Biochemical genetics of hydrogen metabolism in Escherichia coli

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Biochemical genetics of hydrogen metabolism in Escherichia coli purification and characterization of hydrogenase
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Patel, Pramathesh S., 1957-
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Escherichia coli   ( lcsh )
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Thesis (Ph. D.)--University of Florida, 1985.
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Includes bibliographical references (leaves 118-129).
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by Pramathesh S. Patel.
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Typescript.
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Vita.

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BIOCHEMICAL GENETICS OF HYDROGEN METABOLISM IN
ESCHERICHIA COLI: PURIFICATION AND CHARACTERIZATION OF
HYDROGENASE















By



Pramathesh S. Patel


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF
THE UNIVERSITY OF FLORIDA
IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR
THE DEGREE OF DOCTOR OF PHILOSOPHY




UNIVERSITY OF FLORIDA


1985





















ACKNOWLEDGEMENTS


The author wishes to acknowledge, with, gratitude,

the help and guidance of his major professor, Dr. K. T.

Shanmugam. The author would also like to thank the many

fellow students he had the opportunity to interact

with over the course of these studies.

The author would especially like to thank Dr. R.

P. Boyce, Dr. D. E. Duggan, Dr. P. J. Laipis and Dr. P.

H. Smith for their help, advice, support and

encouragement while serving on the authors advisory

committee.

Special thanks are due to Dr. S. G. Zam for his

constant help, advice and encouragement.

















TABLE OF CONTENTS


ACKNOWLEDGEMENTS.................................... ii

LIST OF ABBREVIATIONS............................... v

ABSTRACT ............................................ vi

INTRODUCTION ..........................................

LITERATURE REVIEW....................................7

Physiological Role of Hydrogenase...............8
Methods to Monitor Hydrogenase Activity........ 11
Properties of the Enzyme ...................... 13
Hydrogenase from Escherichia coli.............. 17

MATERIALS AND METHODS...............................22

Bacterial Strains and Culture Conditions.......22
Chemicals...................................... 22
Media ..........................................22
Enzyme Assays.................................. 24
Polyacrylamide Gel Electrophoresis.............26
Molecular Weight Determination................ 27
Iron and Acid Labile Sulfide Determination.....29
Protein Determination ......................... 30
Removal of Triton X-100........................30
Temperature Profile............................30
pH Profile..................................... 31
Effect of Oxygen on Purified Hydrogenase.......31
Regulation of Hydrogenase......................31

RESULTS ............................................. 34

Purification of Hydrogenase....................34
Molecular Weight Determination .................41
Iron and Sulfur Content ........................65
Temperature Profile............................65
pH Profile..................................... 76
Kinetic Characteristics........................76


iii
















Hydrogen Uptake in the Presence of
Different Artificial Electron Acceptors.....90
Inactivation of Hydrogenase by Oxygen..........90
Stability of Hydrogenase at Alkaline pH........92
Hydrogenase Activity in Solublized
Membranes of HUP Mutants of E. coli........99
Regulation of Hydrogenase Activity
in Whole cells............................. 102

DISCUSSION......................................... 111

REFERENCES ......................................... 113

BIOGRAPHICAL SKETCH............................... 130




















LIST OF ABBREVIATIONS


BV...................................... Benzyl viologen

DEAE ........................Diethylaminoethyl cellulose

EDTA....................Ethylenediaminetetraacetic acid

FHL...............................Formate hydrogenlyase

HUP..................................... Hydrogen uptake

MV.................................... Methyl viologen

PAGE.................Polyacrylamide gel electrophoresis

PEG..................................Polyethylene glycol

SDS..............................Sodium dodecyl sulfate

















v



















Abstract of Dissertation Presented to the Graduate School of The
University of Florida in Partial fulfillment of the Requirements
for the Degree of Doctor of Philosophy


Biochemical Genetics of Hydrogen Metabolism in
Escherichia coli: Purification and Characterization of
Hydrogenase

By

Pramathesh S. Patel

December 1985

Chairman: Dr. K. T. Shanmugam
Major Department: Microbiology and Cell Science



A procedure is described for the purification of

membrane-bound hydrogenase from Escherichia coli. This procedure

uses a non-ionic detergent, precipitating agents, and a series of

column chromatography steps to purify the protein to homogeneity.

The molecular weight of the protein is determined in the presence

and absence of detergent by gel filtration and polyacrylamide gel

electrophoresis, under non-denaturing conditions. These results

show that the enzyme exists predominantly as a monomer (59,000 d)

in the presence of Triton X-100 and as a dimer (124,000 d) in its

absence. The subunit molecular weight is 56,000. The enzyme has












4.4 molecules of iron and 4.7 molecules of acid-labile sulfur per

59,000 d protein. The optimum temperature for catalysis of the

exchange reaction is 35 0C and has a broad pH optimum between pH

values of 7.0 and 7.5. The enzyme has an apparent Km of 26.7 mM

for oxidized methyl viologen, 7.7 mM for oxidized benzyl viologen,

1.5 mM for reduced methyl viologen, 4.0 mM for reduced benzyl viologen

and 1.6 mM for hydrogen. The dimer is twice as active as the monomer.

The enzyme is inactivated by air with a half life of 650 minutes.

Comparison of the biochemical properties of the pure hydrogenase

with the hydrogenase produced by the HUP mutants suggests the pure

hydrogenase is involved in the hydrogen uptake reaction in E. coli.

Study of the regulation of hydrogenase activity in whole cells

indicates that the enzyme activity is inducible under anaerobic

conditions and maximal levels of hydrogenase activity can be detected

within 60 minutes after anaerobic shift. Hydrogenase activity

declined rapidly upon the addition of electron acceptors, viz.,

oxygen and nitrate. The rate of decline of hydrogenase activity due

to the addition of nitrate was bi-phasic with an initial half life

of 40 minutes and eighty minutes after addition of nitrate, the

hydrogenase activity declined with a half life of 10 minutes. This

rapid decline in activity was accompanied by the appearance of nitrite

in the medium. The half life of hydrogenase activity of the culture

exposed to oxygen is 8 minutes.















INTRODUCTION


Since the dawn of civilization, mankind has been in search of an

efficient source of energy. It has been shown throughout history

that there is a direct correlation between the energy consumption per

capital and the standard of living.

Today, the world's population is increasing at an alarming rate.

Most of the increase in the population is taking place in the

impoverished nations of the world. As this major part of the world's

population strives to improve their standard of living, there will be

an unprecedented demand on the energy resources of the world. This

increase in demand will have far-reaching consequences on the life of

every living being on this planet.

As we have already seen, the increased demand for energy in the

last couple of decades has sent the price of fossil fuels on a sharp

increase. As a result of this, the developing countries are caught

in a "no win" situation. To become industrialized, a nation has to

have an abundance of energy at its disposal, and to be able to import

the energy it needs for industrialization, it has to have the

economic resources. Also, the increased demand in energy, world

wide, has led to some of the major political crises around the globe.














Political and economic consequences of the increase in energy

demand are not the only concerns that we have to address. Due to the

increase in the energy demand, one of the questions that we have to

ask ourselves is the effect the increase in energy consumption will

have on the environment. Most of our energy demand today is met by

fossil fuel. It is apparent that the fossil fuel reserves will be

eventually depleted, and in any event, the natural environment cannot

readily assimilate the by-products of fossil fuel consumption at much

higher rates than it does without suffering unacceptable levels of

environmental decay. Thus, today more than ever before, we realize

the importance of a fuel source that is economical and efficient, has

no toxic end products, and can be recycled.

With this bleak outlook for the future in the field of energy, a

lot of money and effort have been spent in exploring alternative

sources of energy that are efficient and non-polluting. At present,

the two most promising areas seem to be solar energy and biomass

conversion. Both these fields have the advantage of being present in

nature in abundance.

One of the products that can be obtained from both biomass

conversion and solar energy is hydrogen. Hydrogen as an energy

source of the future has enormous potential. It has the advantage of

being a very clean fuel. Unlike the fossil fuels of today,

combustion of hydrogen does not yield any of the toxic waste

products, viz., SO2, CO, NOx etc. Water is the main product and the













water produced is easily recycled in nature. The biggest advantage

of using hydrogen as a fuel source of the future is the fact that it

can be produced on a large scale using biological systems. Hydrogen

is produced by most procaryotic microorganisms (5,119). We can

exploit these biological systems and utilize the enormous pool of

energy-rich compounds such as lignin, cellulose and cellobiose found

in nature. Hydrogen is also produced by cyanobacteria and

photosynthetic bacteria (115). During photosynthesis, light energy

is used to generate energy-rich compounds such as NADH, NADPH and

ATP. The ATP, NADH and NADPH can be used to reduce protons resulting

in the evolution of hydrogen. Thus, it is conceivable that some day

in the future hydrogen can be produced on a large scale by harnessing

solar energy.

However, if we are to exploit hydrogen as the energy of the

future we should have a detailed understanding of the role it plays

in nature. Biologically, hydrogen is produced by most procaryotic

organisms growing in the absence of inorganic electron acceptors such

as oxygen or nitrate (40,121). During fermentation, many bacteria

use protons as an electron sink, resulting in the evolution of

hydrogen. Hydrogen is also consumed by many bacteria as a source of

reducing power under appropriate conditions (5,119). During the

formation of methane, a widely used fuel, methanogens utilize

hydrogen as a source of reductant to reduce CO2. If we understand














the role of hydrogen gas in these reactions to a greater extent, we

will be able to exploit it for our own good.

At the center of the hydrogen metabolism in biological systems

lies the enzyme hydrogenase. It catalyzes the primary reaction in

hydrogen metabolism, the reversible activation of a hydrogen

molecule. Thus it is imperative that we understand the mechanism of

catalysis of this basic reaction and its regulation in nature.

