeux et al., 1984a; Du Toit and Lambercht, 2002) (cfu stands for colony forming
unit, a measure of the number of viable cells capable of producing new colonies when seeded,
that are contained in a culture). Because of G. oxydans’ low tolerance of ethanol, it disappears in
the first stages of alcoholic fermentation. Acetobacter and Gluconacetobacter species have also
been isolated from unspoiled grapes, although in very low amounts (Du Toit and Lamberchts,
2002).
Damaged, rotten or Botrytis-infected grapes can be infected by yeasts and acetic acid
bacteria. Yeasts can start metabolizing the sugars in grapes into ethanol, which are then oxidized
into acetic acid by acetic acid bacteria. Damaged grapes contain AAB population, mainly
belonging to Acetobacter species (A. aceti and A. pasteurianus) up to 106cfu/g (Joyeux et al.,
1984b; Grossman and Becker, 1984). These grapes contain high concentrations of acetic acid,
ethanol and glycerol, and small amounts of ethyl acetate (Sponholz and Dietrich, 1985; Drysdale
and Fleet, 1989b). Both ethanol and glycerol are the products of yeast metabolism. The glycerol
produced can be metabolized by AAB into dihydroxyacetone, which affects the sensory quality
21
of the wine and can bind to SO2 thus decreasing its antimicrobial properties. Gluconic acid arises
from the metabolism of glucose by AAB (Drysdale and Fleet, 1988), and can be further oxidized
to produce 5-keto and 2-ketogluconic acid.
Thus, grape juice composition can be significantly altered if the berries are infected with
acetic acid bacteria. The changes not only have an adverse effect on the sensory quality of wine
but also on the growth of yeasts during alcoholic fermentation (Drysdale and Fleet, 1989a) and
the possible growth of lactic acid bacteria (Joyeux et al., 1984b)
Adding SO2 to the musts is common practice in cellars (a wine cellar is a storage room
for wine in bottles or barrels, or more rarely in carboys, amphorae or plastic containers, because
it inhibits the microorganism and hinders the development of undesirable organisms such as
AAB. So the presence and growth of AAB in must will depend on the concentration of SO2
whether it is present in the free or the bonded form. The free form consists of molecular sulphur
dioxide, bisulphate ands sulphite ions. Only the molecular SO2 has anti-microbial effects. The
proportion of molecular SO2 represents from 1% to 10% of the free form depending on the pH of
the wine, therefore, the lower the pH is, the higher proportion of molecular SO2 will exist, and
the higher anti-bacterial effect (Ribereau-Gayon et al., 2000). In this process the must may also
be contaminated by AAB resident in the cellar because of such processes as grapes juice racking
and pumping.
1.8.4. Acetobacter during Fermentation
During alcoholic fermentation both Saccharomyces and non-Saccharomyces yeasts
develop enormously and can reach populations up to 107-108 cfu/ml. During this process, sugars
from must are transformed into ethanol by yeasts, which make this new media more suitable for
Acetobacter and Gluconacetobacter species. In this process, a considerable amount of CO2 is
produced because of the yeast metabolism, and this creates anaerobic conditions that are
theoretically unsuitable for AAB growth. Recent studies by Du Toit et al. (2005), however,
suggest that some AAB strains can survive for a long period under relatively anaerobic
conditions in wine. The pH is usually around 3.5, and the optimum pH for AAB development is
5.5-6.3 (Holt et al., 1994), although some AAB have been isolated at pH 3.0. The pH is also
important for the state in which we can find SO2 in wine. Low concentration of SO2 does not
22
affect the culturability of some AAB strains, and sulphur dioxide does not completely eliminate
the presence of AAB (Du Toit et al., 2005). AAB are able to grow in wines containing 20mg/l of
free SO2 (Joyeux et al., 1984a), which means that the common levels of SO2 in wines are not
enough to inhibit AAB growth. Watanabe and Iino (1984) found that 100mg/l of total SO2 were
needed to inhibit the growth of Acetobacter species in grape must.
The temperature at which alcoholic fermentation takes place depends on the type of
vinification. Red wine fermentations take place between 25 and 38oC, which is the same as the
optimum temperature for AAB growth (Holt et al., 1994), and therefore does not seem to prevent
AAB development. The temperature of white and rose fermentations ranges from 18-19oC and
the effect of low-temperature fermentations on the AAB population has not been studied yet.
Growth of these bacteria during alcoholilc fermentation may also be linked to the number of
bacteria and yeast in the must at the start of the fermentation (Watanabe and Iino, 1984). The
predominant species during alcoholic fermentation are commonly A. aceti, A. pasteurianus, Ga.
Liquefaciens and Ga. Hansenii (Joyeux et al., 1984b; Du Toit and Lamberchts, 2002), although
G. oxydans have also been isolated as the only species during the fermentation.
In spite of these adverse conditions during alcoholic fermentation, some authors (Du Toit
et al., 2005) have detected that AAB can survive and even grow during this process. If the
quality of the wines is to be good, it is of vital importance to keep the numbers of AAB low. This
can be done by using healthy grapes, inoculating a high quality of yeast, adding SO2, clarifying
the must and lowering the pH by adding acid (Du Toit and Pretorius, 2002).
If AAB grow a lot in the first stages of alcoholic fermentation, fermentation may become stuck
or sluggish and there may be renewed growth of AAB and the reduction in the quality of the
wines during their storage (Joyeux et al., 1984b)
1.8.5. Acetobacter during ageing and wine maturation
During storage, the major species found belong to Acetobacter (A. aceti and A.
pasteurianus). These bacteria have been isolated from the top, middle and bottom of the tanks
and barrels, suggesting that AAB can actually survive under the semi-anaerobic conditions
occurring in wine containers. This can be explained by the ability of AAB to use compounds,
23
such as quinines and educible dyes, as electron acceptors (Du Toit and Pretorius, 2002). The
main product obtained from the presence of AAB at this point is acetic acid, although
considerable amounts of acetaldehyde and ethyl acetate are produced (Drysdale and Fleet,
1989b) and glycerol metabolises to dihydroxyacetone. The pumping over and racking of wine
may stimulate the growth of AAB and lead to populations up to 108cel/ml (Joyeux et al., 1984b;
Drysdale and Fleet, 1989b), because of the intake of oxygen during these operations. The
number of bacteria usually decreases drastically after bottling, because of the relatively
anaerobic conditions in a bottle. However, the excessive addition of oxygen during bottling can
increase the number of AAB. The ethanol concentration of wine is around 10-15% (v/v). As
mentioned above, ethanol is a good carbon source for AAB, but it can also inhibit AAB growth
at high concentrations. However, it is well known that these bacteria can grow in wine
containing between 10-14% (v/v) (Joyeux et al., 1984a; Drysdale and Fleet, 1989a; Koselbalan
and Ozlingen, 1992; Du Toit and Pretorius, 2002). It has been reported by Saeki et al. (1997) that
AAB can overcome the inhibitory effect, and become tolerant to ethanol. In this respect AAB
have been isolated from sake and tequila (beverages with a much higher ethanol concentration
than wine) (Joyeux et al., 1984a), although Drysdale and Fleet (1989b) observed a weak growth
of AAB even at 10oC.
1.8.6. Acetobacter in vinegar production
Vinegar is a precious food additive and complement as well as effective preservative against
food spoilage that is produced by Acetic acid bacteria and contains essential nutrients such as
amino acids regarding its fruit source (Kocher et al., 2006). Food and Drug
Administration(FDA), USA has explained the vinegar as a 4% acetic acid solution that is
synthesized from sweet or sugary substances through alcoholic fermentation. The neoclassical
fermentation resulted in several vinegar types with different tastes, frangrances and nutritional
values because of applying various acetic acid bacteria in vinegar making procedure. Currently
the vinegar manufacturers are seeking for new types of vinegar using different AAB as their
starter and tradional vinegar production has been improved using various natural substrates and
fruits (Du Toit and Lambrechts, 2002). Acetobacter strains are the major bacteria that are dealing
with vinegar production industrially (Sokollek et al., 1998; Kaeere et al., 2008).
24
Vinegar has been very important in the human diet since ancient times as a condiment
and food preservative; for many centuries, acetic acid from vinegar was the strongest acid, until
sulphuric acid was discovered around the year 1300. Although little is known about the role
played by microorganisms in vinegar production, vinegar has been produced mainly from wine,
alcohol and rice. Nowadays knowledge is much more advanced, above all as far as the analytical
and industrial processes are concerned, but the microbiology of the process is still not well
understood. At the beginning of the 21st century, the species and strains responsible for vinegar
production are still not very clear. Nowadays, there are three different biotechnological processes
for producing vinegar (Greenshields, 1978): the Orleans method (this is the most famous slow
method of vinegar production. Here, barrels are filled with wine and vinegar and fermentation
are carried out slowly by AAB, which will generally metabolize all the alcohol in 1 to 3 months),
the German method (a very quick method also called generator method. In this method, the
alcoholic solution to be acetified is allowed to trickle down through a tall tank or column
(generator) packed with porous solid material on whose surface Acetobacter bacteria are
permitted to grow) and the submerged method (a catalysed fast method involving acetator, a tank
equipped with a variety of system that keep the mixture constantly turning, introducing air into
the mixture to introduce oxygen to keep the bacteria working).
1.9. Alcohol dehydrogenase
Alcohol dehydrogenase (EC.1.1.5.5) (Gomez-Manzo et al., 2008) otherwise called
pyrolloquinoline quinone alcohol dehydrogenases or Alcohol dehydrogenase or type III Alcohol
dehydrogenase or membrane associated quinohaemoprotein alcohol dehydrogenase, an enzyme
with system name alcohol:quinone oxidoreductase, belongs to quinoenzymes and requires
quinoid cofactors (e.g., pyrroloquinoline quinone, PQQ) as enzyme-bound electron acceptors.
They are distinct from other types of alcohol dehydrogenases because of their position in the
cells. While majority of other alcohol dehydrogenases are located in the cytosol, otherwise called
cytosolic NAD+/NAD(P)+-dependent alcohol dehydrogenase located in the cytoplasm,
these family of alcohol dehydrogenase are membrane-bound. Many membrane-bound
dehydrogenases in the periplasmic space or on the outer surface of the cytoplasmic
membrane of acetic acid bacteria and other aerobic Gram-negative bacteria have been
classified as PQQ- or FAD-dependent dehydrogenases (Matsushita et al., 1994). Most of the
25
enzymes are closely associated with oxidative fermentation in industry, catalyzing an
incomplete one-step oxidation, allowing accumulation of an equivalent amount of
corresponding oxidation products outside the cells. The active sites of individual enzymes
face the periplasmic space (Fig 1). Apart from alcohol dehydrogenases, there are other
membrane-bound dehydrogenases such as glucose dehydrogenase and fructose
dehydrogenase. All the enzyme reactions are carried out by periplasmic oxidase systems
including alcohol- and sugar-oxidizing enzymes of the organisms. D-Glucose, ethanol,
and many other substrates are oxidized by the dehydrogenases (shown as PQQ or FAD,
except for aldehyde dehydrogenase) that are tightly bound to the outer surface of the
cytoplasmic membranes of the organism. These membrane-bound enzymes irreversibly
catalyze incomplete one-step oxidation and the corresponding oxidation products
accumulate rapidly in the culture medium or reaction mixture. The electrons (e-)
generated by the action of these dehydrogenases are transferred to ubiquinone in the
membrane. The reducing equivalents are further transferred to the terminal ubiquinol
oxidase in the cytoplasmic membranes. The terminal oxidase generates an
electrochemical proton gradient either by charge separation or by a proton pump or by
both during substrate oxidation by the membrane-bound enzymes, allowing the
organism to acquire bioenergy through substrate oxidation. Thus, the organisms generate
bioenergy through the enzyme activities of PQQ- and FAD-dependent dehydrogenases.
Many different NAD- and FAD- dependent dehydrogenases in the cytoplasm have no
function in oxidative fermentation and thus are not shown in Fig. 2.
26
Fig.2. Membrane-bound PQQ- and FAD-dependent primary dehydrogenases on the outer
surface of acetic acid bacteria. (Adachi et al., 2007).
1.9.1. Classes of alcohol dehydrogenases (EC.1.1.5.5)
There are different classes of alcohol dehydrogenase or Pyroloquinoline quinone (PQQ)-
dependent alcohol dehydrogenases. Among the most comprehensively studied of these enzymes
are the three classes of PQQ-containing quinoprotein alcohol dehydrogenases; Type I are
soluble, periplasmic enzymes containing a single Pyroloquinoline Quinone prosthetic group; this
group includes the methanol dehydrogenase of methylotrophs. Type II dehydrogenases are
soluble, periplasmic quinohemoproteins, having a C-terminal extension containing heme C. Type
III dehydrogenases have similar quinohemoprotein subunits but have two additional subunits
(one of which is a multiheme cytochrome c), bound in an unusual way to the periplasmic
membrane (Anthony, 2004). These membrane enzymes and other quionoprotein dehydrogenases,
their prothetic group, electron acceptors, location and organisms in which they are found are
summarized in the Table 3 below.
27
Table 3. Summary of quinoprotein and quinohemoprotein dehydrogenase
Enzyme Location Prosthetic
group
Electron
acceptor
Organism
Type 1 alcohol
dehydrogenase: eg
-methanol
dehydrogenase
-ethanol dehydrogenase
periplasm
periplasm
PQQ
PQQ
Cytochrome c
Cytochrome c
Methylotroph
Pseudomononas sp
Type 11 alcohol
dehydrogenases
Membrane PQQ
Heme c
Azurin Comamonas
testosterone
Pseudomonas putida
Type 111 alcohol
dehydrogenases
Membrane PQQ
4 heme c
UQ Acetic acid bacteria
Sorbitol dehydrogenase Membrane PQQ
4 heme c.
UQ Acetic acid bacteria
Membrane glucose
dehydrogalnse (m-GDH)
Membrane PQQ UQ Enteric bacteria
Acetic acid bacteria
Acinetobacter
calcoaceticus
Soluble glucose
Dehydrogenase (s-GDH)
Periplasm PQQ ? Acenetobacter
calcoaceticus
Glycerol dehydrogenase Membrane PQQ UQ Acetic acid bacteria
D-arabitol
dehydrogenase
Membrane PQQ UQ Acetic acid bacteria
D-sorbitol
dehydrogenase
Membrane PQQ UQ Acetic acid bacteria
Lupanine hydroxylase Periplasm PQQ heme c Cytochrome c Pseudomonas sp
Sorbose/sorbosone
dehydrogenase
Periplasm PQQ Cytochrome c Acetic acid bacteria
Methylamine Periplasm TTQ Amicyanin Methylotrophs
28
dehydrogenase
Aromatic amine
dehydrogenase
Periplasm TTQ Azurin Alcaligenes
Amine dehydrogenase Periplasm CTQ 2 heme Azurin Pseudomonas putida
Paracoccus
denitrificans
UQ = ubiquinone, PQQ = pyroloquinoline quinone, TTQ = tryptophan tryptophyl quinone, CTQ
= Cysteine Tryptophylquinone
Source: (Matsushita et al., 2002; Davidson, 1993; Anthony, 1996; Goodwin and Anthony, 1996;
Davidson, 2000; Choi et al., 1995; Hyun and Davidson, 1995; ; Anthony, 2000; Adachi et al.,
1998; Hopper and Rogozinski, 1998; Asakura and Hoshino, 1999; Cozier et al., 1999; Takagi et
al., 1999; Yoshida et al., 1999; Afolabi et al., 2001; Elias et al., 2000, 2001; Keitel et al., 2000;
Adachi et al., 2001; Datta et al., 2001; Sugisawa and Hoshino, 2001; Chen et al., 2002; Miyazaki
et al., 2000; Oubrie et al., 2002; Satoh et al., 2002)
1.9.1.1. The Type I Alcohol Dehydrogenase
Methanol dehydrogenase (MDH) belongs to type 1 alcohol dehydrogenase. The MDH of
methylotrophic bacteria oxidizes methanol to formaldehyde during growth of bacteria on
methane or methanol (Anthony, 1982), during which its electron acceptor is a novel acidic
cytochrome (cytochrome cL) (Anthony, 1992). MDH is also responsible for oxidation of ethanol
to acetaldehyde during growth on ethanol. Using phenazine ethosulphate in a dye-linked assay
system the pH optimum is about 9 and ammonia or methylamine is required as activator. MDH
oxidizes a wide range of primary alcohols (very rarely secondary alcohols), having a high
affinity for these substrates; for example, the Km for methanol is 5–20 M. The pH optimum for
the reaction with cytochrome cL is 7.0, and ammonia is not usually required as activator.
