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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. M