The best way to approach the basic questions regarding the

mechanism of catalysis and its optimal conditions for catalysis is to

study the enzyme after it has been obtained in a pure state. The

purified enzyme can also be used to raise antibodies in a mammal.

The antibodies can then be used to elucidate the regulation of

hydrogenase synthesis, identify the structural gene for the enzyme

and also determine the location of hydrogenase in the membrane with

respect to other proteins.

Since E. coli possesses hydrogenase it seems logical to examine

the role it plays in the cells metabolism. E. coli offers the

following advantages: 1) It is a very well studied organism with

respect to metabolism; 2) Has established techniques to investigate

the biochemistry of the various metabolic reactions and for genetic

manipulation of the organism; 3) The ease with which a large number

of mutants can be isolated and characterized. A thorough

understanding of hydrogen metabolism in one organism may help us

better understand the hydrogen metabolism of other micro-organisms.














In Escherichia coli, hydrogenase is known to be a part of the

format hydrogenlyase enzyme complex which is responsible for the

evolution of hydrogen gas from format under anaerobic growth

conditions. Hydrogenase is also involved in the growth of E. coli

under conditions where hydrogen is the source of reductant, in the

presence of a suitable electron acceptor, viz., fumarate. It is not

known whether the same enzyme is involved in both the hydrogen

evolution and uptake reactions, or whether there are two separate

hydrogenases responsible for the two different reactions.

There are a number of reports in the literature concerning

hydrogen metabolism in E. coli. Most of the studies have been

directed toward characterization of the enzyme hydrogenase, and

toward determining the number of hydrogenase proteins in a cell

(details are presented in the Literature Review section). However,

the number of hydrogenases present in the cell has not yet been

established conclusively. Even the reports describing the physical

properties of the enzyme provide conflicting molecular weights for

the enzyme. Adams and Hall (4) reported a molecular weight of

113,000 for the hydrogenase they purified from E. coli. In 1978,

Bernhard and Gottschalk (12), reported a molecular weight of 191,000

for hydrogenase from E. coli. The molecular weight in the latter

report was determined by the sucrose density gradient centrifugation

and gel filtration of a partially purified preparation. On the

other hand, Graham et al. (37), in 1980, reported a molecular weight













of 56,000 for hydrogenase from E. coli. They determined the

molecular weight by an indirect method which involved electrophoresis

of solubilized membrane vesicles in native PAGE, staining for

hydrogenase activity followed by SDS-PAGE of the excised active band

from the gel. It should also be stated that both, Adams and Hall

(4), and Bernard and Gottschalk (12), used proteases to aid in the

solubilization of the enzyme from the membrane. It is possible that

the enzyme obtained after protease treatment may not reflect the

properties of the native enzyme.

This work describes a scheme for purification of the enzyme

hydrogenase to homogeneity without the use of proteolytic agents.

The enzyme is characterized with respect to its biochemical and

physical properties. The regulation of the enzyme in cultures

growing in the presence or absence of electron acceptors such as

oxygen and nitrate is also studied.














LITERATURE REVIEW


Production and consumption of hydrogen gas by microorganisms has

been known since the turn of the century (44). However, it was only

in the 1930's that the enzyme responsible for the evolution of

hydrogen gas during bacterial fermentation was identified and the

physiology of this process studied (24,41,98,113). During an

investigation of a pollution problem in the Great Ouse river,

Stephenson and Stickland (98) observed that the microorganisms

thriving upon the sugar beet waste dumped into the sluggish river

were responsible for the evolution of gases such as hydrogen, carbon

dioxide and methane. They also showed that washed cultures of

bacterial isolates from the river bed were able to reduce methylene

blue in the presence of hydrogen. This observation led them to

conclude that the microorganisms possesed an enzyme capable of

activating a molecule of hydrogen. This enzyme, which they termed

hydrogenase (EC 1.12), catalyses the reversible reaction as

represented by the equation



H2 + e carrier oxidisedd) 2 H+ + e carrier (reduced)













Since the initial discovery of hydrogenase, the enzyme has been

shown to be present in a diverse group of microorganisms, both

bacteria and algae (5,40,119). In vivo, hydrogenase is usually

coupled to other electron carriers or is a part of a multienzymne

complex, and thus, the enzyme generally catalyzes an "irreversible"

reaction, in vivo. However, in the presence of suitable electron

donors or acceptors all pure hydrogenase proteins examined so far,

catalyze reversible reactions.



Physiological Role of Hydrogenase


The physiological role of hydrogenase in the anaerobic

metabolism of microorganisms can be divided into two categories : 1)

evolution of hydrogen and 2) consumption of hydrogen.

In the fermentative bacteria, evolution of hydrogen via

hydrogenase can be seen as a means of oxidizing the electron carriers

reduced during fermentation. This oxidation is required to allow the

electron carriers to recycle so that a continuous supply of ATP can

be generated by substrate level phosphorylation. For example, in

Clostridium pasteurianum, each mole of glucose yields two moles of

pyruvate which is further degraded to acetyl-CoA, CO2 and hydrogen,

via pyruvate:ferredoxin oxidoreductase and hydrogenase (104,105,106).

Some of the excess NADH generated at the level of

glyceraldehyde-3-phosphate dehydrogenase is oxidised to produce













hydrogen by NADH:ferredoxin oxidoreductase and hydrogenase (5,27,59).

Thus, the redox balance is maintained in the cell without the need

for terminal electron acceptors other than protons. In E. coli

pyruvate is metabolized via the pyruvate-formatelyase enzyme complex

to format and acetyl Co-A (56). The format is further metabolized

to CO2 and H2 via the format hydrogenlyase complex of which

hydrogenase is an integral part (5,41,42,59).

Under conditions where hydrogen is the only source of reducing

power and energy, hydrogenase oxidizes hydrogen. The electrons

obtained are utilized to reduce inorganic or organic electron

acceptors (5,42,69,76,110). For example, the genus Desulfovibrio
-2
possesses the ability to use SO4-2 as a terminal electron acceptor.

Electrons obtained from the oxidation of hydrogen are utilized to

reduce SO 4-2 and yield S-2 (6,42). Paracoccus denitrificans has the

ability to utilize nitrate as the terminal electron acceptor for the

electrons obtained from the oxidation of hydrogen by hydrogenase

(96). In methanogens, hydrogen can serve as the sole electron donor

for the reduction of CO2 to methane (12,120). In E. coli, under

appropriate conditions, hydrogen can serve as an electron donor to

reduce electron acceptors, such as fumarate, nitrate, and oxygen

(5,36,69). In all of these reactions, hydrogenase is an essential

enzyme and is associated with the membrane and an electron transport

chain (69). Therefore, the oxidation of hydrogen not only serves as













a source of electrons but the association of hydrogenase with the

membrane helps generate a proton gradient across the membrane.

Hydrogenase also plays a dual role in photosynthetic bacteria.

During photosynthesis, the bacteria utilize hydrogen as a source of

reductant for CO2 fixation (5,115). This ability to utilize hydrogen

as a source of reductant is mediated by hydrogenase (53). In

addition, during dark fermentation, the purple non-sulphur bacteria

metabolize pyruvate via a pyruvate formatelyase and a format

hydrogenlyase system analogous to that found in E. coli (33).

Another group of microorganisms that possess hydrogenase is the

aerobic hydrogen oxidizing bacteria. These microorganisms are

characterized by their ability to grow autotrophically, using

hydrogen as the sole electron donor, CO2 as the carbon source and

oxygen as the terminal oxidant (35,39). Some of the microorganisms

belonging to this group have two different types of hydrogenases.

One of the hydrogenases is soluble in the cytoplasm and reduces NAD

directly with hydrogen (82). The other hydrogenase is membrane bound

and donates electrons to the respiratory chain which in turn reduces

oxygen and thus produces energy for autotrophic growth (34,91,93).

From the examples cited above, it can be seen that hydrogenase

plays an important role in the physiology and metabolism of a diverse

group of microorganisms.













Methods to Monitor Hydrogenase Activity


In 1930, the classical hydrogenase assay involved the reduction

of methylene blue in the presence of hydrogen (98). Today,

hydrogenase activity can be monitored by three different methods: 1)

hydrogen evolution, 2) hydrogen consumption and 3) the exchange

reaction (59).

In the hydrogen evolution reaction, the ability of hydrogenase

to reduce protons in the presence of suitable electron donors is

monitored. Electron donors such as reduced ferredoxin (12,73,77),

cytochrome c3 (77), and reduced viologen dyes (38,58,66,80,102) have

been employed. The rate of hydrogen evolution is monitored either

manometrically or by using a gas chromatograph or a hydrogen

electrode (58,59,80,112).

Conversely, in the hydrogen uptake reaction the ability of

hydrogenase to oxidize hydrogen to protons and electrons is

monitored. The oxidation of hydrogen requires the presence of

suitable electron acceptors. In the presence of such electron

acceptors, the rate of the reaction can be monitored either

spectrophotometrically, or by using a gas chromatograph or a hydrogen

electrode (5,59,100).

In the exchange reaction, hydrogenase activity can be measured

by monitoring the exchange between molecular hydrogen and heavy water

or between tritium gas and water (22,23,58,59,111).













Farkas et al. (24) were the first to demonstrate an exchange reaction

between hydrogen gas and heavy water catalysed by E. coli. The

reaction as represented by the equation



HD + H20 HDO + H2



is catalysed by hydrogenase. The rate of the forward reaction can be

measured by monitoring the appearance of the isotope in the aqueous

phase. In 1963 Gingras et al.(28) introduced a modification of the

exchange assay in which tritium gas was used in place of hydrogen and

heavy water was replaced by water. In this modified assay as

represented by the equation



HT + H20 HTO + H2



hydrogenase activity can be followed by monitoring the accumulation

of tritiated water in the aqueous phase.