The X-ray structure has been determined for the MDH from Methylobacterium
extorquens (Blake et al., 1994; Ghosh and Anthony, 1995; Afolabi et al., 2001), and from
Methylophilus sp. (Xia et al., 1992; White et al., 1993; Xia et al., 1996; Xia et al., 1999; Zheng
et al., 2001). MDH has an α2β2 tetrameric structure; each α subunit (66 kDa) contains one
molecule of PQQ and one Ca2+ ion. The β subunit is very small (8.5 kDa), it cannot be reversibly
29
dissociated, its function is unknown and it is not present in any other quinoproteins. The large α
subunit has a propeller fold making up a superbarrel (Fig. 3)
Fig.3. Propeller structure of type 1 alcohol dehydrogenase (methanol dehydrogenase)
Source: Gosh et al., 1995
The αβ unit of MDH looking down the pseudo 8-fold axis, simplified to show only the β-strands
of the ‘W’ motifs of the α-chain, and the long α-helix of the β-chain, but excluding other limited
β-structures and short α-helices (Ghosh et al., 1995). The PQQ prosthetic group is in skeletal
form and the calcium ion is shown as a small sphere. The outer strand of each ‘W’ motif is the D
strand, the inner strand being the A strand. The ‘W’ motifs are arranged in this view in an anti-
clockwise manner. The exceptional motif W8 is made up of strands A-C near the C-terminus,
plus its D strand from near the N-terminus.
The structure has several important novel features, including novel the ‘tryptophan-docking
motifs’ that link together the eight beta sheets, and the presence in the active site of an unusual
eight-membered disulphide ring structure formed from adjacent cysteine residues, joined by an
30
atypical non-planar peptide bond. The PQQ is sandwiched between the indole ring of Trp243
and the disulphide ring structure (Fig. 4).
Fig.4 The novel disulphide ring in the active site of type 1 alcohol dehydrogenase (methanol
dehydrogenase)
Source: Ghosh et al. 1995.
The ring is formed by disulphide bond formation between adjacent cysteine residues. The
PQQ is ‘sandwiched’ between this ring and the tryptophan that forms the floor of the active site
chamber. The calcium ion is coordinated between the C-9 carboxylate, the N-6 of the PQQ ring
and the carbonyl oxygen at C-5. This structure is seen in all the alcohol dehydrogenases but not
in aldose dehydrogenases. The indole ring is within 15o of co-planarity with the PQQ ring and,
on the opposite side, the two sulfur atoms of the disulphide bridge are within 3.75Å of the plane
of PQQ. The rarity of the disulphide ring structure would suggest some special biological
function. Reduction of the disulphide bond leads to loss of activity but oxidation in air or
carboxymethylation of the free thiols leads to return of activity. The activity of the
carboxymethylated derivative rules out reduction to the thiols during the catalytic cycle. The
disulphide ring is not present in the quinoprotein glucose dehydrogenase in which electrons are
transferred to membrane ubiquinone from the quinol PQQH2, and in which the semiquinone free
radical is unlikely to be involved as a stable intermediate. It is possible, therefore, that this novel
structure might function in the stabilization of the free radical PQQ semiquinone or its protection
from solvent at the entrance to the active site in MDH (Blake et al., 1994; Avezoux et al., 1995).
31
Recent work with the quinohemoprotein (Type II) alcohol dehydrogenase suggests, however,
that, although it does not become completely reduced, the disulphide ring is essential for intra-
protein electron transfer in all the alcohol dehydrogenases (Oubrie et al., 2002). In addition to the
axial interactions, many amino acid residues are involved in equatorial interactions with the
substituent groups of the PQQ ring system (Fig. 5).
.
This Figure also shows Asp303, which is likely to act as the catalytic base, and Arg331
which may also be involved in the mechanism. The equatorial interactions of the
quinohemoprotein alcohol dehydrogenase (QH-ADH) are almost identical to these, an important
exception being that Arg331 is replaced by a lysine (Chen et al., 2002; Oubrie et al., 2002), as is
also the case in glucose dehydrogenase. These are exclusively hydrogen-bond and ion-pair
interactions. Although the number of polar groups involved might indicate at first sight that the
environment of the PQQ is polar, this is not the case. Oxygen of the 9-carboxyl forms a salt
bridge with Arg109 and both groups are shielded from bulk solvent by the disulphide. The
carboxyl group of Glu155 and a 2-carboxyl oxygen of PQQ are also shielded from solvent and it
is probable that at least one is protonated, their interaction thus being stabilized through
Fig. 5. The equatorial interactions of PQQ and the coordination of Ca2+ in the active site of type 1
alcohol dehydrogenase (methanol dehydrogenase)
Source: Ghosh et al., 1995
32
hydrogen bond formation. The active site contains a single Ca++ ion whose coordination sphere
contains PQQ and protein atoms, including both oxygens of the carboxylate of Glu177 and the
amide oxygen of Asn261. The PQQ atoms include the C5 quinone oxygen, one oxygen of the C7
carboxylate and, surprisingly, the N6 ring atom which is only 2.45Å from the metal ion (Fig. 5).
The C4 and C5 oxygen atoms, which become reduced during the catalytic cycle, are hydrogen
bonded to Arg331, which also makes hydrogen bonds between its NH2 and the carboxylate of
Asp303 which is the most likely candidate for the base required by the catalytic mechanism.
Ethanol Dehydrogenase of Pseudomonas species (QEDH) is also a type 1 alcohol
dehydrogenase. This ethanol dehydrogenase (QEDH), induced during growth on ethanol of
Pseudomonas or Rhodopseudomonas, is similar to MDH (Mutzel and Gorisch, 1991; Toyama et
al., 1995; Keitel et al., 2000). It uses a specific cytochrome c550 as electron acceptor (Schobert
and Gorisch, 1999), although this shows no sequence identity to cytochrome cL, the electron
acceptor for MDH. Like MDH, QEDH has a high pH optimum, requires ammonia or
alkylamines as activator in the dye-linked assay system (ferricyanide is not used as electron
acceptor), and is able to oxidize a wide range of alcohol substrates including secondary alcohols,
but it differs in its very low affinity for methanol; the Km for ethanol is about 15 M and that for
methanol is about 1000 times higher. QEDH is homodimeric, the subunits being 65 kDa; it thus
differs from MDH in lacking a small subunit.
Unlike MDH, PQQ dissociates from QEDH after removal of Ca2+ with EDTA, this
process being reversible after reconstitution in the presence of Ca2+ and PQQ (Mutzel and
Goerisch, 1991). It is possible that the additional disulphide bridge in the subunit of MDH and
the complex with the small subunit may lead to a stronger stabilization of the native
conformation of the enzyme.
The X-ray structure of the enzyme from Pseudomonas aeruginosa shows that, apart from
differences in some loops, the folding pattern is very similar to the large (α) subunit of MDH
(Keitel et al., 2000).
There are different loops in the vicinity of the active site and several rather flexible loops
protrude from the molecule surface and partly occupy the space filled by the small subunit of
MDH. The PQQ is located in the center of the superbarrel, coordinated to a calcium ion. Most
amino acid residues that make contact with the PQQ and the Ca2+ are similar to those in MDH.
33
The main differences in the active site region are a bulky tryptophan residue in the active-site
cavity of MDH, which is replaced by a phenylalanine and a leucine side-chain in the QEDH, and
a leucine residue right above the PQQ in MDH which is replaced by a tryptophan side-chain in
QEDH. Both amino acid exchanges appear to have an important influence, causing the different
substrate specificities of these otherwise very similar enzymes. Docking calculations suggest that
one of the tryptophans must be able to change its orientation in order to accommodate the higher
primary alcohols in the active site (Keitel et al., 2000). In addition to the Ca2+ ion in the active-
site cavity, QEDH contains a second Ca2+-binding site at the N terminus, which contributes to its
stability. Although the localization of the interaction surfaces between the subunits is identical in
QEDH and MDH, the residues and the interactions involved are not conserved.
1.9.1.2. The Type II alcohol dehydrogenase
Soluble Quinohemoprotein Alcohol Dehydrogenase (QH-ADH) of Comamonas
testosteroni is a type II alcohol dehydrogenase. The best-known quinohemoprotein ADH is that
isolated from Comamonas testosteroni (Groen et al, 1986; Jongejan et al., 1998; Oubrie et al.,
2002). It has also been described in Pseudomonas putida (Toyama et al., 1995; Chen et al.,
2002) which produces two distinct forms, having different substrate specificities; ADH-IIB is
induced during growth on butanol and ADH-IIG induced on glycerol. This same organism also
produces a Type I alcohol dehydrogenase, induced during growth on ethanol. The electron
acceptor for QH-ADH is a specific blue copper protein, azurin (Matsushita et al., 1999) which is
probably oxidized directly by the membrane oxidase. This periplasmic enzyme is a monomer (71
kDa) containing one molecule of PQQ and a single heme C. In the dye-linked assay system the
pH optimum is 7.7 and there is no requirement for an amine activator. Because electron transfer
from PQQ is by way of heme C this enzyme can also be assayed using ferricyanide. It has a wide
specificity for primary and secondary alcohols, although it is unable to oxidize methanol; it also
oxidizes aldehydes and can accept large molecules such as steroids as substrates. This has been
exploited for enantiospecific oxidation of industrially important precursor molecules (synthons)
(Geerlof et al., 1994). The enzyme has been extensively characterized by EPR, NMR and
Raman resonance spectroscopy with respect to the nature of the heme and its relationship to
PQQ (De Jong et al., 1995a; De Jong et al., 1995b), the conclusions being supported by the X-
ray structures of the enzymes from Comamonas (Oubrie et al., 2002) and from Pseudomonas
34
(Chen et al., 2002). QH-ADH comprises two domains connected by a long linker (23 amino
acids) which spans the whole length of the enzyme.
The N-terminal dehydrogenase domain has the typical β barrel with its propeller fold,
having the active site containing PQQ and a Ca2+ ion. The C-terminal domain, located on top of
the dehydrogenase, is a type I cytochrome c, with 5 α helical segments, which enclose the C-type
heme which is covalently bonded to cysteine residues and which has typical histidine and
methionine heme iron ligands. A channel leads from the periplasm to the region of the PQQ and
a second channel contains a chain of hydrogen-bonded water molecules between the periplasm
and the cavity between the two domains. The N-terminal dehydrogenase domain is very similar
to the α subunit of MDH, the PQQ being located at the top of the superbarrel in a hydrophobic
cavity that is accessible through a deep and narrow channel. It is ‘sandwiched’ between a co-
planar tryptophan and the disulphide ring as in MDH (Fig. 4) and it has in-plane bonding
interactions with almost exactly the same side chains as in MDH, the only significant difference
is that Arg331 which is bonded to the O5 of PQQ in MDH (Fig. 5) is replaced by a lysine side
chain in QH-ADH as it is in mGDH. The ligation of the Ca2+ ion with PQQ and with amino acid
side chains is also exactly the same as in MDH (Fig. 5). QH-ADH is the only alcohol
dehydrogenase whose X-ray structure includes the substrate, or rather a product of substrate
oxidation. In the case of the Comamonas enzyme this is tetrahydrofuran-2- carboxylic acid,
presumably produced from the two step oxidation of tetrahydrofurfuryl alcohol (Oubrie et al.,
2002). The tetrahydrofuran ring makes van der Waal’s contacts with the hydrophobic walls of
the substrate cavity. An oxygen atom of the substrate carboxylate is hydrogen bonded to the
active site aspartate (Asp303 in MDH), and the glutamate carboxylate that coordinates to the
Ca2+ (Glu177 in MDH), and to the two sulfur atoms of the disulphide ring. The enzyme from
Pseudomonas putida was crystallized in the presence of isopropanol, and acetone, its oxidation
product, was shown to be present in the active site, close to the O-4 and O-5 of PQQ and close to
a carboxylate oxygen of the proposed active site aspartate (Asp303 in MDH; Fig. 5) (Chen et al.,
2002). The side chains of the products lie in a cavity lined with mainly hydrophobic side chains-
cysteines, phenylalanines, tyrosine and proline. The volume of this substrate cavity is about 120
Å3 which is about twice that of the Type I ethanol dehydrogenase and much larger than seen in
the Xray structure of MDH (~18 Å3) (Chen et al., 2002). MDH is unable to oxidize secondary
alcohols or primary alcohols with substituents on the C2 atom but it is still able to oxidized a
35
wide range of large alcohols and it is possible that the entrance to the active site might be flexible
in order to accommodate these substrates. Because the catalytic machinery of MDH is strictly
conserved in QH-ADH, the mechanism of alcohol oxidation is most likely identical for the two
enzymes (Oubrie et al., 2002). The positions of the substrate products together with the results
obtained by site directed mutagenesis of Asp303 in MDH (Afolabi et al., 2001), all suggest that
Asp308 is the catalytic base. The mechanism for aldehyde oxidation is presumably essentially
similar to that for alcohol oxidation: it is proposed that Asp308 abstracts a proton from a
hydrogen-bonded water and the resulting hydroxyl ion performs a nucleophilic attack on the
aldehyde C1 atom in concert with hydride transfer from this atom to the C5 of PQQ, to give the
carboxylic acid product (Oubrie et al., 2002). The shortest distance between PQQ and the heme
is 13–15 Å which is close to the maximum travel distance for electrons but the predicted rate of
transfer through the protein is much higher than the measured rate of substrate oxidation. A
number of paths are possible for the electron flow but they all involve the disulphide bridge and
probably at least one water molecule (for example, see Fig. 8) (Chen et al., 2002; Oubrie et al.,
2002). During oxidation of the reduced PQQ, protons are released into the periplasm. This is
likely to be by way of a hydrogen bonded network involving a water filled chamber between the
two domains, Lys335, Asp308 and Glu185 (Oubrie et al., 2002); these are equivalent to the
MDH residues Arg331, Asp303 and Glu177 (Fig. 5). Azurin isolated from P. putida is a good
electron acceptor for the QH-ADH, the interaction being mediated by hydrophobic forces
(Matsushita et al., 1999). The heme is buried within the cytochrome domain except for one edge
which is surrounded by a charge-neutral surface area which may form a binding site for azurin,
in which one of the histidine ligands to the buried copper is exposed to the surface and is
surrounded by a surface patch of hydrophobic residues (Chen et al., 2002).
1.9.1.3. Type III alcohol dehydrogenase
Membrane Associated Quinohemoprotein Alcohol Dehydrogenase of acetic acid bacteria
is a type III alcohol dehydrogenase. This enzyme is a quinohemoprotein-cytochrome c complex
and has only been described in the acetic acid bacteria, Acetobacter and Gluconobacter
(Matsushita and Adachi, 1993; Matsushita et al., 1994, Goodwin and Anthony, 1998). Together
with the membrane-bound aldehyde dehydrogenase, it is responsible for the oxidation of alcohol
to acetic acid in vinegar production. It does not require ammonia as activator and has a pH
36
optimum of 4-6. Its substrate specificity is relatively restricted, oxidizing only a few primary
alcohols (chain length, C2-C6) (but not methanol), or secondary alcohols and has some activity
with formaldehyde and acetaldehyde.
The Type III ADH has 3 subunits and is tightly bound to the periplasmic membrane,
requiring detergent for its isolation. Translation of the gene sequences shows that all the subunits
have N-terminal signal peptides typical of periplasmic proteins. Its natural electron acceptor is
ubiquinone in the membrane. Subunit I (72-80 kDa) is a quinohemoprotein similar to the soluble
Type II Quinoheamoprotein alcohol dehydrogenase, with a single molecule of PQQ and a single
heme C. Its N-terminal region has sequence similarity to the soluble methanol dehydrogenase but
with a C terminal extension having a single heme binding site. Subunit II (48k-53 kDa) has 3
hemes that can be distinguished by biochemical techniques in the pure protein (Matsushita et al.,
1996). Subunits I and II therefore have a total of 4 hemes. Most Type III ADHs have a third
subunit (subunit III, 14-17 kDa) in which the gene is not linked to the genes encoding the other
two subunits, and whose predicted amino acid sequence indicates that its processed size is
greater (about 20 kDa) than that obtained by SDS -PAGE (14 kDa). The Type III ADH may be
assayed with phenazine methosulfate, or with ferricyanide which reacts at the level of one or
more of the heme C prosthetic groups on subunits I and II. It differs from all other ADHs in
using short-chain ubiquinone homologues (Q1 and Q2) as electron acceptors and native
ubiquinone (Q9 and Q10) when reconstituted in membrane vesicles (Matsushita et al., 1992).
There is good evidence that electron transfer from reduced PQQ to the membrane ubiquinone
takes place by way of the hemes on the cytochrome subunit II but that only two of them may be
involved in this electron transfer process (Matsushita et al., 1996; Frebortova et al., 1998). It has
been suggested that the cytochrome subunit II is firmly embedded in the membrane, that subunits
I and III are firmly attached to each other and that this attachment helps the dehydrogenase
subunit I couple with the cytochrome c (subunit II). This raises the question of how the
ubiquinone in the membrane reacts with subunit II to accept electrons from its heme. Clearly part
of the protein must be embedded in the membrane for this to occur but subunit II does not appear
to have typical hydrophobic transmembrane helices (Kondo and Horinouchi, 1997). The Type III
ADH thus appears to be unique in a number of ways; it requires detergent for its isolation from
membranes and so seems to be a typical integral membrane protein, although none of the
subunits appears (from their gene sequences) to have characteristic membrane protein structural
37
domains. Furthermore, the electron acceptor for the quinohemoprotein/cytochrome c complex is
membrane ubiquinone, so we have the unusual situation where a c-type cytochrome precedes
ubiquinone in the electron transport chain (Anthony, 2004).
1.10. Pyroloquinoline Quinone (PQQ) Cofactor
4, 5-dihydro-4, 5-dioxo-1H-pyrrolo- [2, 3- ] quinoline-2, 7, 9-tricarboxylic acid (PQQ)
(Fig. 6) is an aromatic, tricyclic ortho-quinone that serves as the redox cofactor for several
bacterial dehydrogenases. Among the best-known examples are methanol dehydrogenase and
glucose dehydrogenase. PQQ belongs to the family of quinone cofactors that has been
recognized as the third class of redox cofactors following pyridine nucleotide- and flavin-
dependent cofactors.
PQQ is a prosthetic group required by several bacterial dehydrogenases, including methanol
dehydrogenase (MDH) of Gram negative methylotrophs, quinohemoprotein alcohol
dehydrogenase and some glucose dehydrogenases. PQQ is derived from two amino acids,
tyrosine and glutamic acid (Houck, 1991; Van Kleef, 1988) (i.e all carbon and nitrogen atoms of
PQQ are derived from conserved tyrosine and glutamate residues), but the pathway for its
biosynthesis is unknown.
PQQ is an important cofactor of bacterial dehydrogenases, linking the oxidation of many
different compounds to the respiratory chain. PQQ was the first of the class of quinone cofactors
that have been discovered in the last 18 years and make up the prosthetic group of quinoproteins
(Duine, 1991)
PQQ was discovered in 1979 from a bacterium, and afterward it was reported to be in common
foods. Because PQQ-deprived mice showed several abnormalities, such as poor development
and breakable skin, PQQ has been considered as a candidate for vitamin. It was a mystery, that
until 2003 it was not identified as vitamin. Since the first vitamin (now called vitamin B1) was
discovered in 1910 by Dr. U. Suzuki, thirteen substances have been recognized as vitamins; the
latest one was vitamin B12 found in 1948.So it takes 55 years to discover “PQQ” a previously
identified substance as new vitamin ( Choi, 2008; Kashara and Kato, 2003).
38
1.11. Mechanisms of oxidation of alcohols in alcohol dehydrogenases,
Two mechanisms have been proposed for the oxidation of alcohols in quinoprotein
dehydrogenases, both of which begin with the pyroloquinoline quinone in an oxidized state.
Initially, an addition/elimination mechanism was proposed, a suggestion that is now
considered unlikely; rather, a hydride transfer mechanism is preferred (Oubrie et al., 1999;
Oubrie and Dijkstra., 2000; Anthony and Williams, 2003) ( Fig. 7)
Fig 6: Chemical structure of PQQ (4, 5-dihydro-4, 5-dioxo-1H-pyrrolo- [2, 3-f] quinoline-
2, 7, 9-tricarboxylic acid)
(Source:http://www.dlarborist.com/treetrends/2005/05/27/auxin_action_s.jpg)
39
Fig. 7. The current accepted reaction cycle for alcohol oxidation in quinoproteins
Source: Anthony and Williams (2003)
The mechanism is based on a hydride transfer from the alcohol to the C-5 position of the
pyroloquinoline quinone (Oubrie et al., 1999). The back-reaction is via a radical intermediate
protonated at O-4 or O-5. The IUPAC numbering scheme of PQQ is also shown. Following
substrate binding, the reaction is initiated by amino acid (Asp(11) or Glu (25)) base-catalyzed
proton abstraction of the hydroxyl proton of the alcohol. Nucleophilic attack of the hydride from
the substrate to the C-5 position of PQQ then occurs. Subsequently, the PQQ enolizes to form the
quinol. The reduced PQQ is reoxidized by two sequential single electron transfers (ET) to
cytochrome c1 in MDH, cytochrome c550 in QEDH, or the cytochrome c domain in QH-ADH
via the intermediate free radical (Duine and Frank, 1980; Frank, et al., 1988; Dijkstra et al.,
40
(1989), a process that is thought to be mediated by the disulfide bridge (Avezoux et al., 1995, Oubrie et
al., 2002; Chen et al., 2002).
Information that is not usually obtained from x-ray analysis but is necessary for
obtaining full understanding of a dehydrogenation reaction cycle concerns the protonation states
of the single and doubly reduced species. As shown in Fig. 8, apart from one review (Duine,
1999), reduced PQQ is usually shown protonated at both O-4 and O-5, whereas the radical is
depicted singly protonated at either O-4 (Zheng and Bruice, 1997; Duine et al., 1984) or O-5
(Anthony,1996; Anthony and Williams, 2003; Oubrie, 2003), although a deprotonated radical
was recently postulated (Sato et al., 2001). Knowledge of the protonation states is crucial if the
electron transport (ET) and proton transfer pathway in both reoxidation steps are to be
understood, because depending on the protonation states of the initial and final molecules, the
reaction is either simple ET or must be accompanied by the release of a proton. Furthermore,
apart from the driving force and the reorganization energy, according to ET theory (Marcus and
Sutin.,1985), the rate of ET is dependent on the electroninc coupling between the donor (PQQ)
and the acceptor (heme). Therefore, a full understanding of ET kinetics in quinoproteins will
only be possible with knowledge of both the spatial and electronic structures of the ET partners
(Davidson, 2004). The later may be provided by electron nuclear double resonance (ENDOR)
via determination of hyperfine coupling (hfcs) in combination with density functional theory
(DFT) calculations (Buttner et al., 2005). These methods enable us to establish that the PQQ
radical is deprotonated when bound in QEDH from Pseudomonas aeruginosa
The other enzyme involved in the oxidation of ethanol is aldehyde dehydrogenase. It is
also a NADP+ independent enzyme and located in the cytoplasmatic membrane. Its optimum pH
is between 4 and 5, although it can catalyse the oxidation of acetaldehyde to acetate at lower pH
values (Adachi et al., 1980). It is an enzyme that is sensitive to oxygen concentrations, and when
these are low its activity decreases, accumulating acetaldehyde. It is also more sensitive to the
presence of ethanol than alcohol dehydrogenase (Muraoka et al., 1983).
1.12. In vitro and in vivo properties of Alcohol dehydrogenase
The particulated alcohol dehydrogenase could be assayed in vitro in the presence of one
of the following dyes as an electron acceptor; 2,6-dichlorophenolindophenol, phenazine
methosulfate or potassium ferricyanide. NAD or NADP were not effective as an electron
41
acceptor at all (Adachi et al., 1978). Many people still believe that acetate is produced by the
cytosolic NAD(P)-dependent alcohol dehydrogenase and keto-D-gluconate by the cytosolic
NAD(P)-dependent D-gluconate dehydrogenase located in the cytoplasm. Such a serious
confusion is probably caused by the confused of localization of the enzymes concerned. Before
describing the the actions of the individual PQQ- and FAD-dependent dehydrogenases, it is
worth clarifying the common physiological roles and localizations of PQQ-and FAD-dependent
dehydrogenases in acetic acid bacteria and other microorganisms. At present, of the enzymes
exploited as either PQQ-dependent or FAD-dependent dehydrogenases, aldehyde dehydrogenase
is the only one that is known to use a molybdopterin coenzyme. Unlike the cytoplasmic
oxidoreductases, no energy is required for substrate intake into the periplasm and pumping out
the oxidation products across the outer membrane. Microbial production of L-sorbose, aldehyde
(which is ultimately converted to acetic acid) and keto-D-gluconate are the examples shown in
the Fig 8 below.
Fig 8. Membrane- bound dehydrogenase-dependent periplasmic oxidase systems (Adachi et al.,
2007).
42
All substrate are oxidized by the respective membrane-bound dehydrogenase, of which
active site faces the periplsmic space formed between the outer membrane and the cytoplasmic
membrane. The dehydrogenase then donates electrons to ubiquinone than in turn transfers them
to the terminal ubiquinol oxidase. The terminal oxidase generates an electrochemical proton
gradient either by charge separation or by a proton pump or by both during substrate oxidation
by the membrane-bound enzyme, allowing the organism to acquire bioenergy through substrate
oxidation (Adachi et al., 2007). Both membrane-bound enzymes and NAD(P)-dependent
enzymes sometimes occur in the same cell-free exract when bacterial cells are broken down and
the cell-free extract is prepared (Anthony, 1992). Some periplasmic enzymes, such as
quinoprotein methanol dehydrogenase in methylotrophs, are readily solubilized when the cell-
free extract is prepared. Given that oxidative fermentation is only functional under fairly acidic
conditions, D-gluconate oxidation fermentation is only functional under fairly acidic conditions,
D-gluconate oxidation with an NADP-dependent enzyme observed at alkaline pH is unlikely to
participate directly in keto-D-gluconate production under acidic conditions. Moreover, unlike the
NAD(P)-dependent alcohol dehydrogenase, alcohol oxidation is catalised under fairly acidic
condition at pH 3-6. Ethanol-grown cells of Acetobacter or Gluconobacter show a strong
ethanol-oxidizing activity with the membrane fraction while a little enzyme ctivity of NAD(P)-
dependent alcohol dehydrogenase is observed (Adachi et al., 2007)
Ethanol oxidation with an NAD(P)-dependent alcohol dehydrogenase usually shows a pH
optimum under highly alkaline conditions at pH 9-11 and aldehyde reduction to alcohol
favourably occurs under acidic conditions at pH 5-7
Although FAD is linked covalently to FAD-dependent enzymes and PQQ is tightly
bound to enzyme proteins (though all PQQ-dependent enzyme (quinoproteins) contain PQQ as
dissociable form), most of the membrane-bound dehydrogenases indicated earlier were stable
and active without exogenous addition of the responsible coenzyme, giving the impression that
they were coenzyme-independent or NAD(P)-independent dehydrogenases.
Most PQQ and FAD- dependent dehydrogenase can be assayed using artificial electron
acceptors such as potassium ferricyanide or phenazine methosulfate (PMS) (Emeyama, 1982). In
the case of potassium ferricianide, enzyme activity can be assayed over a broad pH range, from
highly acidic to highly alkaline conditions. On the other hand, enzyme activity measurement
43
with PMS combined with dichlorophenol indophenols (DCIP) is invalid at acidic pH below 6
due to non-enzymatic decolorization of elelctron acceptor used. Thus the assay with PMS-DCIP
is valid in the neutral to alkaline region (Adachi et al., 2007).
It is also worthy noting that when ezymes containing a heme c component in the enzyme
molecule or membrane fraction are used, the enzyme activity can be easily assayed with
potassium ferricanide. However, enzyme activity measurement with PMS-DCIP is invalid if the
enzymes do not contain the heme c component after solubilsation from the membrane (Adachi et
al., 2007).
1.13. Other Alcohol dehydrogenases
A dehydrogenase is an enzyme that oxidizes a substrate by a reduction reaction that
transfers one or more hydrides (H-) to an electron acceptor, usually NAD+/NADP+, flavin
coenzyme such as FAD or FMN or to pyroloquinoline quinone cofactor. Examples include
aldehyde dehydrogenase, acetaldehyde dehydrogenase, alcohol dehydrogenase, glutamate
dehydrogenase (an enzyme that can convert glutamate to α-ketoglutarate and vice versa), lactate
dehydrogenase, pyruvate dehydrogenase (common enzyme that feeds the TCA cycle in
converting pyruvate to acetyl-CoA), glucose dehydrogenase (involved in the pentose phosphate
pathway), glyceraldehydes-3-phosphate dehydrogenase (involved in glycolysis) or sorbitol
dehydrogenase. In TCA cycle, we have isocitrate dehydrogenase, alpha-ketoglutarate
dehydrogenase, succinate dehydrogenase and malate dehydrogenase (Murray et al., 2000)
Alcohol dehydrogenases (ADH) (EC 1.1.1.1) are a group of dehydrogenase enzymes that occur
in many organisms and facilitate the interconversion between alcohols and aldehydes or ketones
with the reduction of nicotinamide adenine dinucleotide (NAD+ to NADH). In humans and many
other animals, they serve to break down alcohols that otherwise are toxic, and they also
participate in generation of useful aldehyde, ketone, or alcohol groups during biosynthesis of
various metabolites. In yeast, plants, and many bacteria, some alcohol dehydrogenases catalyze
the opposite reaction as part of fermentation to ensure a constant supply of NAD+. The first ever
isolated alcohol dehydrogenase (ADH) was purified in 1937 from Saccharomyces cerevisiae
(baker's yeast) (Negelein and Wulff, 1937). Many aspects of the catalytic mechanism for the
horse liver ADH enzyme were investigated by Hugo Theorell and coworkers (Theorell and
44
McKee, 1961). ADH was also one of the first oligomeric enzymes that had its amino acid
sequence and three dimensional structure determined (Jornvall and Harris, 1970; Branden et
al.,1973; Hellgren, 2009). The alcohol dehydrogenases comprise a group of several isozymes
that catalyse the oxidation of primary and secondary alcohols to aldehydes and ketones,
respectively, and also can catalyse the reverse reaction (Sofer and Martin, 1987). In mammals
this is a redox (reduction/oxidation) reaction involving the coenzyme nicotinamide adenine
dinucleotide (NAD+) and cofactor Pyrroloquinoline Quinone, PQQ. Alcohol dehydrogenase is a
dimer with a mass of 80 kDa (Hammes-Schiffer and Benkovic, 2006).
1.13.1. Human Alcohol dehydrogenase
In humans, alcohol dehydrogenase exists in multiple forms as a dimer and is encoded by at least
seven different genes. There are five classes (I-V) of alcohol dehydrogenase, but the hepatic
form that is primarily used in humans is class 1. Class 1 consists of α, β, and γ subunits that are
encoded by the genes ADH1A, ADH1B, and ADH1C (Sultatos et al., 2004). The enzyme is
present at high levels in the liver and the lining of the stomach (Farres et al., 1994). It catalyzes
the oxidation of ethanol to acetaldehyde as shown in the reaction below.
CH3CH2OH + NAD+ → CH3CHO + NADH + H+
This allows the consumption of alcoholic beverages, but its evolutionary purpose is probably the
breakdown of alcohols naturally contained in foods or produced by bacteria in the digestive tract
(Kovacs and Stoppler, 2011)
Another evolutionary purpose may be metabolism of the endogenous alcohol vitamin A (retinol),
which generates the hormone retinoic acid, although the function here may be primarily the
elimination of toxic levels of retinol (Duester, 2008; Hellgren et al., 2007)
1.13.2. Yeast and bacteria alcohol dehydrogenase
Unlike humans, yeast and bacteria (except lactic acid bacteria, and E. coli in certain conditions)
do not ferment glucose to lactate. Instead, they ferment it to ethanol and CO2. The overall
reaction can be seen below:
45
Glucose + 2 ADP + 2 Pi → 2 ethanol + 2 CO2 + 2 ATP + 2 H2O (Cox and Nelson, 2005)
In yeast (Leskovac et al., 2002) and many bacteria, alcohol dehydrogenase plays an important
part in fermentation: pyruvate resulting from glycolysis is converted to acetaldehyde and carbon
dioxide, and the acetaldehyde is then reduced to ethanol by an alcohol dehydrogenase called
ADH1. The purpose of this latter step is the regeneration of NAD+, so that the energy-generating
glycolysis can continue. Humans exploit this process to produce alcoholic beverages, by letting
yeast ferment various fruits or grains. It is interesting to note that yeasts can produce and
consume their own alcohol.