If the hydrogen evolution or hydrogen consumption reaction is

used to determine hydrogenase activity, one has to bear in mind that

the rate of the reaction may not be a true reflection of hydrogenase

activity. Since hydrogenase is usually associated with other

electron transport proteins in the cell, it is possible that the

activity of hydrogenase in whole cells or intact membranes monitored

using artificial electron acceptors or donors may reflect the













activity of a multienzyme complex, of which hydrogenase is one of the

many components. Gitilitz and Krasna (29) did observe this

phenomenon. They found that the activity of hydrogenase from

Chromatium, measured by reduction of artificial electron carriers

decreased during purification as compared to the exchange activity,

indicating the presence of various cellular electron carriers in the

crude extract that enhanced oxidation/reduction of artificial

electron carriers. Also, since the reaction catalysed by hydrogenase

is an oxidation/reduction reaction, the redox potential (Eo') of the

substrate determines the rate of the reaction. The exchange

reaction, on the contrary, is a direct and simple assay for

hydrogenase and it is extremely sensitive. Considering these facts,

it appears that the exchange reaction is the method of choice to

assay for hydrogenase activity.



Properties of the Enzyme


Purification of any protein is immensely simplified if some of

the physical properties and special characteristics of the protein

are known. Usually, information on the protein from other systems

are used as indicators for "do's and don'tts. There are a number of

reports, in the literature, describing the purification of

hydrogenase from a diverse group of microorganisms

(3,4,12,16,19,29,31,38,51,70,90,92,94,107,116). Review of the














literature indicates that the enzyme hydrogenase is inactive in the

presence of oxygen. Thus, it is important to protect the enzyme from

oxygen during the purification procedure. Another factor to be taken

into consideration is the location of the enzyme. Hydrogenase in the

microbial world is found either in the periplasmic space, membrane

associated or as a soluble cytoplasmic enzyme. If the hydrogenase is

membrane associated, as in the case of E. coli (4,35), one has to

incorporate special procedures during the purification procedure.

Membrane proteins have been solubilized using various detergents

(32). Hydrogenase has been successfully solubilized from the

membrane using detergents, viz., Triton X-100 (8,35,63,71,95,97), and

sodium deoxycholate (4,29,71,83). Some researchers have used

proteolytic agents, such as trypsin or pancreatin to aid in the

solublization of hydrogenase from the membrane (2,4,94,117).

However, one has to realize that the use of proteases during the

purification procedure may yield a protein that is altered due to

proteolytic digestion.

The sensitivity of the enzyme to inactivation by oxygen has been

overcome by deoxygenating the buffers with pre-purified nitrogen or

argon, adding 1mM sodium dithionite to scavenge any contaminating

oxygen and carrying out all procedures in closed vessels under

positive pressure of nitrogen, argon or hydrogen

(18,43,45,55,75,109).













Most of the other techniques used are the standard techniques

used for protein purification, viz., ion exchange chromatography and

gel filtration (48,49). In addition to these established procedures

many researchers have taken advantage of the hydrophobic nature of

the membrane protein and included chromatography on matrices such as

Octyl- or Phenyl-Sepharose (46,49,94,108).


Physical Properties of Hydrogenases: Hydrogenase is one of the few

anaerobic proteins that has attracted considerable attention in the

recent past. One of the first hydrogenases to be purified to

homogeneity was from Clostridium pasteurianum (17) and is

a well characterized enzyme. Since the time the first hydrogenase

was purified, the enzyme has been obtained in pure state from a

number of other organisms (5).

All the hydrogenases studied so far can be grouped into three

major groups based on their molecular weight and number of subunits.

Most of the hydrogenases are made up of a single polypeptide with an

apparent molecular weight ranging from 50,000 to 66,000. The second

group of hydrogenases range in molecular weight from 89,000 to

101,000. Hydrogenases belonging to the latter group have two

subunits. The large subunit has a molecular weight of about 62,000

to 67,000 and the small subunit ranges from 26,000 to 34,000.

Hydrogenases studied from two organisms, namely Paracoccus

denitrificans and Alcaligenes eutrophus, belong to a third group with













a molecular weight of about 205,000 and consists of 4 subunits

(5,92). There are two large subunits and two small subunits. The

molecular weight of the large subunits are 63,000 and 67,000 and the

molecular weight of the small subunits range from 31,000 to 33,000

(5,92).

There is some variability with respect to the enzyme's

sensitivity to oxygen. All of the hydrogenases are inactive in the

presence of oxygen (5). However the inactivation is usually

reversible. In Clostridium pasteurianum, Desulfovibrio gigas and

Alcaligenes eutrophus H16, the enzyme is extremely sensitive to

oxygen and irreversibly inactivated (5).


Presence of Metal Ions in Hydrogenase: Hydrogenase is a non-heme iron

sulfur protein. The hydrogenases characterized so far have been

reported to have varying contents of iron and acid labile sulfur.

Most of the active centers contain 4 atoms of Fe and 4 atoms of acid

labile S (5).

A number of reports indicate the presence of at least two other

metal ions, nickel and copper. Nickel has been shown to be present

in the hydrogenases obtained from Rhodospirillum rubrum (3),

Chromatium vinosum (7), E. coli (10), Rhodopseudomonas capsulata

(20), Methanobacterium thermoautotrophicum (34), Desulfovibrio

vulgaris (Hildenborough) (39), Desulfovibrio desulfuricans (61),

Desulfovibrio gigas (101), and Vibrio succinogenes (103). Copper has














also been shown to be a part of the hydrogenase derived from

Desulfovibrio vulgaris (Hildenborough) (39).



Hydrogenase from Escherichia coli


Hydrogenase from E. coli and its hydrogen metabolism has

attracted considerable amount of attention. As indicated above, E.

coli is capable of metabolizing format to yield hydrogen gas and

also utilizing hydrogen as a source of reductant under certain growth

conditions (5,69,81). However, it is not yet known whether E. coli

has two hydrogenases, one for the format hydrogenlyase (FHL)

reaction and the other for the hydrogen uptake (HUP) reaction, or one

single hydrogenase protein that is involved in both the reactions.

There have been reports in the literature suggesting the presence of

multiple hydrogenases in E. coli. Ackrell et al. (1) reported the

presence of three hydrogenase species in E. coli. Yamamoto and

Ishimoto (118), demonstrated that extracts of E. coli cells grown in

different media, either favoring conditions for hydrogen evolution or

hydrogen uptake, when subjected to electrophoresis in polyacrylamide

gels exhibited bands possessing hydrogenase activity with different

electrophoretic properties. More recently, Ballantine and Boxer (10)

have reported the existence of two distinct hydrogenases in E. coli.

This contention is based on the detection of two immuno-precipitin

arcs possessing hydrogenase activity in extracts of cells grown under














anaerobic conditions. The cross-immuno-electrophoresis was performed

using Triton X-100 dispersed membranes as antigen. The antiserum

from rabbits immunised with E. coli membranes, was used as

antibodies. They also demonstrated three distinct bands of

hydrogenase activity upon subjecting Triton dispersed E. coli

membranes to electrophoresis in non-denaturing polyacrylamide gels.

Two of the bands were not detected when the extract was exposed to

alkaline pH (pH 10.0), suggesting that one of the hydrogenase was

inactivated at this pH. They also found that the hydrogenase

resistant to alkaline pH was not easily solubilized from the

membrane. Preliminary experiments in our laboratory also suggest the

presence of two distinct hydrogenases in E. coli.

Even though the number of hydrogenases in E. coli has not been

determined directly, numerous attempts have been made to characterize

the hydrogenase(s) from E. coli. In 1950, Joklik (50) made the first

attempt to characterize the hydrogenase from E. coli, followed by

Gest (26) in 1952. In 1957, Kondo et al. (57) published a procedure

for the solublization of hydrogenase using 4% deoxycholate. It was

not until 1979 that the first report describing the purification of

hydrogenase in E. coli to a high degree of purity was reported.

Bernhard and Gottschalk (12) published a report on the purification

of the enzyme using a modified method of Kondo et al. (57).

Employing trypsin and deoxycholate, they obtained a preparation of

the enzyme estimated to be 80% pure. The molecular weight as













determined by gel filtration and density gradient centrifugation was

reported to be 191,000. They also reported the enzyme to be

irreversibly inactivated in the presence of oxygen with a half life

of 36 hours.

In 1979, Adams and Hall (4) reported the purification of

hydrogenase from E. coli to homogeneity. Their procedure differed in

that they used aerobically grown cells obtained from a commercial

source and solubilized the enzyme with sodium deoxycholate and

pancreatin. The enzyme, a cytoplasmic membrane-bound protein, is

reported to have a molecular weight of 11 3,000 and consists of a

dimer of identical subunits. The enzyme, an iron sulphur protein,

has 12 Fe and 12 acid labile S atoms per molecule. They reported a

half life for hydrogenase of 12 hours under air at room temperature.

In 1980 Graham et al. (37) determined the molecular weight of

Hydrogenase from E. coli to be 53,000. The molecular weight was

determined by subjecting Triton X-100 solubilized E. coli membranes

to Native PAGE and staining the gel for hydrogenase activity. The

band possessing hydrogenase activity was then cut from the gel and

the protein was eluted and subjected to SDS-PAGE in cylindrical gels.

In 1981, however, Graham (35) reported a molecular weight of 63,000.

In the latter case the molecular weight was determined using

polyacrylamide gel electrophoresis in slab gels. He also reported

the enzyme to be a trans-membranous protein.














Genetic studies thus far have not yet identified the structural

gene(s) of hydrogenase in E. coli. Pascal and her co-workers (78)

reported the isolation of a hydrogenase activity-deficient mutant.

However, the mutant also lacked format dehydrogenase activity,

indicating that these mutants could be defective in the format

hydrogenlyase enzyme complex. The mutant strains described by Graham

and his co-workers (37) and Krasna (60) also fall into the same

category. Glick and his co-workers (30) reported the isolation of a

hydrogenase defective mutant, however, analysis of the mutant in our

laboratory has indicated that the mutant does possess hydrogenase

activity. In 1983, Bock and his co-workers (79) used Mudl(Ap,-lac)

insertion mutagenesis to obtain mutants defective in hydrogenase.

They were successful in isolating a mutant that lacked hydrogenase

activity. Using beta-galactosidase activity as a means of monitoring

regulation of synthesis, they showed that the gene affected by the

Mud insertion was synthesized only under anaerobic conditions and in

the absence of electron acceptors such as nitrate. In 1981, Tait and

his co-workers (99) described the isolation of strains of E. coli

that lacked hydrogenase activity. All of the mutants defective in

hydrogen metabolism are affected only in those genes that are

essential for hydrogenase activity. In 1983 Karube and his

co-workers (52) reported the isolation of a hydrogenase mutant from

E. coli. The mutant strain, which is unable to reduce methyl














viologen as tested by the filter-dye reduction method, was isolated

after mutagenesis with N-methyl N'-nitro-N-nitrosoguanidine.

Exploiting a positive selection method to isolate mutants of

E. coli defective in its hydrogen metabolism (65), our laboratory has

successfully isolated a large number of such mutants. Based on

phenotypic characteristics, the mutants have been grouped into two

distinct classes. One of these classes, which is defective in

hydrogen uptake, did produce an active format hydrogenlyase, has a

lesion near 65 minutes in the E. coli chromosome (genetic map (9)

and is 76% co-transducible with metC (65). The other class of

mutants which lack hydrogenase activity have lesions between srl and

cys operons (58 and 59 min, respectively) on the chromosome. Based

on fine structure analysis of this region, the latter class of

mutants have been further subdivided into two sub-groups. These two

sub-groups belong to two distinct operons. Genes from both of these

operons are essential to produce an active hydrogenase in the cell.

Segments of DNA from wild type E. coli containing both of these

operons have been cloned (88). Further genetic studies of the region

have shown that there are at least four distinct genes responsible

for the production of an active hydrogenase in the cell (personal

communication, Sankar, P. and Lee, J.H.).















nATERIL.LS AiD IET.IODS


Bacterial Strains and Culture Conditions


Bacterial strains used in this study are listed in Table 3-1.

Bacterial cultures for each experiment were grown as described for

each individual experiment, in the Results Section.



Chemicals


All the chemicals used were of analytical grade and were

obtained from Fisher Scientific Company, Pittsburgh, PA, or Sigma

Chemical Co., St. Louis, MO.



Media


The mineral base for the minimal medium used to cultivate E.coli

had the following composition: (grams/liter) Na2HPO 6.25; KH2PO,

0.75; NaCl, 2.00; (NH )2SO, 1.00; MgSO4.7H20, 0.20; FeSO .7H20 ,

0.010; NaMo0O.2H2O, 0.010; Na2Se03, 0.000263. The pH of the medium

was 7.5. The carbon source was glucose and was supplied at a

concentration as described for particular experiments.
















Bacterial strains used in this study.


Strain Genotype or Phenotype Source



K-10 Hfr PO2A relAl pit-10 L. Csonka

tonA22 T2r + spoT


JC10244 cysC43 alaS3 srl-300::Tn10 thrl L. Csonka
leu-6 thi-1 lacY1 galK2 ara-14
xyl-5 mtl-1 proA2 his-4 argE3
rpsL31 tsx-33 supE 4V


SE-8 thi-1 leu-6 suc-10 bioA2(?) galT27 Laboratory
rpsL129 chlC3 A- hup101::Tn10 Collection
(65)


SE-49 Same as JC 10244 but alaS+ Laboratory
recA56 and hup103 Collection
(65)

SD 7 gal-25, A topA10, pyrF287, B. Bachmann
fnr-1, rpsL195, gyrB226, iclR7 CGSG 6335
and trp72


* CGSC, Coli Genetic Stock Center.


Table 3-1.













Luria broth (LB) medium had the following composition:

(grams/liter) Bacto tryptone, 10.00; Bacto yeast extract, 5.00; NaC1,

10.00; The pH of the medium was 7.0. All solid media contained 15

grams of agar per liter of medium.



Enzyme Assays


Hydrogen Uptake: Hydrogen uptake activity was measured at room

temperature using a Spectronic 710 spectrophotometer. The reaction

was carried out in a 12 x 75 mm test tube. A 2.5 ml reaction mixture

contained 2.3 ml of 10 mrM phosphate buffer, pH 7.0, and the electron

acceptor at concentrations described for each experiment. The tubes

were capped with serum stoppers, evacuated and filled with hydrogen

several times. The reaction was started by adding hydrogenase, 0.2

ml, to attain a final protein concentration of 12.0 microgram/ml.

The contents were mixed and the reduction of the electron acceptor

was measured at the appropriate wavelength. The electron acceptors

used and their extinction coefficients were : methyl viologen, 12,000
-1 -1 -1 -1
M cm at 600 nm; benzyl viologen, 7,780 M cm at 550 nm;

neutral red, 7600 M-1 at 450 nm; methylene blue, 7000 M at 601 nm;

phenosafranin, 1150 M- at 400 nm; and potassium ferricyanide 13200

M- at 405 nm (The Merck Index, tenth edition, ed. M. Windholz, Merck

and Co., Rahway, N.J.).













Hydrogen Evolution: The assay to measure hydrogen evolution from

reduced viologen dyes was carried out in 9.0 ml serum vials, at 230C.

Eight hundred and seventy five microliters of 10 rimM phosphate buffer,

pH 7.0, containing either methyl viologen or Benzyl Viologen

concentrations described for each experiment was placed in serum

vials. The vials were capped with serum stoppers, evacuated, and

filled with nitrogen, six times. The reaction was started by adding

75 microliters of hydrogenase, to attain a final protein

concentration of 2.0 microgram/ml. To reduce the viologen dye, 50

microliters of sodium dithionite was added anaerobically, with a

syringe, to attain a final concentration of 50 mM. The final

reaction volume was 1.0 ml. The rate of hydrogen evolution was

monitored using a Varian Model 910 gas chromatograph.


Tritium Exchange : The reaction was carried out in a 12 x 75 mm test

tube. Ninety microlitres of 10 mM phosphate buffer, pH 7.0, was

placed in a tube (12 x 75 mm), and the tube was sealed with a serum

stopper. The 5.1 ml gas phase was replaced with helium by evacuating

the tube and filling it with helium. The procedure was repeated six

times. Hydrogenase was added to a final concentration of 0.25

microgram/ml, and 0.1 ml of sodium dithionite, pH 7.0, to attain a

final concentration of 1.0 mM. Eight hundred microliters of hydrogen

gas was added to each of the assay tubes with a syringe. Tritium gas

(11.2 mCi/mmol; New England Nuclear Corp. Boston, MA.) was added (25














microliters) to a final concentration of 0.55 micro-curie per assay

as a means to monitor the exchange reaction. After 1 hour of

incubation at 370C, the serum stopper was removed and the tritium gas

was vented in a fume hood for 10 min. To 100 microliters of the

assay mixture, 2.5 ml of a water-based scintillant was added.

Tritiated water present in the 100 microliter fraction was determined

with the aid of a scintillation counter. The assay used to monitor

hydrogenase activity during the purification procedure was the same,

except the sample assayed was 100 microliters in a final assay volume

of 200 microliters, tritium gas was added to a final concentration of

0.22 micro-curie per assay and no hydrogen gas was added.



Polyacrylamide Gel Electrophoresis


Polyacrylamide gel electrophoresis under non-denaturing

conditions was performed as described by Davis et al. (21). Sodium

dodecyl sulfate polyacrylamide gel electrophoresis was performed as

described by Laemmli (62). The gels were run as either tube gels, or

slab gels. The dimensions of the tube gels were 0.6 cm x 8.8 cm.

The volume of the separatory gel used was 2.2 ml. The dimensions of

the slab gels were 17 x 14.5 x 0.15 cm. The volume of the separatory

gel was 30 ml. Location of hydrogenase, after electrophoresis in

non-denaturing gels was determined by incubating the gel in 10 mM

phosphate buffer, pH 7.0, containing benzyl viologen at a final














concentration of 0.2% and under an atmosphere of hydrogen. The

reduced benzyl viologen, which is auto-oxidizable, was made to

further react with 2,3,5-triphenyl tetrazolium chloride to produce a

bright red permanent band of reduced formazan. The gels were stained

for protein with either the silver stain method described by

Morrissey (72), or with the aid of coomassie blue R-250 as described

by Wilson (114). The molecular weight standards used for the

determination of the molecular weight of hydrogenase using SDS-PAGE

were obtained from Sigma Chemical Co., St. Louis, MO, and consisted

of alpha-lactalbumin, 14,200; trypsin inhibitor, soybean, 20,100;

trypsinogen, PMSF treated, 24,000; carbonic anhydrase, bovine

erythrocytes, 29,000; glyceraldehyde-3-phosphate dehydrogenase,

rabbit muscle, 36,000; albumin, egg, 45,000; and albumin, bovine,

66,000.



Molecular Weight Determination


Gel Filtration: The Sephadex G-200 used for gel filtration was

swollen by incubating the beads in deionised water (15 g of beads

per liter of water) for 72 hours at room temperature. After 72 hours

a column of 1.8 x 48 cm was packed with the swollen Sephadex G-200.

The column was equilibrated with 10 mM phosphate buffer, pH 7.0,

(Triton X-100 concentration was 0.3%, if present). The column was

maintained at a flow rate of 11 ml/hr at 4C for 16 hours before use.