1.13.3. Plant alcohol dehydrogenase
In plants, ADH catalyses the same reaction as in yeast and bacteria to ensure that there is a
constant supply of NAD+. Maize (Zea mays) has two versions of ADH: ADH1 and ADH2,
Arabidopsis (Arabidopsis thaliana) contains only one ADH gene. The structure of Arabidopsis
ADH is 47% conserved, relative to ADH from horse liver. Structurally and functionally
important residues, such as the seven residues that provide ligands for the catalytic and
noncatalytic zinc atoms, however are conserved suggesting that the enzymes have a similar
structure (Chang and Meyerowitz, 1986). ADH is constitutively expressed at low levels in the
roots of young plants grown on agar, if the roots lack oxygen, the expression of ADH increases.
Its expression is also increased in response to dehydration, low temperatures and to abscisic acid
and it plays an important role in fruit ripening, seedling and pollen development (Thompson et
al., 2010). Differences in the sequences of ADH in different species have been used to create
phylogenies showing how closely related different species of plants are (Jarvinen et al., 2010). It
is an ideal gene to use due to its convenient size (2–3 kb in length with an approximately 1000
nucleotide coding sequence) and low copy number (Thompson et al., 2010).
1.13.4. Iron-containing alcohol dehydrogenase
Another family of alcohol dehydrogenases is iron-containing ones. They occur in bacteria and
fungi. In contrast to human and plant alcohol dehydrogenases described above, these enzymes
are oxygen-sensitive. Members of the iron-containing alcohol dehydrogenase family include
46
Saccharomyces cerevisiae alcohol dehydrogenase 4 (gene ADH4), Zymomonas mobilis alcohol
dehydrogenase 2 (gene adhB) (Conway et al., 1987), Escherichia coli propanediol
oxidoreductase EC 1.1.1.77 (gene fucO), an enzyme involved in the metabolism of fucose
(hexose deoxy sugar with the chemical formula C6H12O5) and which also seems to contain
ferrous ion(s) (Conway and Ingram,1989), Clostridium acetobutylicum NADPH- and NADH-
dependent butanol dehydrogenases EC 1.1.1.- (genes adh1, bdhA and bdhB),enzymes which
have activity using butanol and ethanol as substrates (Walter et al.,1992), E. coli alcohol
dehydrogenase (gene: adhE), an iron-dependent enzyme which harbours three different
activities: alcohol dehydrogenase, acetaldehyde dehydrogenase (acetylating), EC 1.2.1.10, and
pyruvate-formate-lyase deactivase (Kessler et al., 1991), Bacterial glycerol dehydrogenase EC
1.1.1.6 (gene gldA or dhaD) (Truniger and Boos 1994), Clostridium kluyveri NAD-dependent 4-
hydroxybutyrate dehydrogenase (4hbd) EC 1.1.1.61, Citrobacter freundii and Klebsiella
pneumoniae 1,3-propanediol dehydrogenase EC 1.1.1.202 (gene dhaT), Bacillus methanolicus
NAD-dependent methanol dehydrogenase EC 1.1.1.244 (de Vries., 1992), E. coli and Salmonella
typhimurium ethanolamine utilization protein eutG.
AIMS AND OBJECTIVES
The aim of this study is to extract and partially purify alcohol dehydrogenase from Acetobacter
and to investigate the heat and pH stability of the enzyme.
The specific objectives include
� Isolation of alcohol dehydrogenase through:
� Ultrasonication
� Solubilization
� Partial purification of alcohol dehydrogenase from Acetobacter
� Characterization of the partially purified alcohol dehydrogenase viz checkin the effect of
pH, temperature and substrate concentration on its activity
� pH
� Temperature and
47
� Substrate concentration
� Determination of temperature stability of alcohol dehydrogenase at different temperature
values
� Determination of pH stability of alcohol dehydrogenase at different pH values
48
CHAPTER TWO
MATERIALS AND METHODS
2.1 Materials
2.1.1 Reagents
The chemicals used in the study were of analytical grade and were sourced as follows:
polyethelene glycol 6000, sephadex G-50, sephadex G-200 and Folin-Ciocalteau were obtained
from Sigma-Aldrich (USA), Bovine serum albumin (BSA), Bio Rad Laboratories (India) and
Folin-Ciocalteau from Sigma-Aldrich (USA).
2.1.2 Apparatus
Weighing balance Ohaus Dial-O-Gram, Ohaus Cooperation, N.J. USA.
Water bath: Model DK.
Magnetic stirrer: AM-3250B Surgi Friend Medicals, England.
Milling machine: Thomas Willey laboratory Mill Model 4, Anthor H (Thomas Company,
Philadelphia, USA).
Autoclave: UDAY BURDON’s Patent Autoclave, made in India.
Incubator: B and T Trimline incubator.
Centrifuge: Finland Nigeria 80-2B.
Oven Gallenkamp Hotbox, made in England.
pH meter: Ecosan pH meter, made in Singapore.
Sensitive weighing balance: B2404-5 mettler Toledo, made in Switzerland.
Uv/visible spectrophotometer: Jenway 6405
Microscope: WESO microscope.
49
Glass ware, Pyrex.
Thermometer
2.2. Methods
2.2.1. Collection of palm wine
Fresh palm wine was collected from a wine tapper at Okpaligbo-Ogu in Nsukka local
Government Area of Enugu State of Nigeria at around 6 o’clock in the morning with a sterile
container, transported to Classical Biomedical Laboratory Nsukka, and stored at room
temperature for use.
2.2.2. Collection of Microorganism
Acetobacter sp was isolated from a three day old palm wine using the method described by
Ronald (2010).
2.2.3. Preparation of medium for isolation of Acetocter aceti
The Basal medium (Acetic acid bacterium medium) for the isolation contained agar (15g), glucose (5g), yeast extract (5g), peptone (5g), MgSO4.7H2O (1g). Components were added to distilled water and volume was brought to 1000ml. the mixture was autoclaved for 20mins at 15psi pressure, 121oC in conical flasks (Ronald, 2010). Present in the medium also are 98% ethanol (10ml) and 98% glacial acetic acid (10ml) and Nystatin (250mg/ml)). The autoclaved medium was kept on the bench until the temperature lowered to about 40oC before the mixture was distributed into agar plates, allowed to cool and solidify.
2.2.4. Preparation of Carr medium for characterization of Acetobacter sp.
Carr medium contained yeast extract (3%), ethanol (2%), acetic acid (1%), bromocresol
green, (0.002%) and distilled water (1000ml) (Maal and Shafiee, 2009). Components were added
to distilled water and volume was brought to 1000ml. the mixture was autoclaved for 20 minutes
at 15psi pressure, 121oC in conical flasks.
2.2.5. Isolation of Acetobacter sp from palm wine
A sterile loopful of palm wine was streaked on the medium repeatedly until a pure culture
is obtained
50
2.2.6. Characterisation of isolated Acetobacter sp
2.2.6.1. Gram Staining
A smear of the pure organism was made onto a glass slide using wire loop and allowed to
air dry. This was heat-fixed by passing it three times through a flame. This was flooded with
crystal violet. 30secs later, the crystal violent was rinsed with water and flooded again with
iodine. 30secs later, the iodine was gently rinsed with water. The slide was held in slant position
and decolourised using alcohol. The excess alcohol was also gently rinsed off using water, and
was flooded with safarin counter stain. This was rinsed again with water. The slide was drained,
allowed to dry and examined under WESSO microscope.
2.2.6.2. Oxidase Test
Each disk was wetted with four inoculating loop of distilled water. Large mass of the
organism was transferred into the disk aseptically using a loop. Three minutes later, change in
colour was observed.
2.2.6.3. Catalase Test
A drop of hydrogen peroxide was placed on microscopic slide. Using applicator stick, the
colony of the Acetobacter was touched and used to make a smear onto the hydrogen peroxide
drop and the result was observed.
2.2.6.4. Characterization of Acetobacter sp using Carr medium
Little mass of the organism was streaked onto Carr Medium agar plate and the changes
were observed on the plate for 48hrs according to Maal and Shaifee (2009)
2.2.7. Preparation of liquid broth and mass production of Acetobacter isolate
Muller Hilton agar (3.8g) was dissolved in 100ml of water and the mixture was allowed
to settle. The containing agar settled to the bottom of the conical flask used. The mixture was
filtered and filterate was autoclaved for 20mins at 15psi pressure, 121oC in conical flasks. When
this cooled, Acetobacter isolate was transferred aseptically from the medium onto the liquid
broth, well covered and allowed to grow for 48hrs.
51
2.2.8. Harvesting of cell
Bacteria cells were harvested by centrifugation at 4,000rpm for 10 min and washed with
cold water. The cell paste were suspended in 0.01M potassium phosphate buffer, pH 6.0 (1g of
wet cell/10 ml of buffer) (Adachi et al., 1978). This is cell suspension.
2.2.9. Homogenization of cell
The harvested cells (already suspended in 0.01M potassium phosphate buffer, pH 6.0 (1g
of wet cell/10 ml of buffer)) were subjected to untrasonication using 500watt untrasonicator for 3
steps (10mins each) with intervolves of 3 minutes. This was centrifuged at 4000rpm 6 mins.
Enzyme assay was carried out on both the supernatant and pellet (Adachi et al., 1978)
2.2.10. Determination of percentage Triton X-100 Suitable for solubilization of alcohol
dehydrogenase from the membrane fraction.
The membrane fraction is suspended in 0.01M buffer, pH 6.0. Triton X-100 was added to
different concentration of 0.05.0%, 0.1%, 0.2%, 0.3%, 0.4% and 0.5% in different test tubes. The
suspensions were gently stirred for 3hrs at 0oC and centrifuged at 16000rpm for 60 mins.
Supernatant is obtained as a solubilized enzyme (Abolhassan et al., 2007). Protein concentration
and enzyme activity were determined on each of the solubilized enzyme. The percentage Triton
X-100 that gave the highest enzyme activity was used for mass solubilization of the enzyme
from membrane fraction following the same procedure.
2.2.11. Alcohol dehydrogenase Assay
Assay method was done using potassium ferricyanide as an electron acceptor and ethanol as the
substrate according to Adachi et al. (1978). The rate of reduction of ferricyanide to ferrocyanide
gives a quantitative amount of ethanol oxidation. The reaction mixture contains 0.1ml potassium
ferricyanide 0.1M, 0.6ml McIlvaine buffer 0.1M (pH 4.0), 0.1ml TritonX-100 10%, 0.1ml
ethanol 1M, enzyme solution in a total volume of 1ml. The reaction is carried out at 37oC by
addition of ethanol solution and stopped by adding 0.5ml of ferric dopanol reagent. 3.5ml of
water is further added to the reaction mixture and well mixed. The resulting stabilized Prussian
blue colour formed was measured with uv spectrophotometer: Jenway 6405 at 660nm after
52
standing for 20 min at 37oC. One unit of the enzyme activity is defined as the amount of the
enzyme catalizing the oxidation of 1μmol of ethanol per min under these assay conditions.
2.2. 12. Determination of protein concentration
Protein concentration was determined using the method described by Lowry et al. (1951).
The reaction mixture contained 0.0-1.0ml of protein stock solution (2mg/ml Bovin Serum
Albumin) in test tubes arranged in triplicates. The volume was made up to 1ml with distilled
water. But for the test mixture, 0.1ml of the enzyme solution was mixed with 0.9ml of distilled
water. In either case, 5ml of solution E was added to each tube and allowed to stand at room
temperature for 10min. Then 0.5ml of solution C (dilute Folin-Ciocalteau phenol reagent) was
added with rapid mixing. After standing for 30min, absorbance was read at 750nm using UV
spectrophotometer. Absorbance values were converted to protein concentrations by extrapolation
from the protein standard curve.
2.2.13. Determination of percentage Polyethelene glycol 6000 Suitable for precipitation of
alcohol dehydrogenase (quinine) from solution
To the solubilized enzyme solution, Polyethelene glycol 6000 was added to different
concentration of 0%, 5%, 10%, 15%, 20%, 25% and 30% in different test tubes containing same
volume of enzyme solution. After 30 minutes of stirring in an ice bath, the enzyme solution is
centrifuged at 4000*g for 1hour. The precipitate is suspended in small volume of 0.01M
potassium phosphate and protein and enzyme activity is determined for each of the concentration
value. The concentration that gave the highest activity was used for mass precipitation of the
enzyme using the same method.
2.2.14. Sephadex G-50 Column Chromatography
Enzyme precipitate is suspended in small volume of 0.01M potassium phosphate buffer and the
thick suspension is introduced onto Sephadex G-50 column. The enzyme was introduced onto
Sephadex G-50 column (1.4 × 61.50cm) pre-equilibrated with 0.01M potassium phosphate
buffer, pH 6.0. The protein was eluted with 0.02M potassium buffer, pH 6.0 The fractions with
high alcohol dehydrogenase activity, collected at a flow rate of 1 ml/ 120secs, were pooled
together for subsequent purification stage.
53
2.2.15. Sephadex G-200 gel filteration
Thee desalted enzyme was introduced onto Sephadex G-200 column (1.4 × 61.50cm) pre-
equilibrated with 0.01M potassium phosphate buffer, pH 6.0. The column was washed with 1L
of the same buffer and was eluted with 0.02M potassium phosphate buffer, pH 6.0.The fractions
with high alcohol dehydrogenase activity, collected at a flow rate of 1 ml/ 120secs, were pooled
and designated as the partially purified alcohol dehydrogenase.
2.2.16. Characterizationof the Partially Purified alcohol dehydrogenase
2.2.16.1. Optimum pH
The optimum pH for enzyme activity was determined using 0.05M sodium acetate buffer
pH 3. - 4, 0.1M McIlvain buffer, pH 5-7 and Tris-HCl buffer pH 8.0 - 10.0 at intervals of 1.0.
Alcohol dehydrogenase activity was determined using 0.6ml of each of the buffers as described
in the assay method, Section 2.2.8.
2.2.16.2. Optimum Temperature
The optimum temperature was determined by incubating the enzyme with alcohol
solution at 30-90oC for 20min and at pH 5 using 0.1M McIlvain buffer. The activity was then
assayed for using the method described in section 2.2.8.
2.2.16.3. Effect of Substrate Concentration on alcohol dehydrogenase Activity.
The effect of substrate concentration on the activity of alcohol dehydrogenase was
determined by incubating the enzyme with 20, 40, 60, 100, 120, 140, 160, 180 and 200mM
ethanol using a temperaure controlled-water bath (Model DK) at pH 5.0 and 50oC. Lineweaver-
burk plot of v
1 against
][
1
s was used to calculate maximum velocity (Vmax) and Michealis
constant (Km).
54
2.2.17 Thermodynamic studies of alcohol dehydrogenase
The thermodynamic parameters for the thermal inactivation of alcohol dehydrogenase were
determined on the basis of isothermal inactivation experiments for varying periods of time in a
temperature-controlled water bath (Model K) using a slight modification of Eze (2012). The
enzyme solution was placed in a pre-warmed tube at the specified temperature (30, 40, 50, 60,
70, 80 and 90oC). The residual enzyme activity was then measured as described in the alcohol
dehydrogenase assay method with 0.1ml of enzyme solution withdrawn using a micropipette at
every 10min time intervals. The stability of the enzyme was expressed as percentage residual
enzyme activity. Afterwards, the samples were immediately cooled at room temperature to stop
the thermal inactivation process.
2.2.18. Determination of pH stability of alcohol dehydrogenase
The pH stability for alcohol dehydrogenase was determined using each of 0.05M sodium acetate
buffer (pH 3- 4), 0.1M McIlvain buffer (pH 4 -7) and 0.05M Tris-HCl buffer (pH 7.0 – 10). Each
buffer is mixed with alcohol dehydrogenase solution in the ratio of 0.6:0.1. The residual enzyme
activity was then measured as described in section 2.2.11 with 0.7ml of the mixture withdrawn
using a micropipette at every 10min time intervals.. The stability of the enzyme was expressed as
percentage residual enzyme activity.
55
CHAPTER THREE
RESULTS
3.1 Characterization of Acetobacter
The microscopic examination, gram staining, oxidase test, catalase test and confirmatory test on
the isolated Acetobacter aceti are as shown in Table 4
Table 4. Result for characterization of Acetobacter sp
Test Result
color Milk
morphology Short rod
Gram stain Pink
Oxidase test No colour change
Catalase test Bubbles were formed
Confirmatory test using Carr medium Region colour changed from green to yellow
and back to green
Morphological studies, Gram staining, catalase test, oxidase test and the growth of the isolated
bacteria on Carr medium (Table 4) show that it has short rod morphology, gram negative,
catalase positive, oxidase negative and has the ability to overoxidize acetic acid typical of
Acetobacter aceti
56
3.2. Isolation of alcohol dehydrogenase
3.2.1. Sonication
The harvested cells (already suspended in 0.01M potassium phosphate buffer, pH 6.0 (1g of wet
cell/10 ml of buffer)) were subjected to untrasonication using 500watt untrasonicator for 3 steps
(10mins each) with intervolves of 3 minutes. The sonicated sample was centrifuged. Fig. 9 and
10 show the alcohol dehydrogenase activity and protein concentration of of the sonicated sample.