Blue Dextran (MW = 2,000,000) was used to determine the void volume

(Vo) of the column. The proteins used for generating a molecular

weight calibration curve for the Sephadex G-200 column were obtained

from Sigma Chemical Co. and consisted of cytochrome c, horse heart,

12,400; carbonic anhydrase, bovine erythrocytes, 29,000; albumin,

bovine serum, 66,000; alcohol dehydrogenase, yeast, 150,000 and

beta-amylase, sweet potato, 200,000. The proteins were dissolved in

the equilibration buffer at the following concentrations: albumin, 10

mg/ml; alcohol dehydrogenase, 5 mg/ml; beta-amylase, 4 mg/ml;

carbonic anhydrase, 3 mg/ml and cytochrome c, 2 mg/ml. The sample

volume in all cases was maintained at 1.5 ml. Fractions of 0.93 ml

were collected. The calibration curve for the column was generated

by plotting the log of the molecular weight of the standard proteins

against the ratio of their elution volumes and the void volume for

the column. Hydrogenase was loaded at a final concentration of 2.6

micrograms/ml and its elution profile was determined by assaying the

fractions collected, for tritium exchange activity. The molecular

weight of hydrogenase was determined from the calibration curve.


Native PAGE: To determine the molecular weight of hydrogenase, the

enzyme was subjected to electrophoresis in the presence of various

concentrations of acrylamide (47). Triton X-100, when present, was

incorporated at a final concentration of 0.3%. The electrophoresis

was performed in tube gels prepared as described above. After













polymerization, the tube gels were subjected to electrophoresis for 1

hour at 1 mA/gel. After the initial run, samples were loaded and run

at 1 mA/gel for the first hour and then the current was increased to

2 mA/gel. The sample volume was 100 microliters, in all cases. The

molecular weight standards used were obtained from Sigma Chemical Co.

and consisted of alpha-lactalbumin, bovine serum, 14,200; carbonic

anhydrase, bovine erythrocytes, 29,000; albumin, chicken egg, 45,000;

albumin, bovine serum, 66,000 (monomer); 132,000 dimerr); urease,

jack bean, 240,000 dimerr); 480,000 (tetramer). Hydrogenase bands

were detected by staining the gels for hydrogenase activity after

electrophoresis. The relative migration of the molecular weight

standard proteins was determined by staining the gels for protein

using the coomassie blue method as described by Wilson (114). A

Ferguson plot (25) was generated using the Rf values obtained for the

molecular weight standard proteins for each experiment. The

molecular weight of hydrogenase was obtained from the Ferguson plot

generated for each experiment.



Iron And Acid Labile Sulfide Determination


Total iron was determined by the ortho phenanthroline method as

described by Lovenberg et al. (68). Acid labile Sulfide was

determined by the method of King and Morris (54).













Protein Determination


Protein was determined using Coomassie Blue G-250 as described

by Bradford (15). Albumin, Bovine Serum was used as the standard.



Removal of Triton X-100


Triton X-100 was removed from a sample in two steps. Initially,

the Triton X-100 concentration was lowered by dialysing the protein

sample against a 100-fold excess of an appropriate buffer, at 4C,

for 4 hours. After dialysis the residual amount of Triton X-100 was

removed by incubating the sample with Bio-Beads SM-2 (20% w/v) on a

rocking platform at 4C for 2 hours. The sample was separated from

the Bio-Beads SM-2 by centrifugation at 5,000 rpm for 5 min.at 40C.



Temperature Profile


Tritium exchange assay was used to determine the temperature

profile for hydrogenase activity. The assay was performed as

described earlier, both, in the presence and absence of Triton X-100.

The only difference was the incubation temperature. The reaction

vials were incubated in water baths maintained at 15, 20, 25, 30, 35,

40, 45,and 50 degrees centigrade for one hour.














pH Profile


Tritium exchange assay was used to determine the pH profile for

hydrogenase activity. The assay was performed as described earlier.

However, 10 mM phosphate buffer was replaced by one of the following

buffers present at a final concentration of 100 mM: Tris, pH, 9.5,

9.0, 8.5, 8.0, and 7.5; piperazine-N,N'-bis[2-ethane-sulfonic acid]

(PIPES), pH, 8.0, 7.5, and 7.0; and phosphate, pH, 7.5, 7.0, 6.5, 6.0

and 5.5.



Effect of Oxygen on Purified Hydrogenase


To determine the effect of oxygen on purified hydrogenase, 1.0

ml of the enzyme (at a concentration of 2.6 microgram/ml, in 10 mM

phosphate buffer, pH 7.0) was placed in a 12 x 75 mm tube. The gas

phase in one of the tube was air whereas the gas phase of the control

tube was hydrogen. The tubes were rocked (20 oscillations/min.) on a

rocking table at room temperature. Samples were withdrawn at various

time intervals to determine hydrogenase activity by monitoring the

tritium exchange reaction as mentioned above.



Regulation of Hydrogenase


Induction of Hydrogenase: To study the induction of hydrogenase

activity in E. coli, strain K-10 was used. The culture used as the














inoculum for the experiment was grown aerobically in 10 ml of LB +

1.5% glucose at 370C. The culture was grown to an optical density of

0.170 at 420 nm. At this stage, the culture was used to inoculate 40

ml of LB + 1.5% glucose medium in a 70 ml serum bottle. The inoculum

size was 25% of the final culture volume. The culture bottle was

capped with a serum stopper and flushed with nitrogen. The gas phase

in the culture vessel was replaced with nitrogen and the bottle was

incubated at 370C. Every twenty minutes, samples were withdrawn to

monitor growth, by measuring the optical density at 420 nm in a

spectrophotometer (Bausch and Lomb, Spectronic 710), and the

hydrogenase activity by assaying for tritium exchange activity.


Effect of Oxygen on Hydrogenase Activity in Whole Cells: To study the

effect of oxygen on hydrogenase activity in whole cells, a culture of

E. coli fully induced for hydrogenase was used. E. coli K-10 was

grown overnight in LB +1.5% glucose at 37 C under an atmosphere of

nitrogen. These cells were used to inoculate 40 ml of LB + 1.5%

glucose in two 70 ml serum bottles. The serum bottles were capped

with rubber stoppers and the gas phase replaced with nitrogen. The

medium was warmed to 37 C and inoculated anaerobically with a syringe

with the overnight grown culture. The inoculum size was 1% of the

final culture volume. The culture was incubated at 37 C. After 130

minutes, one of the culture bottles was opened and 35 ml of the

culture was transferred aseptically to a 500 ml flask and incubated













at 370C under aerobic conditions shaking at 200 rpm. Samples were

withdrawn at various time intervals to monitor growth by measuring

the optical density at 420 nm and monitoring the hydrogenase activity

by assaying for the tritium exchange reaction.


Effect of Nitrate on Hydrogenase Activity in Whole Cells: To study

the effect of nitrate on hydrogenase activity in whole cells, E. coli

cells fully induced for hydrogenase activity were exposed to nitrate.

E. coli strain K-10 was grown overnight in LB + 1.5% glucose under an

atmosphere of nitrogen. This culture was used to inoculate 40 ml of

LB + 1.5% glucose in two 70 ml serum bottles. The contents of the

bottles were flushed with nitrogen and the gas phase was replaced

with nitrogen. The pre-warmed medium was inoculated anaerobically

with a syringe. The inoculum size was 1% of the final culture

volume. The cultures were incubated at 37 C. After 170 minutes,

sodium nitrate was added to one of the cultures anaerobically, with a

syringe, to a final concentration of 11.76 mM. Samples were

withdrawn periodically to monitor growth by measuring the optical

density at 420 nm, and hydrogenase activity by assaying for tritium

exchange activity. Accumulation of nitrite in the medium was also

determined as described by Van'T Reit et al. (110).














RESULTS


Purification of Hydrogenase


Hydrogenase was purified from a prototrophic strain of

Escherichia coli K-12 (strain K-10). The details of the purification

procedure and the properties of the enzyme are presented below and in

Table 4-1.


Growth of the Cells: Escherichia coli, strain K-10, was grown

anaerobically in one liter fleakers filled to the top with LB

containing 1.5% glucose and incubated overnight at 370C. These cells

were used to inoculate 15 liter carboys containing minimal medium

supplemented with glucose at a final concentration of 3% and casamino

acids at a final concentration of 0.1%. The inoculum consisted of

10% of the final culture volume. The carboys were filled to the top

(anaerobic growth) and incubated at room temperature for 8 hours.

During the later stage of incubation, visible gas production could be

observed. The cells were harvested using a De Lavall separator, at

room temperature.


Cell Lysis: One hundred and eighteen grams of wet cell paste was

suspended in 750 ml of 0.1 M phosphate buffer, pH 7.0, containing














0.01M K-EDTA. The cells were lysed, using lysozyme, egg white

(Sigma). A freshly prepared stock solution of Lysozyme was added to

the cell suspension to attain a final concentration of

100 microgram/ml. The suspension was incubated at 37 C, shaking at

200 rpm, for 1 hour. The resulting cell lysis increased the

viscosity of the suspension. The viscosity was reduced by adding

deoxyribonuclease-I, 100 microgram/ml, ribonuclease-A, 100

microgram/ml and MgC12.6H20 to a final concentration 10 rm. The

extract was incubated further for 1 hour in a 37 C shaker, mixing at

200 rpm. The extract was centrifuged at 12,000xg for 10 minutes at

4C, to remove cell debris. The supernatant containing membrane

vesicles which had 83,433 units of hydrogenase activity (micromoles

of H20 produced/mg protein. hour) was used for further purification

of the enzyme, as described below.


Isolation and Solubilization of Membranes: Since hydrogenase in E.

coli is associated with the membrane (4,35), the next step was to

separate the membrane vesicles from other soluble proteins. To

obtain the membrane vesicles, the extract was centrifuged at

100,000xg in a swinging bucket rotor for 1 hour, at 4C (all of the

procedures henceforth were performed at 4 C and all the buffers were

incorporated with sodium dithionite at a final concentration of 1mM,

unless indicated otherwise). The pellet containing the membrane

vesicles was resuspended in 1,100 ml of 0.1 M phosphate buffer, pH














7.0. Approximately 327 of the hydrogenase activity was recovered in

the membrane pellet. To solubilize hydrogenase from the membrane,

Triton X-100, a non-ionic detergent, was added to a final

concentration of 1.0% and the extract was incubated for 1 hour with

gentle rocking. The Triton X-100 solubilized membrane fraction was

centrifuged at 100,000xg for 1 hour to remove the non-solubilized

membrane vesicles. The supernatant, containing the solubilized

membrane proteins, including hydrogenase, was collected.

Approximately 55% of the hydrogenase activity present in the

membranes was solubilized.


Enrichment of Hydrogenase: To remove some of the lipo-protein

complexes that interfere with the purification of the enzyme,

polyethylene glycol (PEG) was used. Solid polyethylene glycol 6,000

(recently renamed as PEG 3,000, Sigma Chemical Co., St. Louis, MO)

was added to the solubilized membrane protein fraction, to a final

concentration of 35% and incubated for 1 hour with gentle mixing.

The extract was centrifuged at 12,000xg for 30 minutes. The pellet

which contained the hydrogenase activity was resuspended in 500 ml of

0.1 M phosphate buffer, pH,7.0. The hydrogenase apparent specific

activity increased to 6.34 units from 4.19 units. Further enrichment

of hydrogenase was achieved by ammonium sulfate fractionation. Solid

(NH )2SO4 was added to the resuspended pellet to reach a final

concentration of 25% saturation. The mixture was incubated for 1













hour with gentle mixing. The proteins that precipitated were removed

by centrifugation at 12,000xg for 1 hour. The supernatant which

contained the hydrogenase activity was collected. Solid (NH4)2SO4

was added to the supernatant to attain a final concentration of 60;'

and incubated for 1 hour. The precipitated proteins were obtained by

centrifugation, at 12,000xg for 1 hour. The pellet, which contained

hydrogenase activity was resuspended in 200 ml of 0.01 M Tricine

buffer, pH 8.0 containing NaCI at a final concentration of 130 mM and

Triton X-100 at a final concentration of 1.0%. The resuspended

pellet was dialyzed against 6 liters of the same buffer for 6 hours.

The dialysis procedure was repeated one more time. This ammonium

sulfate enrichment procedure resulted in a 4.5 fold purification of

the enzyme without loss of total activity.


DEAE-Cellulose Chromatography: The enzyme was further purified by

loading the dialyzed extract on a DEAE-cellulose column

(2.8 x 90 cm.), equilibrated with 0.01 M Tricine buffer, pH 3.0,

containing NaCl at a final concentration of 130 mM and Triton X-100

at a final concentration of 0.3% equilibrationn buffer) and the flow

ratemaintained at 45 ml/hr. The column was washed with 600 ml of the

equilibration buffer. Hydrogenase was eluted with a linear gradient

of 150mM 225 mM NaCIl in a volume of two liters. Five milliliter

fractions were collected and assayed for hydrogenase activity as

described in the Materials and Methods section. Only the fractions














containing high hydrogenase activity (> 35,000 cpm per assay) were

pooled together and dialyzed against 12 liters of 10 mM phosphate

buffer, pH 7.0, containing 100 mM (NH4)2SO to remove the bulk of

Triton X-100. The protein fraction was dialysed twice against 6

liters of buffer for 6 hours. After dialysis, the remaining traces

of Triton X-100 were removed by adding Bio-Beads SM-2 (Bio-Rad) to

the extract and incubating the mixture for 2 hours, with gentle

mixing. As a result of DEAE-cellulose chromatography, hydrogenase

apparent specific activity increased by a factor of eight, although

the recovery was less than 20%.


Octyl-Sepharose Chromatography : To remove most of the hydrophilic,

and some of the hydrophobic protein contaminants, an Octyl-Sepharose

column (2.8 x 40 cm.)was used. The column, fitted with a reverse

flow adaptor and maintained at a flow rate of 25 ml/hour was

equilibrated with 100 mM (NH4)2SO4 in 10 mM phosphate buffer, pH 7.0

equilibrationn buffer). The partially purified hydrogenase was

applied to the column and the column was washed with 150 ml of the

equilibration buffer. Hydrogenase was eluted with a 500 ml linear

gradient containing, initially, the equilibration buffer and finally,

sodium deoxycholate (0.50 % w/v) and Triton X-100 (0.05 % v/v) in 1

mM phosphate buffer. Two and one half milliliter fractions were

collected. The fractions were assayed for hydrogenase activity as

described in the Materials and Methods section. Hydrogenase eluted













as a single peak at 0.225% Sodium Deoxycholate and 0.0225% Triton

X-100. Fractions containing hydrogenase activity (> 45,000 cpm per

assay) were pooled together. Even though hydrogenase was enriched

only by a factor of 1.03 during this step, it was important to

include this step in the purification procedure, to separate the

hydrogenase from some of the other hydrophobic contaminants which

interfered with further purification. The pooled fractions were

dialyzed against 12 liters of 25 mM Histidine HC1, pH 5.5, containing

0.3% Triton X-100. The dialysis was performed in two steps against 6

liters of buffer for 6 hours each time.


Chromatofocussing: Further purification of the enzyme was achieved by

taking advantage of the fact that each protein has its own unique

iso-electric point. Chromatofocussing achieves separation of

proteins based on their isoelectric points. The dialyzed protein

solution containing hydrogenase was applied to a column (1.3 x 45 cm)

of Poly Buffer Exchanger (Pharmacia) equilibrated with 25 mM

Histidine HC1, pH 5.5 containing 0.3% Triton X-100 and maintained at

a flow rate of 20 ml/hour. Hydrogenase activity was eluted by

passing 1,200 ml of Polybuffer 74 adjusted to pH 4.0 with HC1 and

supplemented with 0.3% Triton X-100, through the column. Two ml

fractions were collected. The fractions were assayed for hydrogenase

activity as described in the Materials and Methods section.

Hydrogenase activity eluted as a sharp peak at pH 4.4. Fractions













containing hydrogenase activity, (>23,000 cpm per assay), were pooled

together and dialyzed against 6 liters of 50 mM NaCI in 10 mM

phosphate buffer, pH 7.0, containing 0.3% Triton X-100.

Chromatofocussing resulted in a 4 fold enrichment of hydrogenase

activity.


DEAE-Cellulose Chromatography: To purify the hydrogenase to

homogeneity, the dialyzed sample obtained after chromatofocussing was

applied to a DEAE-cellulose column (1.2 x 30 cm). The column was

equilibrated with 50 mM NaCl in 10 mM phosphate buffer, pH 7.0,

containing 0.3% Triton X-100 equilibrationn buffer) and maintained

at a flow rate of 15 ml/hour. The column was washed with 100 ml of

the equilibration buffer. Hydrogenase was eluted with a 500 ml

linear gradient of 50-125 mM NaCl in 10 mM phosphate buffer, pH 7.0,

containing 0.3% Triton X-100. Two ml fractions were collected and

assayed for hydrogenase activity. Hydrogenase eluted at

approximately 90 mM NaCl. Fractions containing hydrogenase activity

were analysed for purity, using 7.5% non-denaturing and 12% SDS-PAGE.

The gels were stained for protein using the silver stain method.

Fractions containing pure hydrogenase, based on SDS-PAGE, were pooled

together and concentrated by ultrafiltration using an Amicon PM-10

membrane ultrafilter. The concentrated sample was also checked for

purity by subjecting the sample to electrophoresis as mentioned

above. The hydrogenase sample subjected to electrophoresis under














non-denat'ring conditions was also stained for hydrogenase activity

as described in the Materials and Methods section. Results from

native PAGE (Figure 4-1) showed a single band, when stained for

protein using the silver stain method. The Rf of the protein band

was comparable to the Rf of a protein possessing hydrogenase

activity. Figure 4-2 presents the results obtained after the

purified enzyme was subjected to SDS-PAGE and the gel stained for

protein. A single protein band was detected with a molecular weight

of 56,000. The procedure described above yielded a protein which is

enriched for by a factor of 690 as compared to its presence in the

crude extract. The final yield is 1.43%.



Molecular Weight Determination


To determine the molecular weight of the native enzyme, two

different methods were used, namely, gel filtration and native

polyacrylamide gel electrophoresis. Also, since the enzyme was

solubilized using Triton X-100, and the detergent was present during

the purification procedure, the molecular weight was determined, both

in the presence and absence of Triton X-100.




















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Figure 4-1.


Polyacrylamide gel electrophoresis of purified
hydrogenase under non-denaturing conditions.
A, Gel stained for protein using the silver stain
method;
B, Gel stained for hydrogenase activity as described
in the Materials and Methods section.
Amount of protein added to each lane was
2.6 micrograms








45








A B



































Figure 4-2. SDS-Polyacrylamide gel electrophoresis of purified
hydrogenase.
Lane A, purified hydrogenase, (2.0 micrograms);
Lane B, Molecular Weight standard proteins.
Gel stained for protein using the coomassie blue
method.








47


A B







a




e






e



-














Gel Filtration in the Presence of 0.3% Triton X-100: To determine the

molecular weight of hydrogenase by gel filtration, a Sephadex G-200

column (2.6 cm x 48 cm) was used at 4C. The column was equilibrated

with 10 mM phosphate buffer, pH 7.0, containing 0.3% Triton X-100 and

maintained at a flow rate of 11 ml/hr. The void volume (Vo) of the

column was determined using blue dextran (MW=2,000,000). A

calibration curve for the column was generated with proteins of known

molecular weights as described in the Materials and Methods section.