Fig. 9. Alcohol dehydrogenase activity after untrasonication
The uncentrifuged sonicated sample has the highest alcohol dehydrogenase activity, followed by
the pellet while low activity was observed in the supernatant.
57
Fig. 10. Protein concentration after ultrasonication
The protein concentration was more on the uncentrifuged sonicated sample followed by the
pellet and the supernatant had the least concentration
58
3.2.2. Solubilization profile using Triton X-100
Different concentration of triton X-100, 0.05%, 0.1%, 0.2%, 0.3%, 0.4%, and 0.5%, were used to
escertain the concentration that would give the highest yield of alcohol dehydrogenase. Fig.11
and 12 show alcohol dehydrogenase activity and protein concentration respectively.
Fig. 11. Triton X-100 Solubilization profile of Alcohol dehydrogenase.
From the result, effective solubilization of alcohol dehydrogenase increased from o.05% to 0.3%
where it had maximum activity before the activity began to decline.
59
Fig 12. Triton X-100 Solubilization profile for Protein concentration
The protein concentration after solubilization using Triton x-100 had highest yield of protein at
0.4% as against alcohol dehydrogenase activity that was more at o.3%
60
3.3. Partial purification of alcohol dehydrogenase
3.3.1 Precipitation profile using polyethelene glycol 6000 as a precipitant.
Polyethelene glycol precipitation profile was carried out to determine the concentration of the
salt that would precipitate greater percentage of alcohol dehydrogenase. The alcohol
dehydrogenase activity and protein concentration of Polyethelene Glucol 6000 precipitation
profile are shown in Fig. 13 and 14.
Fig. 13. Polyethelene glycol 6000 precipitation profile of alcohol dehydrogenase activity
The alcohol dehydrogenase activity after precipitation was more at 15% polyethelylene glycol
6000. It was this 15% concentration that was used for mass precipitation of the enzyme.
61
Fig. 14. Polyethelene glycol 6000 precipitation profile for Protein concentration
Protein concentration after precipitation was more at 20% followed by 25% polyethelylene
glycol 6000 concentration.
62
3.3.2. Column chromatography using sephadex G-50 and sephadex G-200
The enzyme was further purified using sephadex G-50 and sephadex G-200 as shown in the
Figures 15 and 16.
Fig. 15. Desalting using sephadex G-50.
The peaks with very high alcohol dehydrogenase activity were pooled together for gel filteration
using sephadex G-200
63
Fig. 16. Elution profile using sephadex G-200
In the gel filteration using sephadex G-200, highest peaks for alcohol dehydrogenase activity
were obtained in tubes 19, 21 and 22. These tubes were pooled together and designated as
partially purified alcohol dehydrogenase
64
Table 5. Summary of purification steps of alcohol dehydrogenase
Purification
step
Volume
(ml)
Protein conc. (mg/ml)
Activity
(µmole/min)
Spec.
Activity
(U/mg)
Total
Activity (U)
Purification
fold
% Yield
Crude Enzyme
Triton x-100
solubilization
Polyethelene
glycol 6000
precipitation
Gel filteration
191
155
75
20
1.63
0.41
0.336
0.321
62.20
46.38
57.00
65.38
38.87
113.17
169.64
203.66
11880.20
7188.13
4275.00
1307.50
1.00
2.91
4.36
5.23
100
60.50
35.98
11.01
The gel filteration result, the last partial purification step, shows specific activity of 203.66
U/mg, total activity of 1307.50 U, purification fold of 5.23 and percentage yield of 11.01
65
3.4. Characterization of partially purified alcohol dehydrogenase
3.4.1. Effect of pH on the activity of alcohol dehydrogenase
Effect of pH on the activity of alcohol dehydrogenase was studied using different pH values 3-10
at the interval of 1. The result is as shown in Fig. 16.
Fig. 16. Effect of pH on the alcohol dehydrogenase activity
Increase in pH from 3 to 5 was accompanied by an increase in enzyme activity, beyong which
the enzyme activity declined, making 5 the optimum pH for alcohol dehydrogenase activity.
66
3.4.2. Effect of temperature on the alcohol dehydrogenase activity
Different temperature values, 30, 40, 50, 60, 70, 80 and 90oC were used to study effect of
temperature on the alcohol dehydrogenase activity. The result is as shown in Fig. 17.
Fig.17. Effect of temperature on the alcohol dehydrogenase activity
Increase in temperature from 30oC to 50oC was accompanied by increase in enzyme activity
beyong which the enzyme activity declined, thereby making 50oC the optimum temperature for
alcohol dehydrogenase activity
67
3.4.3. Effect of ethanol concentration on alcohol dehydrogenase activity.
Different concentration of ethanol, 20, 40, 60, 80, 100, 120, 140, 160, 180, and 220mM were
used to study the effect of ethanol concentration on the activity of alcohol dehydrogenase, Fig
18.
Fig. 18. Effect of substrate concentration on the alcohol dehydrogenase activity
In Fig 18, the increase in ethanol concentration from 20mM to 120mM was accompanied by
increase in enzyme activity beyong which alcohol dehydrogenase activity declines gradually
Ethanol concentration (mM)
68
3.4.4. Double reciprocal plot to determine the kinetic parameters of alcohol dehydrogenase
Fig. 19. Lineweaver-Burk plot
Figure 19 shows the Lineweaver-Burk plot of against . the micheallis constant, Km and
maximum velocity Vmax were calculated to be 36mM and 90.9�mole/min respectively
69
3.5. Temperature and pH stability studies
3.5.1. Temperature inactivation studies
Fig. 20. Thermal inactivation of alcohol dehydrogenase.
Thermal inactivation OF alcohol dehydrogenase for 180 mins at 30 and 40oC showed little or no
change in enzyme activity but at 50, 60, 70, 80 and 90oC, enzyme rapidly lost its activity.
70
3.5.2. Determination of thermoinactivation parameters of alcohol dehydrogenase
Table 6. Thermoinactivation parameters of alcohol dehydrogenase
T (K) K (Min-1) ∆H (JMol-1) ∆G (JMol-1) ∆S (JMol-1K-1)
303
313
323
333
343
353
363
0.0001978
0.0006975
0.0037955
0.0079749
0.0086438
0.0010078
0.1018100
-97697.157
-97762.297
-97845.440
-97928.579
-98011.717
-98094.857
-98177.990
-67012.7
-65809.6
-63287.7
-63106.3
-64581.6
-66036.9
-67793.8
-101.209
-102.084
-106.998
-104.570
-97.460
-90.815
-83.702
Where T = temperature, K = thermoinactivation constant, ∆H = enthalpy change, ∆G = change
in Gibbs free energy, ∆S = enthropy change, R is the gas constant = 8.314JMol-1K-1, Ea =
activation energy for denaturation, Kb = Boltzmann constant = 1.38 × 10-23JK-1, h = Planck’s
costant = 6.6 × 10-34JMin-1.
71
3.5.3. Reactivation of alcohol dehydrogenase
After thermal inactivation of alcohol dehydrogenase at the different temperature values
for the for time intervals as described in section 2.2.8.3, the enzyme was quickly removed from
the waterbath and left at room temperature for 72 hours for reactivation. Alcohol dehydrogenase
assay was carried on the enzyme after every 24 hours. The result is as shown in Fig 21 below
Fig. 21. Reactivation of alcohol dehydrogenase
72
3.5.4. pH stability of alcohol dehydrogenase
pH stability studies was done at pH 3, 4, 5, 6, 7, 8, 9 and 10, each incubated with alcohol
dehydrogenase solution for three hours. Percentage residual activity was determined and plotted
against time of incubation. The result is shown in Fig. 22.
Fig. 22. pH stability of alcohol dehydrogenase
The enzyme was stable at pHs 3, 4 and 5 beyond whick it rapidly lost activity during a 180mins
pH stability studies.
73
CHAPTER FOUR
DISCUSSION AND CONCLUSION
4.1. Discussion
Acetic acid bacteria occur in sugar and alcoholised, slightly acid niches such as flowers,
fruits, beer, wine, cider, vinegar, souring fruit juices and honey. On these substrates, they oxidize
the sugars and alcohols, resulting to an accumulation of organic acids as final products. Acetic
acid is produced, and this is where the name of the bacterial group comes from. This was the
reason for the choice of palm wine, the sap of the oil palm tree (Elaeis guinneesis) as the source
of isolating the organism for this work. Palm wine, according to Amoa-Awua et al. (2006),
serves as a rich substrate for various types of micro-organisms to grow. Previous studies on the
microbiology of oil palm tree (E. guineensis) and R. hookeri have incriminated several bacterial
and yeast flora to be involved in the fermentation process (Okafor, 1975). Acetobacter species
were earlier isolated from oil palm wine (Faparusi, 1973; Okafar, 1975). Yeast growth
dominated by S. cerevisiae starts immediately after tapping begins and alcohol concentrations
become substantial in the product after the third day. The growth of acetic acid bacteria
involving both Acetobacter and Gluconobacter species become pronounced after the buildup in
alcohol concentrations.
Morphological studies, Gram staining, catalase test, oxidase test and the growth of the
isolated bacteria on Carr medium (Table 4) show that it has short rod morphology, gram
negative, catalase positive, oxidase negative and has the ability to overoxidize acetic acid typical
of Acetobacter sp. According to Gonzalex (2004), Acetic acid bacteria (AAB) are gram negative,
ellipsoidal (regular oval) to rod-shaped, and can occur singly, in pairs or in chains. Grouped.
AAB that had peritrichous flagella and were able to completely oxidize ethanol into CO2 and
water are grouped into the genus Acetobacter and those that had polar flagella and unable to
perform the complete oxidation into the genera Gluconobacter. The change in colour in the
region around the bacteria on Carr medium is as a result of the ability of Acetobacter to oxidize
acetic acid into CO2 and H2O especially when ethanol is exhausted in the medium. This is called
overoxidation and peculiar to Acetobacter among other genera of acetic acid bacteria.
74
After production using Muller Hilton liquid broth, the bacterial cells were subjected to
untrasonication using ultrasonicator. The alcohol dehydrogenase activity observed was extremely
low relative to the amount of protein present in the crude enzyme sample (Figure 9 and 10). The
high alcohol dehydrogenase activity in the pellet (ie membrane fraction) may suggest that the
enzyme is located in the cytoplasmic membrane the same way as other typical membrane-bound
dehydrogenase in acetic acid bacteria (Matsushita et al., 1994).
Since alcohol dehydrogenase was located in the cytoplasmic membrane, there was then
the need to solubilize the enzyme from the membrane fraction using detergent. The importance
of detergents as tools for the study of membrane proteins cannot be underestimated (Annela,
2004). They are usually vital in the isolation and purification of the protein. Of the different
types of detergents namely: ionic detergent (eg sodium dodesyl sulphate), Zwitterionic
detergents (eg (3[(3-Cholamidopropyl) dimethylammonio] propanesulfonic acid)(CHAPS)), and
nonionic detergent, solubilization of the enzyme was done using 0.3% Triton X-100 which
belongs to the group of nonionic detergent. The choice of Triton X-100 was due to its
effectiveness and at the same time does not denature proteins compared to other types of
detergent used for solubilization (Annela, 2004). Solubilization of alcohol dehydrogenase with
0.3% Triton-X-100 followed the Triton X-100 solubilization profile as explained in section 2.8.
In the solubilization profile, 0.3% of Triton X-100 gave the highest alcohol dehydrogenase
activity (Figure 11 and 12). This concentration is in contrasts with the report of Moonmangmee
and Moonmangmee (2002) and that of Abolhassan et al. (2007) who used 1% each to solubilized
glucose dehydrogenase from Gluconobacter frateiurii and alcohol dehydrogenase from
Acetobacter respectively. However, while these researchers worked solubilized within an hour,
protein was solubilized in this work for three hours in a cold ice bath. The low activity of alcohol
dehydrogenase observed at higher concentration of Triton X-100 could be that the high
concentration of the detergent inhibited the enzyme activity probably by binding to the enzyme
active site or by changing the protein conformation generally. In this study, alcohol
dehydrogenase activity increased by 475% in the cell homogenate after the solubilization using
Triton X-100.
Polyethelene glycol 6000 (a non-ionic polymer of ethelene oxide of molecular weight
6000Da) was used for precipitation of the enzyme from solution. The use of nonionic polymers
75
for the precipitation is a method that can help prevent protein denaturation and assist in removal
of detergents. Typically, larger proteins precipitate at lower concentrations of nonionic polymers.
Several watersoluble uncharged polymers used for precipitation include dextrans, polyvinyl
pyrrolidone, polypropylene glycols and polyethylene glycols (Harrison, 1993). Polyethylene
glycols (PEG) are the preferred non-ionic polymers for protein precipitation because the
viscosity of concentrated solutions is lower than other nonionic polymers (Harison, 1993). PEG
is very soluble in water due to the ether oxygens spread along the length of the polymer, which
are strong Lewis bases and form hydrogen bonds with water molecules. In addition, the
formation and equilibration of precipitates take significantly less time with PEG as the
precipitating agent than with ammonium sulfate or ethanol (Asenjo, 1990; Deutscher, 1990).
Another benefit of PEG precipitation is the removal of nonionic detergents (TritonX-100 &
Tween series) from the proteins. Often nonionic detergents improve the solubility of proteins,
especially membrane proteins, but they can interfere with downstream purification. Precipitation
with PEG can separate the proteins from these nonionic detergents. The precipitation profile
carried out showed 15% polyethelene glycol 6000 as the best concentration for precipitation of
alcohol dehydrogenase (Figure 13). This 15% concentration was therefore used for mass
precipitation of the protein.
After desalting of the precipitate using sephadex G-50 coloum chromatography (Fig. 15),
gel filteration using sephadex G-200 was carried out to further purify the protein. Alcohol
dehydrogenase assay was carried out in each of the eluted fractions containing 5ml of the
enzyme solution and the tubes that gave us maximum activity were pooled together and
designated as partially purified alcohol dehydrogenase solution (Fig 16.)
After each of the purification steps, alcohol dehydrogenase activity assay was carried out
using potassium ferricynide as an in vitro electron acceptor. The alcohol dehydrogenase could be
assayed in vitro in the presence of one of the following dyes as an electron acceptor; 2,6-
dichlorophenolindophenol, phenazine methosulfate or potassium ferricyanide. NAD or NADP
were not effective as an electron acceptor at all (Adachi et al., 1978). In the case of potassium
ferricianide, enzyme activity can be assayed over a broad pH range, from highly acidic to highly
alkaline conditions. On the other hand, enzyme activity measurement with PMS combined with
dichlorophenol indophenols (DCIP) is invalid at acidic pH below 6 due to non-enzymatic
76
decolorization of electron acceptor used. Thus the assay with PMS-DCIP is valid in the neutral to
alkaline region (Adachi et al., 2007).
Partially purified enzyme was characterized based on effects of pH change, temperature
change and increasing substrate concentration on alcohol dehydrogenase activities. An optimal
pH of 5 was obtained (Figure 17) with an optimal temperature of 50oC (Figure 18). The enzyme
lost its activity at more acidic pH and at alkaline pH. When the ethanol oxidation was assayed
with intact cells by Adachi et al., (1978), appreciable amount of enzyme activity was observed
even at pH 2. This means that the enzyme detached from the cell membrane became acid-labile
than intact cell. Abolhassan et al (2007) and Adachi et al.(1978) reported pH optima of 4 each
for membrane-bound quinonprotein alcohol dehydrogenase from a native strain of acetobacter.
These fall within a close range and indicate that the enzyme can be regulated by pH. Although
the occurrence in the inactive alcohol dehydrogenase seems to be strange with respect to alcohol
oxidation, some suggestive evidence has been reported for emergence of some kinds of inactive
forms of alcohol dehydrogenase in acetic acid bacteria. In Acetobacter, ethanol oxidation ability
was greatly decreased concomitantly with decreasing pH in culture medium (Duine et al., 1989).
So by shift in the pH of the culture medium, inactive alcohol dehydrogenase can presumably be
converted to active form and vice versa.Thus alcohol spoilage can be controlled by change in the
pH of acohol medium.