Table 4-2 lists the void volume (Vo) of the column, the elution

volumes (Ve) of the molecular weight standard proteins and the ratio

of Ve/Vo for each of the proteins. To determine the molecular weight

of hydrogenase, 1.5 ml of pure hydrogenase (2.6 microgram/ml) in 10

mM phosphate buffer, pH 7.0 + 0.3% Triton X-100 was loaded on the

column. The elution of hydrogenase was monitored by assaying the

different fractions collected (0.93 ml) for tritium exchange

activity. The elution profile of hydrogenase from a representative

experiment is presented in Figure 4-3. Hydrogenase activity was

detected in two major peaks. The molecular weight of hydrogenase

from each peak was determined using the caliberation curve (Figure

4-4). Based on three independent determinations, 22% (+/- 6W) of the

activity loaded on the column was detected in peak I and 68% (+/- 9%)

of the activity eluted in peak II. Hydrogenase in peak I had a

molecular weight of 133,000 (+/- 6,000) and the hydrogenase in peak

II had a molecular weight of 62,500 (+/- 4,000).

























*H *O
) oe

oO -
0-
,-I
CL X










,0 c
-4


0



.-

-o
Q IUn

tO 0

3 )



















0
. S=
0 "3
0














,-- 10
4-)
(1)
0 *
TH





C 0)
r














'-4 -':

m a
C 0)
c co


0 0
r-^1>


C-

- 0 *





r1> -






(1)

0 :r















r-
0
0
S..
0.L


LC








LM






0
Ll


















0
L'D




0
0
L l
















a)
U)
U)
i-



















0)
L0













0
0
00
HOn










hO
**1 0


0
0
0











S-H












-0


O 0
0 0


0 0



CM











'-.'
00

CL
a



U Q)


4:* *
0 0

Oc)





0 0





4-)!-
LL 0











-4-I -
'- '-,



Co
OO




WO
+ 1
0
0

cu
>1: Q)
= 0









































0 C

I 0

0

(-J
Q. 0











DE
CQ.q








,-4 CO


0 C.
Sx-
0
CO






O 0
Q0)






S-
oJ Q
*C -





0)













51





Q








C















c c



c
























( 0 x wdo)

,1"TAtIZV aSEUaaO-IPAH


































Figure 4-4. Calibration curve for the Sephadex G-200 column
(2.6 cm x 48 cm) used to determine the molecular
weight of hydrogenase in the presence of Triton
X-100. 0 molecular weight standards;
,i hydrogenase.




























































Ve / Vo


20.0-


10. 0o


8.0

6.0




4.0


2..0 L


1.0


1.25


1.75


2.25


2.75













Gel Filtration in the Absence of Triton X-100: To determine the

molecular weight of the enzyme in the absence of Triton X-100, a

column of sephadex G-200 (2.6 cm. x 48 cm.)was used and the Ve/Vo

values for the molecular weight standard proteins were determined in

the same manner as for gel filtration in the presence of Triton X-100

(Table 4-2). The Ve/Vo did not change appreciably for these

proteins, in the absence of Triton X-100 except for albumin and

carbonic anhydrase which migrated little faster through the column.

The Ve/Vo for the enzyme (2.6 microgram/ml; 1.5 ml sample volume) was

determined after removing Triton X-100 as described in the Materials

and Methods section. The elution profile of hydrogenase was

monitored by assaying the different fractions for tritium exchange

activity (Figure 4-5). Hydrogenase activity was present in two major

peaks. Table 4-2 lists the elution volume (Ve) and the ratio of

Ve/Vo for the two hydrogenase peaks and the standard proteins used.

The molecular weight of hydrogenase in the two peaks was determined

using the caliberation curve (Figure 4-6). Based on three

independent determinations, 57% (+/- 9%0) of the activity loaded on

the column was accounted for in peak I and approximately 33% (+/- 7%)

of the activity was detected in peak II. Hydrogenase in peak I had

a molecular weight of 125,000 (+/- 10,000) and the hydrogenase in

peak II had a molecular weight of 58,000 (+/- 5,000). The minor peak

between peak I and peak II had a molecular weight of 77,000 and

accounted for O% of the total activity.






































0
I
X


0
C-.
0 ,-i




x 0
C)
o a)
Cto











0 -



- co
Co"





o--
r-)
taO
0





-1

4*-O
0.


0
*r-f




40






bO
-4











































Q z
C !


















-4)
-0


























(C-OI x ido)
AITA13.V aVseuaojpOAH



































Figure 4-6. Calibration curve for the Sephadex G-200 column
(2.6 cm x 48 cm) used to determine the molecular
weight of hydrogenase in the absence of
Triton X-100. Q(, standard proteins;
*, hydrogenase.





























Peak I


Peak II


1.75 2.25


Ve / Vo


20.0







10.0

8.0



6.0




4.0


2.0 L1


1.25


2.75














Native PAGE In The Presence of Triton X-100: Another independent

method used to determine the molecular weight of hydrogenase involved

electrophoresis of the enzyme in polyacrylamide gels of different

concentrations under non-denaturing conditions. A set of tube gels

containing different concentrations of acrylamide was prepared and

the samples were subjected to electrophoresis, as described in the

Materials and Methods section. The relative mobility of hydrogenase

was determined after staining the gels for hydrogenase activity

(Figure 4-7, Table 4-3). To determine the molecular weight of

hydrogenase, the relative mobilities of proteins with known molecular

weights were determined as described in the Materials and Methods

section. The Ferguson Plot generated based on the Rf values obtained

for the standard proteins is as illustrated in Figure 4-8. The

average molecular weight of hydrogenase, based on three independent

determinations was 58,000 (+/- 6,000).


Native PAGE in the Absence of Triton X-100: The molecular weight of

hydrogenase, using the electrophoresis method was also determined in

the absence of Triton X-100. The tube gels were prepared and the

samples were subjected to electrophoresis as described in the

Materials and Methods section. The only difference in this case was

that Triton X-100 was not incorporated in the gels and hydrogenase

was free of Triton X-100. The gels were stained for hydrogenase

activity after electrophoresis. Under these conditions, two distinct




























Figure 4-7.


Molecular weight determination of hydrogenase in
the presence of Triton X-100 using native PAGE.
Polyacrylamide tube gels with different
concentrations of acrylamide were loaded with
purified hydrogenase (2.6 microgram).
Electrophoresis was performed and the gels stained
for hydrogenase activity as described in the
Materials and Methods section. 1, 5.0% acrylamide;
2, 6.0 % acrylamide; 3, 7.0 % acrylamide;
4, 7.5 % acrylamide; 5, 8.0 % acrylamide and
6, 9.0 % acrylamide.









































12 3 4 5


Mom*

























b3






.-4
00




O ) o
7-4

c m





0 0.





00

0


Cl) 4-
.) -4

-4 i)


0 .)
O0




0 0.

C))
L C





0L




o. 0.
4.) CO 1
) r-







(13 0
Q 0 4.)





CL
caO
S0 -4


a) c -O
-4> 0


I.-
E )

m 00
.C 4.0 >
Q. O C) -:
r a)


0
0
0









b0
a)
c c
2CC

. .4
3 0



0
0

a)
L-0 2
0I--.
\-O E
0




3
-.o



5 a)
.0 >
0
<. .-


(\ -'-
mC L
- Ci


E


S-
,I)



.0 >
0
<* C3


t0
0
































Figure 4-8.


Standard curve (Ferguson plot) for the determination
of the molecular weight of hydrogenase using native
PAGE in the presence of 0.3 % Triton X-100.
Q hydrogenase; *, molecular weight standard
proteins.











64


















10.0

-t 8.0 -


x 6.0



S .0.



4.0



2.0






1.0 t I I I I
1.0 2.0 4.0 6.0 8.0 10.0


Slope













bands that stained for hydrogenase activity can be observed in the

gels (Figure 4-9). The relative migration of both these bands

possessing hydrogenase activity and the relative mobilities of the

standard proteins are listed in Table 4-4. Figure 4-10 illustrates

the Ferguson Plot generated using the Rf values obtained for the

molecular weight standard proteins. Based on three independent

determinations, hydrogenase with a higher Rf value (band I)

corresponds to a molecular weight of 53,000 (+/- 5,000) and the other

band (band II) corresponds to a molecular weight of 115,000 (+/-

10,000).



Iron and Sulfur content


The iron and acid labile sulfur content of the enzyme was

determined as described in the Materials and Methods section. The

two forms of the enzyme were separated by gel filtration. The enzyme

with a molecular weight of 125,000 had 8.87 (+/- 0.34) moles of iron

and 8.91 (+/- 0.56) moles of sulfur per mole of the enzyme. The

58,000 d enzyme had 4.4 (+/- 0.15) moles of iron and 4.74 (+/- 0.48)

moles of sulfur per mole of the enzyme.



Temperature Profile


The optimum temperature for catalysis of the exchange reaction

was determined, as described in the Materials and Methods section.






























Figure 4-9.


Molecular weight determination of hydrogenase in
the absence of Triton X-100.
Polyacrylamide tube gels with different
concentrations of acrylamide were loaded with
hydrogenase (2.6 micrograms). After
electrophoresis the gels were stained for
hydrogenase activity as described in the
Materials and Methods section. 1, 4.0 acrylamide;
2, 6.0 % acrylamide; 3, 7.5 % acrylamide;
4, 10% acrylamide; and lane 5, 12% acrylamide.








67

















i


1 2 3 4



























"-4-




0
r-l






0
CI
S-








C-







0
0c









C C
-O




0 ,







aO -




.-.
0-c-










~- (


'-4 D-
Q(U)


p


1D0
LC


,c.c
- 0


0
O

0 S-
.o


E
0
0
E




C
0
L
0

^ 0
E C
3 *r-l
.0 >
-O 0


0



r-
0








L.
0
CO
E
S-4



E 0

ef -4


O
0
0
-1









CO



0
" 7
0)

o c


*
0
0
0









0
1)
(-:







c

0
-. '0
S"O C
3: CO


0
0
C-D
r\J









c
E
-3
8
-Q

'-4m
0. 0
r- ..
<.,C1

































Figure 4-10.