The result of temperature studies shows that as temperature increased from 30oC to 50oC,
the alcohol dehydrogenase activity increased and maximum alcohol dehydrogenase activity of
92.46 µmole/min was obtained at 50oC. Further increase in temperature beyond 50oC decreased
the alcohol dehydrogenase activity till the end of incubation. Hence optimum temperature was
50oC and was used for further studies. The decrease in enzyme activity at higher temperature
may be due to enzyme denaturation. In effect, change in temperature of the medium of alcohol
dehydrogenase can be used to modulate the catalytic activity of the enzyme.
Alcohol dehydrogenase activity increased as ethanol concentration increased until 120mM
(Figure 18) which increase in ethanol concentration did little change in the activity of alcohol
dehydrogenase. Abolhassan et al. (2007), reported ethanol saturation concentration of 100mM
which is comparable with the result of this study
77
The michaelis-Menten constant (Km) and maximum velocity (Vmax) obtained from the
Lineweaver-Burk plot of v
1 against
s
1 at 50oC and pH of 5 were found to be 36mM and
90.9µmole/min respectively.
From the thermal and pH stability studies, alcohol dehydrogenase was stable at 30oC,
40oC, pHs 4, 5 and 6 but rapidly lost its activity at temperatures values of 50, 60, 70, 80 and
90oC pHs of 3, 7, 8, 9 and 10 (Figure 20 and 22).
Using first order rate equation (equation 1), the Arhenius equation (equation 2), and equations 3,
4 and 5, Activation energy for denaturation (Ea) was calculated as -95160.015JMol-1, and
enthalpy changes (∆H), changes in Gibbs free energy (∆G) and changes in entropy (∆S) were
calculated (table 2). The values of thermodynamic inactivatioin constant increased progressively
as temperature increased from 30 to 900C indicating that alcohol dehydrogenase became thermal
unstable as temperature increased. From the calculated changes in enthalpy (ΔH), the reaction is
exothermic, changes in Gibbs free energy (ΔG) suggests a spontaneous reaction while changes in
entropy (ΔS) indicates that the entropy of the system increases with rise in temperature.
At = Aoe-kt (1)
Kd = (2)
∆H = Ea – RT, (3)
∆G = - RTlin T and (4)
T∆S = ∆H - ∆G, (5)
Regeneration studies (Figure 21) show that beyond 50oC, alcohol dehydrogenase
inactivation up to three hours, the enzyme could have been permanently denatured that
regeneration of the enzyme activity by cooling at room temperature was not possible.
78
4.2. CONCUSSION
From the results obtained in this work, it can be inferred that the detergent, triton X-100
and the salt, polyethelene glycol 6000 respectively are effective in solubilization of alcohol
dehydrogenase from the cell membrane and precipitationn of alcohol dehydrogenase from
solution.
Naturally, when palm wine is produced on a very cold day, like during harmattan season
when the environmental temperature remains cold throughout the day, palm wine usually retain
their organoleptic characteristics for a long time. The palm wine does not easily turn sour. This
knowledge has pre-existed, though the molecular mechanism for the retention of the organoleptic
characteristic has not been established. From this work, this phenomenom can be attributed to
low activity of alcohol dehydrogenase that is responsible for converting ethanol in palm wine to
acetic acid. This work is therefore a justification of the pre-existing information and an insight
into the molecular mechanism involved. The low activity of alcohol dehydrogenase both at low
and high temperature of the medium of alcohol dehydrogenase shows that tempterature can be
used to modulate the catalytic activity of the alcohol dehydrogenase.
It is also known that on the day when temperature increases rapidly with time,
deterioration of palm wine ie the souring of palm wine is very fast. This shows that as
temperature increases, alcohol dehydrogenase activity also increases concomitantly as was
observed in this work. However, this deterioration does not continue indefinitely. With time, the
rate of deterioration ceases even as time progresses. This work is also able to explain this
phenomemon. During the characterisation using pH, it was noted that at high acidic pH the
activity of alcohol dehydrogenase decreased ie the activity of alcohol dehydrogenase decreased
at lower pH values. The ceasation of deterioration at some time implies that as more acetic acid
were produced, the pH of palm wine reduced, thereby reducing the deteriorating properties of
alcohol dehydrogenase through further production of acetic acid as shown in this work. In
essence, by shift in the pH of the palm wine, inactive alcohol dehydrogenase can presumably be
converted to active form and vice versa.
79
REFERENCES
Abolhassan M.F., Sepehr, S. I. M., Shabani, A., Soudi, M.R. and Moosavi-Nejad, S.Z. (2007),
Purification and characterization of Membrane-Bound Quinoprotein Alcohol Dehydrogenase from a Native Strain of Acetobacter, Journal of Biological Sciences,
7(2): 315-320
Adachi, O., Ano, Y., Toyama, H. and Matsushita, K. (2007). Biooxidation with PQQ and FAD-Dependent dehydrogenases, In: Modern Biooxidation, Enzymes, Reactions and Application, Rolf D, S and Urlacher V.B (eds),WILEY-VCH Verlag GmbH and Co., pp 1-41.
Adachi, O., Fujii, Y., Ghaly, MF., Toyama, H., Shinigawa, E. and Matsushita, K. (2001) Membrane-bound quinoprotein D-arabitol dehydrogenase of Gluconobacter suboxydans
IFO 3257: A versatile enzyme for the oxidative fermentation of various ketoses. Bioscience, Biotechnology and Biochemistry 65: In Press
Adachi, O., Kubota, T., Hacisalihoglu, A., Toyama, H., Shinigawa, E., Duine, J.A. and
Matsushita, K. (1998) Characterization of quinohemoprotein amine dehydrogenase from Pseudomonas putida. Bioscience, Biotechnology and Biochemistry, 62: 469–478.
Adachi, O., Miyagawa, E., Shinagawa, E., Matsushita, K. and Ameyama, M. (1978). Purification
and properties of particulate alcohol dehydrogenase from Acetobacter aceti, Agricultural
and Biological Chemistry, 42: 2331-2340.
Adachi, O., Tayama, K., Shinagawa, E., Matshuta, K. and Ameyama, M. (1980). Purification and characterization of membrane-bound aldehyde dehydrogenase from Gluconobacter suboxydans. Agricultural and Biological Chemistry, 44: 503-515
Afolabi P.R., Mohammed F., Amaratunga, K., Majekodunmi, O., Dales, S.L., Gill, R., Thompson, D., Cooper, J.B., Wood, S.P., Goodwin, P.M., and Anthony, C. (2001). Site-directed mutagenesis and Xray crystallography of the PQQ-containing quinoprotein methanol dehydrogenase and its electron acceptor, cytochrome cL. Biochemistry, 40: 9799–9809
Ali, S.A. (2008) Toddy and palm wine. http://itdg.org/docs/technical informations services/toddy
palmwine.pdf. Retrieved on 15 April, 2014.
Ameh, S.J; Obodozie, O.O., Olorunfemi, O.P., Okoliko, E.I. and Ochekpe, N.A. (2011). Potetntial of gladiolus corms as antimicrobial agent in food processing and traditional medicine. Journal of Microbiology and Antimicrobials, 3(1): 8-12.
Amoa-Awua, W.K., Sampson, E. and Tano-Debrah, K. (2006). Growth of yeasts, lactic and acetic acid bacteria in palm wine during tapping and fermentation from felled oil palm (Elaeis guneensis) in Ghana, Journal of Applied Microbiology, 101: 599-606.
80
Annela M. Seddon, P. C. and Paula J. B. (2004). Membrane proteins, lipids and detergents: not just a soap opera, Biochimica et Biophysica Acta, 1666: 105– 117.
Anthony, C. (1982) The Biochemistry of Methylotrophs. Academic Press, London, pp 1 – 431.
Anthony, C. (1992). Methanol dehydrogenase in Gram-negative bacteria. In: Principles and Applications of Quinoproteins, Davidson, V.L (ed).. Marcel Dekker New York, pp 17-45
Anthony, C. (1992). The c-type cytochromes of methylotrophic bacteria. Biochimica et
Biophysica Acta, 1099: 1–15 Anthony, C. (1996) Quinoprotein-catalysed reactions. Biochemistry Journal, 320: 697–711 Anthony, C. (2000) Methanol dehydrogenase, a PQQ-containing quinoprotein dehydrogenase.
Subcellullar Biochemistry, 35: 73–118 Anthony, C. (2004), the PQQ-containing quinoporotein dehydrogenases. In: Respiration in
Archaea and Bacteria, Zannoni D (ed), Kluwer Academic Publisher,Netherland, pp 1-10. Anthony, C., and Williams, P. (2003) The structure and mechanism of methanol dehydrogenase.
Biochimica et Biophysica Acta, 1647: 18 –23
Asai, T. (1934). Acetic acid bacteria. Classification and biochemical activities. University of Tokio Press. TOkio, pp 121-126.
Asakura, A. and Hoshino, T. (1999) Isolation and characterization of a new quinoprotein dehydrogenase, L-sorbose / L-sorbosone dehydrogenase. Bioscience, Biotechnology and
Biochemistry 62: 469–478 Asenjo, J.A. (1990). Separation Processes in Biotechnology, Marcel Dekker, New York, pp 329
358. Attwood, M.M., Van Dijken, J.P. and Pronk, J. (1991). Glucose metabolism and gluconic aacid
production by Acetoboacter diazotrophicus. Journal of Fermentation Bioengineering, 72: 101-105
Avezoux, A. Goodwin, M.G. and Anthony, C. (1995). The role of the novel disulphide ring in the active site of the quinoprotein methanol dehydrogenase from Methylobacterium
extorquens. Biochemical Journal, 307:735–741 Bartowsky, E.J., Xia, D., Gibson, R.L., Fleet, G.H. and Henschke, P.A. (2003). Spoilage of
bottled red wine by acetic acid bacteria. Letter in Applied Microbiology, 36: 307-314
Bechem, E.E T., Omoloko, C., Nwaga, D. and Titanji, V.P.K. (2007). Characterization of palm wine yeasts using osmiotic, ethanol tolerance and the isozyme polymorphism of alcohol dehydrogenase, Archiv fur Mikrobiologie, 83: 237-245
81
Blake, C.C.F., Ghoshm M,, Harlosm, K., Avezoux, A. and Anthony, C. (1994) The active site of methanol dehydrogenase contains a disulphide bridge between adjacent cysteine residues. Nature and Structural Biology, 1: 102–105.
Boesch, C., Trcek, J., Sievers, M. and Teuber, M., (1998). Acetobacter intermedius, sp. Nov.
Systematic and Applied Microbiology, 21: 220-229
Brändén, C., Eklund, H., Nordström. B., Boiwe, T, Söderlund, G., Zeppezauer, E., Ohlsson, I., Akeson, A. (1973). Structure of liver alcohol dehydrogenase at 2.9-angstrom resolution". Proceedings of the National Academy of Sciences of the United States of America 70 (8): 2439–42.
Buchanan RE, Gibbons NE (1974). Bergey’s_s Manual of Determinative Bacteriology (8th ed.). The Williams and Wilkins Co., Baltimore. pp. 267-278.
Buchanan RE, Gibbons NE (1984). Family VI. Acetobacteraceae. In: Bergey’s_s Manual of
Systematic Bacteriology, Vol.1 (9th ed.). Holt JG (eds). The Williams and Wilkins Co., Baltimore, pp. 267-278.
Buttner, T., Geier, J., Frison, G., Harmer, J., Calle, C., Schweiger, A., Schonberg, H., and Gru
tzmacher, H. (2005) Science 307: 235–238.
Caetano-Anolles, G., Bassam, B. J. and Gresshoff, P.M. (1991). DNA amplification fingerprinting using very short arbitrary oligonucleotide primers. Biotechnology, 9: 553-557.
Carlotti, A. and Funke, G. (1994). Rapid distinction of Brevibacterium species by restriction
analysis of rDNA generated by polymerase chain reaction. Systematic and Applied
Microbiology, 17: 380-386 Carr, J.G. and Passmore, S.M. (1979). Methods for identifying acetic acid bacteria. In,
Identification methods for microbiologists F.A. Skinner and D.W. Lovelock (ed). Academic Press, London, p 333-347
Chang, C and Meyerowitz, E.M. (1986). Molecular cloning and DNA sequence of the Arabidopsis thaliana alcohol dehydrogenase gene". Proceedings of the National
Academy of Sciences of the United States of America 83 (5): 1408–1412.
Chen, Z-W, Matsushita, K., Yamashita, T., Fujii, T., Toyama, H., Adachi, O., Bellamy, H. and Mathews, SF. (2002) Structure at 1.9 Å resolution of a quinohemoprotein alcohol dehydrogenase from Pseudomonas putida HK5. Structure 10: 1–20
Choi, O., K. Jinwoo, K. Jung-Gun, J. Yeonhwa, J. S. Moon, C. S. Park and I. Hwang, (1995) Pyrroloquinoline Quinone Is a Plant Growth Promotion Factor Produced by Pseudomonas fluorescens B161. Plant Physiology, 146: 657–668 .
82
Cleenwerck, I., Dellallglio, F., Felis, G.E., ENgelbeen, K., Jansens,D. and Marzotto, M. (2005). Description of Gluconacetobacter sweingsii sp. Nov. and Gluconacetobacter rhaeticus sp. Nov., isolated from Italian apple fruit. Vinegars and Acetic Acid Bacteria Internation Symposium. Reggio Emilia, 2005.
Conway, T. and Ingram, L.O. (1989). "Similarity of Escherichia coli propanediol oxidoreductase (fucO product) and an unusual alcohol dehydrogenase from Zymomonas mobilis and Saccharomyces cerevisiae". Journal of. Bacteriology. 171 (7): 3754–9.
Conway T, Sewell GW, Osman YA, Ingram LO (June 1987). "Cloning and sequencing of the alcohol dehydrogenase II gene from Zymomonas mobilis". Journal of. Bacteriology. 169 (6): 2591–7.
Cozier, G.E, Salleh, RA. and Anthony, C. (1999). Characterization of the membrane glucose dehydrogenase from Escherichia coli and characterization of a site directed mutant in which His262 has been changed to tyrosine. Biochemical Journal, 340: 639–647.
Datta, S., Mori, Y., Takagi, K., Kawaguchi, K., Chen, Z.W., Okajima, T., Kuroda, S., Ikeda, T.,
Kano, K., Tanizawa, K. and Mathews, FS. (2001). Structure of a quinohemoprotein amine dehydrogenase with an uncommon redox cofactor and highly unusual crosslinking. Proceedings of National Academy of Science 98: 14268–14273
Davidson, V. L. (1993) Electron transfer in quinoproteins Archives for Biochemical and
Biophyisical, 428: 32– 40
Davidson, V.L. (2000) Methylamine dehydrogenase: structure and function of electron transfer complexes. Subcellullar Biochemistry, 35: 119–144
De Ley, J. (1959). On the formation of acetoin by Acetobacter. Journal of General
Microbiology, 21: 352-365 De Ley, J., Gossele, F. and Swings, J. (1984). Genus I Acetobacter. In: Bergey’s Manual of
Systematic Bacteriology. Vol 1, Williams and Wilkens, Maryland, U.S.A. pp. 268-274. De Vero, L., Giudici, P. (2008). Genus-specific profile of acetic acid bacteria by 16S rDNA
PCR-DGGE. Internation. Journal of. Food Microbiogy. 125(1): 96- 101.
De Vries, E., Arfman, N., Terpstra, P., Dijkhuizen, L. (1992). Cloning, expression, and sequence analysis of the Bacillus methanolicus C1 methanol dehydrogenase gene. Journal of.
Bacteriology. 174 (16): 5346–53
DeJong, G.A.H., Caldeira, J., Sun, J., Jongejan, J.A., Devries, S., Loehr, T.M., Moura, I., Moura, J.J.G. and Duine, J.A. (1995a) Characterization of the interaction between PQQ and heme C in the quinohemoprotein ethanol dehydrogenase from Comamonas testosteroni. Biochemistry 34: 9451–9458.
83
DeJong, G.A.H., Geerlof, A., Stoorvogel, J., Jongejan, J.A., Devries, S. and Duine, J.A. (1995b)
Quinohaemoprotein ethanol dehydrogenase from Comamonas testosteron—purification, characterization, and reconstitution of the apoenzyme with pyrroloquinoline quinone analogues. European Journal of Biochemistry. 230: 899–905.
Deppenmeier, U., Hoffmeister, M. and Prust, C. (2002). Biochemistry and biotechnological
applications of Gluconobacer strains. Applied Microbiology and Biotechnology 60:, 233-242.
Dijkstra, M., Frank, J., and Duine, J. A. (1989) studies on electron transfer from methanol
dehydrogenase to cytochrome cl both purified from Hyphomicrobium X. Biochemical
Journal, 257: 87–94
Drysdale, G.S. and Fleet, G.H. (1989a). The growth and survival of acetic aacid bacteria in wines at different concentration of oxygen. American Journal of Enology and Viticulture, 40: 99-105.
Drysdale, G.S. and Fleet, G.H. (1989b). the effect of acetic acid bacteria upon the growth and
metabolism of yeast during the fermentation of grape juice. Journal of Applied
Bacteriology, 67: 471-481. Drysdale, G.S. and Fleet, G.H.I. (1988). Acetic acid bacteria in winemaking: A Review.
American Journal of Enology and Viticulture, 39: 143-154. Du Toit, W.J. and Lamberchts, M.G. (2000). The enumeration and idenfication of acetic acid
bacteria from South African red wine fermentations. International Journal of Food
Microbiology, 74: 57-64. Du Toit, W.J. and Pretorius, I.J. (2002). The occurrence , control and esoteric effect of acetic
acid bacteria in winemakinig. Annals of Microbiology, 52: 155-179. Du Toit, W.J., Pretorius, I.J. and Lonvaud-Funel, A. (2005). The effect of sulphur dioxide and
oxygen on the viability and culturability of a strain of Acetobacter pasteurianus and a strain of Brettanomycces bruxellensis isolated from wine. Journal of Applied
Microbiology, 98: 862-871. Duester, G. (2008). Retinoic acid synthesis and signaling during early organogenesis. Cell, 134
(6): 921–31.
Duine, J. A. (1991). Quinoproteins: enzymes containing the quinonoid cofactor pyrroloquinoline quinone, topaquinone or tryptopha tryptophan quinone. European Journal of
Biochemistry 200:271-284.
84
Duine, J. A. (1999) The Pyroloquinoline quinone story, Journa of Bioscience and Bioengineering. 88: 231–236
Duine, J. A. and Joengjan, J. (1989). Quinoproteins, enzyms with pyrroloqunoline quinone as cofactor. Annual Review of Biochemistry, 58, 403-426
Duine, J. A., and Frank, J. (1980) The prosthetic group of methanol dehydrogenase. Purification and some of its properties, Biochemical. Journal, 187: 221–226
Duine, J. A., Frank, J., and De Beer, R. (1984) An electron-nuclear double-resonance study of methanol dehydrogenase and its coenzyme radical Archives for Biochemical and.
Biophysical, 233: 708 –711
Elias, MD, Tanaka, M., Izu, H., Matsushita, K., Adachi, O. and Yamada, M. (2000). Functions of amino acid residues in the active site of Escherichia coli pyrroloquinoline quinonecontaining quinoprotein glucose dehydrogenase. Journal of Biological
Chemistry, 275: 7321–7326. Elias, MD, Tanaka, M., Sakai, M., Toyama, H., Matsushita, K., Adachi, O. and Yamada, M..
(2001). C-terminal periplasmic domain of Escherichia coli quinoprotein glucose dehydrogenase transfers electrons to ubiquinone. Journal of Biological Chemistry, 276: 48356–48361
Emeyama, M. (1982). Microdetermination of D-glucose, D-fructose, D-gluconate, 2-keto-D-gluconate, aldehyde, and alcohol with membrane-bound dehydroganse, Methods in
Enzymology, 89: 20-29.
Entani, E., Ohmori, S., Masai, H. and Suzuki, K.I. (1985). Acetobacter polyoxogenes sp. Nov., a new species of an acetic acid bacterium useful for producing vinegar with high acidity. J
Gen Appl Microbiol 31: 475-490. Enwefa, C., Uwajeh, R. and Oduh, R. (2004). Some studies on Nigerian palm wine with special
reference to yeasts. Acta Biotechnological, 12(2): 117-125.
Eze, O.O. (2012). The kinetic analysis of the thermostability of peroxidase from African oil bean (Pentaclethra macrophylla Benth) seeds, Journal of Biochemical Technology, 4(1): 459-463.
Faparusi, S.I. (1973). Origin of initial microflora of palm wine from oil palm trees (Elaeis
guineensis). Journal of. Applied Bacteriology., 36: 559-565.
Farrés, J., Moreno, A., Crosas, B., Peralba, J.M., Allali-Hassani, A., Hjelmqvist, L., Jörnvall, H., Parés, X., (1994). Alcohol Dehydrogenase of Class IV (σσ-ADH) from Human Stomach cDNA Sequence and Structure/Function Relationships". European Journal of
Biochemistry, 224 (2): 549–557.
85
Frank, J., Dijkstra, M., Duine, J. A., and Balny, C. (1988). Kinetic and spectral studies on the redox forms of methanol dehydrogenase from Hyphomicrobium X, European Journal of
Biochemistry, 174: 331–338
Franke, I.H., Fegan, M., Hayward, C., Leonard, G., Stakebrandt, E. and Sly L.I. (1999). Description of Gluconacetobacter sacchari sp. Nov., a new species of acetic acid bacterium isolated from the leaf sheath of sugar cane and from the pink sugar cane mealy bug. Internation Journal of Systematic Bacetiology, 49, 1681-1693.
Frateur, J. (1950). Essai sur la sytematique des Aceto bacters. La Cellule 53: 287 Frebortova, J., Matsushita, K., Arata, H. and Adachi, O. (1998) Intramolecular electron transport
in quinoprotein alcohol dehydrogenase of Acetobacter methanolicus: A redox-titration study. Biochimica and Biophysica Acta, 1363: 24–34.
Geerlof, A., Stoorvogel, J., Jongejan, J.A., Leenen, E.J.T.N., Vandooren, T.J.G.M.,
Vandentweel, W.J.J. and Duine, J.A. (1994) Studies on the production of (s)-(+)-solketal (2,2-dimethyl-1,3-dioxolane-4-methanol) by enantioselective oxidation of racemic solketal with Comamonas testosteroni. Applied Microbiology and Biotechnology 42: 8–15.
Ghosh, M., Anthony, C., Harlos, K., Goodwin, M.G. and Blake, C.C.F. (1995). The refined
structure of the quinoprotein methanol dehydrogenase from Methylobacterium
extorquens at 1.94 Å.Structure, 3: 177–187. Gillis, M., Kersters, K., Hoste, B., Janssens, D., Droppenstedt, M., Stephan, M.P., Teixeira,
K.R.S., Dobereiner, J. and De Ley, J. (1989). Acetobacter diazotrophicus sp. nov., a nitrogen fixing acetic acid bacterium associated with sugar cane. International Journal of
Systematic Bacteriology, 39: 361-364
Gomez-Manzo, S., Contreras-Zentella, M., Gonzalez-Valdez, A., Sosa-Torres, M., Arreguin Espinoza, R. and Escamilla-Marvan, E. (2008). The PQQ-alcohol dehydrogenase of Gluconacetobacter diazotrophicus". Internationa. Journal of. Food Microbiology. 125: 71–78.
Gonzalex, A., Hierro, N., Guillamon, J.M., Mas, A. and Poblet, M. (2004). Applications of molecular methods for the differentiation of acetic acid bacteria in a red wine fermentation, Journal of Applied Microbiology, 96: 853-860.
Goodwin, MG. and Anthony, C. (1996) Characterization of a novel methanol dehydrogenase containing barium instead of calcium. Biochemical Journal, 318: 673–679.
Greenshields, R.N. (1978). Acetic acid: vinegar. In: Primary Products Metabolism, Economic
Microbiology, Vol 2, Rose, A.H. (ed), London Academic Press, London, pp 121-186.
86
Groen, B.W., van Kleef, M.A.G. and Duine, J.A. (1986) Quinohaemoprotein alcohol dehydrogenase apoenzyme from Pseudomonas testosteroni. Biochemical Journal, 234: 611–615.
Grossman, M.K. and Becker, R. (1984). Investigation on bacterial inhibition of wine
fermentation. Kellerwirtschaft, 10: 272-275.
Hammes-Schiffer, S. and Benkovic, S.J. (2006). Relating protein motion to catalysis. Annual
Review of Biochemistry 75: 519–541.
Harrison, Roger G. (1993), Protein Purification Process Engineering, Marcel Dekker, New York, pp 115-208.
Hauge, J.G. (1964) Glucose dehydrogenase of Bacterium anitratum: An enzyme with a novel
prosthetic group. Journal of Biological Chemistry, 239: 3630—3639
Hellgren, M. (2009). Enzymatic studies of alcohol dehydrogenase by a combination of in vitro
and in silico methods, Ph.D. thesis. Stockholm, Sweden: Karolinska Institute. p. 70.
Hellgren, M., Strömberg, P., Gallego, O., Martras, S., Farrés, J., Persson, B., Parés, X., Höög, J.O. (2007). "Alcohol dehydrogenase 2 is a major hepatic enzyme for human retinol metabolism.". Cellular and molecular life sciences, 64 (4): 498–505.
Holt, J.M., Krieg, N.R., Sneath, P.H.A., Staley, J.Y. and Williams, S. T. (1994). Genus
Acetobacter and Gluconobacter. In: Bergey’s Manual of Determinative Bacteriology (9th edn), Williams and Wilkens, Marylands, pp, 71-84.
Hopper, D.J. and Rogozinski, J. (1998). Redox potential of the haem c group in the
quinocytochrome, lupanine hydrolase, an enzyme located in the periplasm of a Pseudomonas sp. Biochimica and Biophysica Acta, 1383: 160–164
Houck, D. R., Hanners, J. L. and Unkefer, C. J. (1991). Biosynthesis of pyrroloquinoline quinone. Biosynthetic assembly from glutamate and tyrosine. Journal of American.
Chemical Society, 113:3162-3166.
Hyun, Y.L. and Davidson, V.L. (1995) Mechanistic studies of aromatic amine dehydrogenase, a tryptophan tryptophylquinone enzyme. Biochemistry, 34: 816–823
Jarvinen, P., Palme, A., Orlando, M.L., Lannenpaa, M., Keinanen, M., Sopanen, T., Lascoux, M. (2010). "Phylogenetic relationships of Betula species (Betulaceae) based on nuclear ADH and chloroplast matK sequences - American Journal of Botany, 91 (11): 1834.
Jojima, Y., Mihara, Y., Suzuki, S., Yokozeki, K., Yamanaka, S. and Fudou, R. (2004). Saccharibacter floricola gen. nov., sp. nov., a novel osmophilic acetic acid bacterium
87
isolated from pollen. Internationa Journal of Systematic Evolutionary Microbiology, 54: 2263-2267.
Jongejan, A., Jongejan, J.A. and Duine, J.A. (1998). Homology model of the quinohaemoprotein
alcohol dehydrogenase from Comamonas testosteroni. Protein Eng 11: 185–198
Jörnvall, H., Harris, J.I., (1970). Horse liver alcohol dehydrogenase. On the primary structure of the ethanol-active isoenzyme". European Journal of Biochemistry, 13 (3): 565–76.
Joyeux, A., Lafon-Lafourcade, S. and Ribereu-Gayon, P. (1984b). Metabolism of acetic acid bacteria in grape must: consequences on alcoholic and malolactic fermentation. Sci
Aliments 4: 247-255. Joyeux, A., Lafon-Lafourcase, S. and Ribereu-Gayon, P. (1984a). Evolution of acetic acid
bacteria during fermentation and storage of wine. Applied Environronmental
Microbiology, 48: 153-156. Katsura, K., Kawasaki, H., Potachaoren, W., Saono, Seki, T., Yamada, Y., Uchimura, T. and
Komagata, K. (2001). Asaia siamensis sp. nov., an acetic aacid bacterium in the alpha proteobacteria. Internationa Journal of Systematic Evolutionary Microbiology, 5: 559-563.
Keitel, T., Diehl, A., Knaute, T., Stezowski, J.J., Hohnem,W. and Gorisch, H. (2000). X-ray
structure of the quinoprotein ethanol dehydrogenase from Pseudomonas aeruginosa: Basis of substrate specificity. Journal of Molecular Biology, 297: 961–974
Kessler, D., Leibrecht, I., Knappe, J.(1991). "Pyruvate-formate-lyase-deactivase and acetyl CoA reductase activities of Escherichia coli reside on a polymeric protein particle encoded by adhE". FEBS Letter, 281 (1-2): 59–63.
Kocher, G.S., Kalra, K.L. and Phutela (2006). Comparative Production of Sugarcane Vinegar by Different Immobilization Techniques, Journal of the Institute of Brewing, 112 (3): 264–266.
Kondo, K. and Horinouchi, S. (1997) Characterization of the genes encoding the three component membrane-bound alcohol dehydrogenase from Gluconobacter suboxydans
and their expression in Acetobacter pasteurianus. Appl Environ Microbiol 63:1131–1138. Kovacs, B., Stöppler, M.C. (2011) "Alcohol and Nutrition". MedicineNet, Inc. Archived from
the original on 23 June 2011. Retrieved 2011-06-07. Lasko, D.R., Zamboni, N. and Sauer, U. (2000). Bacterial response to acetate challenge:a
comparison of tolerance among species. Appl Microbiol Biotechnol 54: 243-247. Leskovac, V., Trivic, S., Peričin, D. (2002). "The three zinc-containing alcohol dehydrogenases
from baker's yeast, Saccharomyces cerevisiae". FEMS Yeast Research, 2 (4): 481–494.
88
Lisdiyanti, P., Kawasaki, H., Seki, T., Yamada, Ya., Uchimura, T. and Komagata, K. (2001). Identification of Acetobacter strain isolated from Indonesia sources and proposals of Acetobacter syzgii sp. nov., Acetobacter cibinongenesis sp. nov. and Acetobacter orientalis sp nov. Journal of General Applied Microbiology, 47: 119-131.
Lisdiyanti, P., Kawasaki, H., Widyastuti, Y., Saono, S., Seki, T., Yamada, Y., Uchimura, T. and
Komagata, K. (2002). Kozakia baliensis sp. nov., a novel acetic acid bacterium in the α-Proteobacteria. Internation Journal of systematic Evolutionary Microbiology, 52: 813-818.
Longanathan, P. and Nair, S. (2004). Swaminathania salitolerans gen. nov., sp. nov., a salt-
tolerant, nitrogen-fixing and phosphate-solubilizing bacterium from wild rice (Porteresia coarctata Tateoka). Internation Journal of systematic Evolutionary Microbiology, 54: 1185-1190.
Lowry, O.H., Rosebrough, N.J., Farr, A. l. and Randall. R.J .(1951). Protein measurements
with follin –phenol reagents. Journal of Biological Chemistry, 93:265-275.
Lu, S.F., Lee, F.L. and Chen, H.K. (1999). A thermotolerant and high acetic acid-producing bacterium Acetobacter sp. J Appl Microbiol 86: 55-62.
Maal, B. and Shafiee, R. (2009). Isolation, and identification of an acetobacter strain from
Iranina White-Red Cherry with High acetic acid productivity as a potential strain for cherry vinegar production in food and agriculture biotechnology, World Acedemy of
Science, Engineering and Technology, 30: 201-204.
Margalith, P.Z. (1981). Flavor Microbiology. Charles Thomas (ed), Illinois. U.S.A. Matsushita, K., Takaki, Y., Shinagawa, E., Ameyama, M. and Adachi, O. (1992). Ethanol
oxidase respiratory chain of acetic acid bacteria-reactivity with ubiquinone of pyrroloquinoline quinone-dependent alcohol dehydrogenases purified from Acetobacter
aceti and Gluconobacter suboxydans. Bioscience Biotechnology and Biochemistry, 56: 304–310
Matsushita, K., Toyama, H., Adachi, O (1994). Respiratory chains and bioenergetics of
acetic acid bacteria. Advances in Microbial Physiology, 36: 247–301.
Matsushita, K. and Adachi, O. (1993) Bacterial quinoproteins glucose dehdyrogenase and alcohol dehydrogenase. In: Principles and Applications of Quinoproteins, Davidson VL (ed), Marcel Dekker, New York pp 47–63.
Matsushita, K., Honobe, M., Shinagawa, E., Adachi, G. and Ameyama, M. (1985). Isolation and
characterization of outer and cytoplasmatic membranes from spheroplasts of Acetobacter. Antonie van Leewenhoek 20: 102.
89
Matsushita, K., Toyama, H. and Adachi, O. (1994) Respiratory chains and bioenergetics of acetic acid bacteria. Adv Microbial Physiol 36: 247–301
Matsushita, K., Toyama, H., Yamada, M. and Adachi, O. (2002). Quinoproteins: structure, function, and biotechnological applications. Applied Microbiology and Biotechnology, 58: 13–22.
Matsushita, K., Yakushi, T., Toyama, H., Shinagawa, E. and Adachi, O. (1996) Function of multiple heme c moieties in intramolecular electron transport and ubiquinone reduction in the quinohemoprotein alcohol dehydrogenase cytochrome c complex of Gluconobacter
suboxydans. J Biol Chem 271: 4850–4857. Matsushita, K., Yamashita, T., Aoki, N., Toyama, H. and Adachi, O. (1999) Electron transfer
from quinohemoprotein alcohol dehydrogenase to blue copper protein azurin in the alcohol oxidase respiratory chain of Pseudomonas putida HK5. Biochemistry, 38, 6111–6118.
Menzel, U. and Gottshalk, G. (1985). The internal pH of Acetobacter wieringae and Acetobacter
aceti during growth and production of acetic acid . Archives of Microbiology, 143: 47-51. Meunier, J.R. and Grimont, P.A.D. (1993). Factors affecting reproducibility of random amplified
polymorphic DNA fingerprinting. Researches in Microbiology, 144: 373-379. Miyazaki, T., Tomiyama, N., Shinjoh, M. and Hoshino, T. (2000) Molecular cloning and
functional expression of D-sorbitol dehydrogenase from Gluconobacter suboxydans
IF03255, which requires pyrroloquinoline quinone and hydrophobic protein SldB for activity development in E. coli. Bioscience, Biotechnology and Biochemistry, 66: 262–270.
Moonmangmee, D. and Moonmangmee, S. (2012) Purification and characterization of
membrane-bound glucose dehydrogenase from mutant Gluconobacter frateurii THD32N, Ist Mae Fah Luang University International Conference
Muraoka, H., Watab, Y., Ogasawara, N. and Takahashi, H. (1983). Trigger damage by oxygen deficiency to the acid production system during submerged acetic acid fermentation with Acetobacter aceti. Journal of Fermentation Technology, 61, 89-93.
Murray, K.M., Granner, D.K., Mayes, P.A. and Rodwell, V.W. (2000). Harper’s Biochemistry (25th edition), Appleton and Lange, New York, pp 182-189
Mutzel, A. and Gorisch, H. (1991) Quinoprotein ethanol dehydrogenase: preparation of the apo form and reconstitution with pyrroloquinoline quinone and Ca2+ or Sr2+ ions. Agric Biol
Chem 55: 1721–1726 Namba, A., Tamura, A. and Nagai, S. (1984). Synergistic effects on acetic acid ad ethanol on the
growth of Acetobacter sp. J Ferment Technol 62: 501-505.
90
Negelein, E. and Wulff, H.J. (1937). Diphosphopyridinproteid ackohol, acetaldehyd. Biochemistry, 293: 351.
Ojimelukwe, P. C. (2002). Effect of preservation with saccoglottis. Gabonesis on the
microbiology of fermenting palm wine, Journal of Food Biochems, 25: 411-424
Okafor, J.C. (1975). Varietal delimination in Irvingia gabonensis (Irvingiaceae). Bulletin du
Jardin Botanique Nationale de Belgique, 45(1-2): 211-221. Okunade, AL, Clark AM, Hufford, CD, Oguntimein, B.O. (1999). Azaanthraquinone: an
antimicrobial alkaloid from Mitracarpus scaber, Planta Med., 65(5): 447-448. Olijve, W. and Kok, J.J. (1979). Analysis of growth of Gluconobacter oxydans in glucose
containing media. Arch of Microbiology, 121: 283-290. Onwuka, U. N. (2011). Performance evaluation of ohmic heating under a static medium on the
pasteurization and quality parameters of palm wine (Raphia Hokeri). Journal of
Emerging Trends in Egineering and Applied Science, 2(1): 160-165
Opara, C.C., Ajoku, G. and Madumelu, N.O. (2013). Palm wine mixed culture fermentation kinetics, Greener Journal of Physical Sciences, 3(1): 028-037.
Oubrie, A. (2003) structure and mechanism of soluble glucose dehydrogenase and other PQQ-dependent enzymes, Biochimica and. Biophysica. Acta, 1647: 143–151
Oubrie, A., and Dijkstra, B. W. (2000), Structural rearrangement of Pyroloquinoline quinone enzymatic reactions, Protein Science, 9, 1265–1273
Oubrie, A., Rozeboom, H. J., Kalk, K. H., Huizinga, E. G., and Dijkstra, B. W. (2002) Crystal structure of quinoheamoprotein alcohol dehydrogenase from Comanomas testosterone: structural basis for substrate oxidation and electron transfer, Journal of Biolocial Chemistry,
277: 3727–3732
Oubrie, A., Rozeboom, H. J., Kalk, K. H., Olsthoorn, A. J., Duine, J. A., and Dijkstra, B. W.(1999) Structure and mechanism of soluble quinoprotein glucose dehydrogenase, EMBO Journal 18: 5187–5194.
Oyeku, O.M., Adeyemo, F.S., Kupoluyi, F.C., Abdulhadi, T.M. Davies, O.S., Yussuf, I.G.,
Sadiq, A.O. and Olatunji, O.O. (2009) techno-economic packaging of palm wine preservation and bottling technology for entrepreneur, Global Journal of Social Sciences, 8(1): 21-26.
Qazi, G.N., Parshad, R., Verma, V., Chopra, C.L., Buse, R., Trager, M. and Onken, U. (1991). Diketo-gluconate fermentation by Gluconobacter oxydans. Enzyme and Microbial
Technology Journal, 13: 504 507.
91
Qazi, G.N., Sharma, N. and Parshad, R. (1993). Role of dissolved oxygen as a regulator for the
direct oxidation of glucose by Erwinia herbicola and Gluconobacter oxydans. Journal of
Fermentation Bioengineering, 76: 336-339. Ribereu-Gayon, P., Dubordieu, D., Doneche, B. and Lonvaud, A. (2002). Handbook of Enology.
The microbiology of wine and vinifications. Coordinating, Ribereeau-Gayon (ed), John Eiley and Sons Ltd, West Sussex, England.
Romano, P. and Suzzi, G. (1996). Origin and production of acetoin during wine yeast
fermentation. Applied Environnental Microbiology, 62: 309-315. Ronald, M.A. (2010). Handbook of Microbiological Media (4th ed), CRC Press, Washington, D.C,
pp 23-28
Ruiz, A., Poblet, M., Mas, A. and Guillamon, J.M. (2000). Identification of acetic acid bacteria by RFLP of PCR-amplified 16S rDNA and 16S-23S rDNA intergenic spacer. International Journal of Systematic Evolutionary Microbiology, 50: 1981-1987.
Saeki, A., Teeragool, G., Matsushita, K., Toyama, H., Lotong, N. and Adachi, O. (1997).
Development of thermotolerant acetic acid bacteria useful for vinegar fermentation at higher temperatures. Bioscience, Biotechnology and Biochemistry, 61: 138-145.
Sato, A., Takagi, K., Kano, K., Kato, N., Duine, J. A., and Ikeda, T. (2001) Ca(2+) stabilizes the semiquinone radical of pyrroloquinoline quinone. Biochemical. Journal. 357: 893 898
Satoh, A., Kim, J.K., Miyahara, I., Devreese, B., Vandenberghe, I., Hacisalihoglu, A., Okajima, T., Kuroda, S., Adachi, O., Duine, J.A., van Beeumen, J., Tanizawa, K. and Hirotsu, K. (2002) Crystal structure of quinohemoprotein amine dehydrogenase from Pseudomonas
putida. Identification of a novel quinone cofactor encaged by multiple thioether cross-bridges. Journal Biological Chemistry: 277: 2830–2834
Schobert, M. and Gorisch, H. (1999) Cytochrome c550 is an essential component of the
quinoprotein ethanol oxidation system in Pseudomonas aeruginosa: cloning and sequencing of the genes encoding cytochrome c550 and an adjacent acetaldehyde dehydrogenase. Microbiology, 145: 471–481
Shorter Oxford English dictionary: 6th edition. United Kingdom: Oxford University Press. 2007
Sievers, M., Lorenzo, A., Gianotti, S., Boesch, C. and Teuber, M. (1996). 16-23S ribosomal RNA spacer regions of Acetobacter europaeus and A. xylinum, tRNA genes and antitermination sequences. FEMS Microbiology Letter, 142: 43-48.
Skerman VBD, McGowan V, Sneath PHA (1980). Approved lists of bacterial names.
International. Journal of Systematic Bacteriology, 30: 225-420.
92
Sofer, W., Martin, P.F. (1987). Analysis of alcohol dehydrogenase gene expression in Drosophila, Annual Review of Genetics 21: 203–25.
Sokollek, S.J., Hertel, C. and Hammes, W.P. (1998). Description of Acetobacter oboediens sp. nov. and Acetobacter pomorum sp. nov., two new species isolated from industrial vinegar fermentation. International Journal of Systematic Bacteriology, 48: 935-940.
Sponholz, W.R. and Dittrich, H.H. (1985). Origin of gluconic, 2- and 5- oxo-gluconic,
glucoronic and galactouronic acids in must and wines. Vitis Journals, 24: 41-58. Sugisawa, T. and Hoshino, T. (2001) Purification and properties of membrane-bound D-sorbitol
dehydrogenase from Gluconobacter suboxydans IFO 3255. Bioscience, Biotechnology
and Biochemistry, 65, In Press Sultatos LG, Pastino GM, Rosenfeld CA, Flynn EJ (March 2004). Incorporation of the genetic
control of alcohol dehydrogenase into a physiologically based pharmacokinetic model for ethanol in humans". Toxicological Sciences : an Official Journal of the Society of
Toxicology 78 (1): 20–31. Swings, J. and De Ley, J. (1981). The genera Acetobacter and Gluconobacter. In: The
Prokaryotes, Starr M.P. (ed). Springer-Verlag, Berlin, Germany pp 771-778. Tagaki, K., Torimura, M., Kawaguchi, K., Kano, K. and Ikeda, T. (1999) Biochemical and
electrochemical characterization of quinohemoprotein amine dehydrogenase from Paracoccus denitrificans. Biochemistry, 38: 6935–6942
Theorell, H, and McKee, JS (1961). Mechanism of action of liver alcohol dehydrogenase". Nature 192 (4797): 47–50.
Thompson, C., Fernandes, C., De Souza, O., De Freitas, L. and Salzano, F. (2010). Evaluation of the impact of functional diversification on Poaceae, Brassicaceae, Fabaceae, and Pinaceae alcohol dehydrogenase enzymes". Journal of molecular modeling, 16 (5): 919–928.
Toyama, H., Fujii, K., Matsushita, K., Shinagawa, E., Ameyama, M. and Adachi, O. (1995) Three distinct quinoprotein alcohol dehydrogenases are expressed when Pseudomonas
putida is grown on different alcohols. Journal of Bacteriology, 177: 2442–2450 Trcek, J and Teuber, M. (2002). Genetic restriction analysis of the 16S-23S rDNA internal
transcribed spacer regions of the acetic acid bacter. FEMS Microbiol Letter, 19: 69-75.
Truniger, V. and Boos, W. (1994). Mapping and cloning of gldA, the structural gene of the Escherichia coli glycerol dehydrogenase, Journal of Bacteriology,. 176 (6): 1796–800.
93
Urakami, T., Tamaoka, J., Suzuki, K and Komogata, K. (1989). Acidomonas gen. nov., incorporating Acetobacter methanolicus as Acidomonas methanolica comb. nov. International Journal of Sistematic Bacteriology, 39: 50-55.
Valero, E., Roldan, P., Jimenez, C., Garcia, I. and Mauricio, J.C. (2003). Contenido en
aminoacidos libres en vinagres procedentes de diffentes sustratos. In: Primeras jornadas de I+D+I en la elaboracion de vinagre de vino. Mas, A. and Guillamon, J.M. (eds), Servei de publicacions, Tarragon, pp 53-58.
Van Kleef, M. A. G. and Duine, J. A.(1988). L-tyrosine is the precursor of PQQ biosynthesis in Hyphomicrobium X. FEBS Letter, 237:91–97 . .
Velizarov, S. and Beschkov, V. (1994). Production of free gluconic acid by cells of Gluconobacter oxydans. Biotechnol Letter, 16: 715-720.
Walter, K.A., Bennett, G.N,, Papoutsakis, E.T. (1992). "Molecular characterization of two Clostridium acetobutylicum ATCC 824 butanol dehydrogenase isozyme genes". J.
Bacteriol. 174 (22): 7149–58.
Watanabe, M. and Iino, S. (1984). Studies on bacteria isolated from Japanese wines. In: growth of the Acetobacter sp. A-1 During the fermentation and the storage of grape must and red wine. Part 2. Yamanashien, Dokuhin. Koyo, Shidojo. Kenkyu. HOkoku. 16: 13-22.
Weenk, G., Olijve, W. and Harder, W. (1984). Ketogluconate formation by Gluconobacter
species. Applied Microbiology and Biotechnology, 20: 400-405. White, S., Boyd, G., Mathews, F.S., Xia, Z.X., Dai, W.W., Zhang, Y.F. and Davidson, V.L.
(1993) The active site structure of the calcium-containing quinoprotein methanol dehydrogenase. Biochemistry 32: 12955–12958
Williamson VM, Paquin CE (September 1987). "Homology of Saccharomyces cerevisiae ADH4 to an iron-activated alcohol dehydrogenase from Zymomonas mobilis". Molecular
Genetics and Genonomics, 209 (2): 374–81..
Xia, Z., Dai, W.W., Zhang, Y., White, S.A., Boyd, G.D. and Mathews, F.S. (1996) Determination of the gene sequence and the three-dimensional structure at 2 ångstrom resolution of methanol dehydrogenae from Methylophilus W3A1. Journal of Molecular
Biology, 259: 480–501. Yamada, Y. (2000). Transfer of Acetobacter oboediens Sokollek et al.1998 and Acetobacter
intermedius Boesch et al. 1998 to the genus Gluconacetobacter as Gluconacetobacter oboediens comb. nov. and Gluconacetobacter intermedus comb. nov. International
Journal of Systematic Evolutionary Microbiology, 50: 2225-2227.
94
Yamada, Y., Hoshino, K. and Ishikawa, T. (1997). The phylogeny of acetic acid bacteria based on the partial sequences of 16S ribosomal RNA. The elevation of the subgenus Gluconobacter to the generic level. Bioscience Biotechnology and Biochemistry, 61: 1244-1251.
Yoshida, H., Kojima. K., Witarto, A.B. and Sode, K. (1999) Engineering a chimeric
pyrroloquinoline quinone glucose dehydrogenase: improvement of EDTA tolerance, thermal stability and substrate specificity. Protein Engineering, 12: 63–70.
Yukuphan, P., Potachaoren, W., Tanasupawat, S., Tantichaoren, M. and Yamada, Y. (2004).
Asaia krungthepensis sp. nov., an acetic acid bacterium in the alpha-proteobacteria. International Journal of Systematic Evolutionary Microbiology, 54: 313-316.
Zheng, Y.J. and Bruice, T. C. (1997) Conformation of coenzyme pyrroloquinoline quinone and role of Ca2+ in the catalytic mechanism of quinoprotein methanol dehydrogenase Proceedings of. National Academy of Science. U. S. A. 94: 11881–11886
Zheng, Y.J., Xia, Z.X., Chen, Z.W. and Mathews, F.S. (2001). Catalytic mechanism of quinoprotein methanol dehydrogenase: A theoretical and x-ray crystallographic investigation, Proceedings of. National Academy of Science. U. S. A 98: 432–434
95
APPENDIX I
PREPARATION OF THE COMPONENT REAGENTS FOR PROTEIN
CONCENTRATION
Solution A: an alkaline sodium carbonate solution was prepared by dissolving 2g of the salt and
0.1M NaOH (i.e 0.4g of sodium hydroxide pellets were dissolved in 100ml of
distilled water).
Solution B: a copper tetraoxosulphate (vi)-sodium potassium tartarate solution was prepared by
dissolving 0.5g of CUSO4 in 1g of sodium potassium tartarate, all in 100ml of
distilled water. It was freshly prepared by mixing the stock solution.
Solution C: Folin-Ciocateau reagent was made by diluting the commercial reagent with water in
a ratio 1:1.
Solution D: Standard protein (bovine serum albumin) solution.
Solution E: This was obtained by mixing 50ml of solution A and 1ml of solution B.
96
APPENDIX II
PROTEIN STANDAR CURVE
Protein standard curve
97
APPENDIX III
CALCULATION OF ENZYME ACTIVITY
Activity = ktle
ODblankODtest
××
− * 106 (µmole/min)
ODtest = optical density of the test sample
ODblank = optical density of the blank
e = molar extinction coefficient of potassium ferrocyanide at 660nm = 2000M-1
l = path length =1cm
t = time of incubation = 20mins
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