Standard curve (Ferguson Plot) for the
determination of the molecular weight of
hydrogenase using native PAGE in the absence of
Triton X-100. hydrogenase; molecular
weight standard proteins.



























Band II


Band I


I I I I


2.0


4.0


6.0 8.0 10.0


Slope


10.0


8.0


6.0



4.0







2.0


1.0


1.0













Figure 4-11 shows the temperature profile for hydrogenase as

determined by the exchange reaction. The data represent the results

of four independent determinations. As can be deduced from the

graph, the optimal temperature for the exchange reaction was 350C.

Figure 4-12 presents an Arrhenius plot for the tritium exchange

reaction catalyzed by hydrogenase between 15 C and 35C. The slope

of the line obtained by plotting the log of the reaction rate versus

the inverse of the absolute temperature at which the reaction was

performed, gives the activation energy for the exchange reaction

catalyzed by hydrogenase. Hydrogenase shows two activation energies

for the exchange reaction, the activation energy is 3657 cal at a

temperature range of 35 C to 20 C and 3,517 cal at temperature below

200C. To determine whether the higher activation energy required at

temperatures below 200C was due to micelle formation by Triton X-100

at lower temperatures, the experiment was also performed in the

absence of Triton X-100. The results obtained in the absence of

Triton X-100 were similar to the results obtained in the presence of

Triton X-100, suggesting that the higher activation energy value

observed at temperatures below 20 C was not due to the presence of

Triton X-100.































U I I I I


I I I


I I I


(.noq uiauoad Jm/paonpoad OzH Jo saiotuo.3itu)


.\' A. i V asEu3o0ipAH


C C-.
O
C;
C 0






C-
0












-4


I I I


I I





































-0 0


NO
>L

0


m 0
00


,) L
C o







0 Q.
-E
V. S-







C)




0
Qo




4 .
QD



00
0.



0 *



cL3
(1-J







*C
ry-




























I


A 'o0















pH Profile


The optimum pH for the catalysis of the exchange reaction was

determined as described in the Materials and Methods section at

different pH and in the presence of three different buffers. The

buffers used and the pH at which the exchange reaction was carried

out were as described in the Materials and Methods section. Figure

4-13 illustrates the pH profile for the exchange activity. A broad

pH optimum between pH 7.0 7.5 was observed for hydrogenase using

the tritium exchange reaction.



Kinetic Characteristics


Since the purified hydrogenase can exist both as a monomer and

a dimer, depending on the presence or absence of detergent, it was

important to determine whether both the monomer and dimer have the

same kinetic properties. To achieve this, the two forms were

separated by gel filtration. The apparent Km and Vmax for the

exchange reaction were determined for both forms of hydrogenase. The

results presented in Table 4-5 show that the apparent Km for hydrogen

and the Vmax for the exchange reaction of the monomer and dimer forms

of the enzyme are comparable to the apparent Km and the Vmax of the

enzyme before the separation. The turn over number calculated based

on the molecular weight values obtained using Gel Filtration suggests




































C~f

LO
C) w
*o
C/ C




r-4 1)
SCL-4







C Q





C) (1*
CO Q


0 3
LC).0

-0
C.Lr c
C fl) n












0 0. 0

-0

O.U
wO '-' s-










C)
0 -






C-
0
to



r.-
60
^ c/









73







































U-






























0 0 0 0
0 0 0
CIA 00
( .inoq *uiaoaid 2w/paanpoId 0 HcJO salowo.oipm)
.,\:AT3)V aSeUaodZOpH














79







C'

hO
0






S0 0
v 0
o '-





0: "- > *- 0 .J










00 0
S- 0
. 0 0(-) cM DC





gI I CC



-0 > 0- C -Y 0. 0








cC 0
0 3 0 0
o C) 3 C







0 0 >

-0 S 4: Iz E
CM- CO




H 0 O.J











^O L OQ S Qm a- i














that the dimer form of the enzym? is composed of two monomers that

are equally active.


The apparent Km was also determined for different electron

carriers of hydrogenase viz: oxidized benzyl vLologen, oxidized

methyl viologen, reduced benzyl viologen, reduced methyl viologen and

hydrogen. The determinations for each of the substrates were done as

described in the Material and Methods section. The computer

generated Lineweaver Burk plots for each of the substrates is as

presented in Figures 14-14, 4-15, 4-16, and 4-17. The results

obtained are an average of three independent determinations and are

summarized in Table 4-6. The apparent Km for oxidized methyl

viologen in the hydrogen uptake reaction was 26.7 mM and the forward

reaction was catalyzed at a maximal velocity of 24.3 micromoles of

methyl viologen reduced/min mg protein. Monitoring the rate of

hydrogen evolution, the apparent Km for reduced methyl viologen was

determined to be 1.5 mM and the Vmax = 35.1 micromoles of hydrogen

produced/min. mg protein. The apparent Km for oxidized benzyl

viologen in hydrogen uptake reaction was 7.7 mM and the Vmax = 49.4

micromoles of benzyl viologen reduced/min mg protein. The apparent

Km for reduced benzyl viologen was 4.0 mM and the maximum velocity at

which the rate of hydrogen evolution reaction proceeds, using reduced

benzyl viologen as a source of electrons was 12.5 micromoles of

hydrogen produced/min. mg protein.










































>
7O 4-)

*M *H

.-4 C)

0

0 )O
loo

0O
- 0






CD
I2 C


a) 0
C -4-
SJ-)


4-)
0 <1)
C)
,-- 0

0
C

0


-.04
*H -4
0






0 ")









to
b 0
.- I
L ._
LL. z













82




























I















I I



OOCC
-4rf








































N>
H *r-

0
- C)


0
OT


0 0

CQ 0


>








i- O
C.
0



S00






.-4
CL
e- G

































C'












C'

CNE






N







































4-)
o -4




0
C) >




0* 0
LO


0
>C







0
" ro
LO






>C

00



4C
OJ



0 0



L.
*o ro
0.4










*
L.O
.r-l








L-H














36























(c






















co o oL



O C
r-i *







































>



0 0


C)

0
LO
s- 0
C3 "


>C
0
0
0C







-4 0
0
OC

S-

L 0
0

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0 )
00


















*-










































c
C











Ln 2
E


--





N--
















89


































to t b .0 *O
O 0 C)O
oI r


L I
c *-








to ..- >


0 0 0 0





*- O O
N N C) C)

o -\ 0

o > 0 >





0 .0 0 0 0





--* C C) C CO C)
o x a














Hydrogen Uptake in the Presence of Different Artificial Electron

Acceptors


The ability of hydrogenase to utilize a variety of artificial

electron acceptors was also determined. The experiment as described

in the Materials and Methods section involved the incubation of

hydrogenase with various electron acceptors in the presence of

hydrogen and determination of the rate of reduction of the acceptor,

spectrophotometrically. Table 4-7 lists the values obtained for the

rate of reduction of the various acceptors and their redox potentials

(Eo'). As shown in Table 4-7, hydrogenase can use all five compounds

as electron acceptors. The results indicate that the rate of

hydrogen uptake proceeds at a more rapid rate in the presence of

methylene blue and potassium ferricyanide, compounds with a positive

Eo'.



Inactivation of Hydrogenase by Oxygen


The inactivation of hydrogenase by oxygen was determined by

incubating the enzyme (1.7 microgram/ml) in the presence of air in a

12 x 75 mm tube on a rocking platform at room temperature. At

various time intervals aliquots were withdrawn to monitor tritium

exchange activity. The control sample of hydrogenase was maintained

under similar conditions, but the gas phase was hydrogen. Details of









































*

0










O>
4-1

-r-4




-0

























0
0
E








L
O

41



w
0O
C)


0
(U








4-)






ci)


0 0 O o













N! -n L-- (












O O CM -

I I I + +













0 0 -

0 0 ) -1
*-14 *- L I) E C
> > C 3O
Q--1 .) -q >



0) 0 (V 0 O)
Sc a i


4

UCD

~C)



bO



*r-1 i
-4-)
0
Wa


















O 0

b0: 0
b- ci)







CL
o 1)


*-1
> 4-'

0 "/

D -










GL a
TO *:S0

0. 0



-c ci


Ca )




C)M
f-4Y
.0ci
C> a













the experiment are as described in the Materials and Methods section.

Figure 4-13 gives the oxygen inactivation profile of the hydrogenase

exposed to air and the hydrogenase sample maintained under hydrogen.

The half-life of the hydrogenase in the presence of air was 650

minutes. The inactivation of the monomer and the dimer form of

hydrogenase due to oxygen was also determined. The two forms of

hydrogenase were obtained by Gel Filtration as described in the

Materials and Methods section. The experiment was performed as

described above. The inactivation profile for the first thirty

minutes is as illustrated in Figure 4-19. It is apparent that both

the monomer and dimer form of hydrogenase are equally stable in air.



Stability of Hydrogenase at Alkaline pH


Ballantine and Boxer (10) recently reported the presence of two

different hydrogenases in E. coli. They detected two distinct bands

of hydrogenase activity when Triton X-100 solubilized membranes were

subjected to PAGE at neutral pH. They observed that one of these

bands was labile at alkaline pH (pH 10.0), and lost the hydrogenase

activity. Thus the stability of the purified hydrogenase at alkaline

pH was checked. Hydrogenase was incubated at pH 10.0 for 15 minutes

and then the activity was determined by monitoring the exchange

reaction and the hydrogen uptake reaction at, pH 7.0 and pH 10.0.

For the exchange reaction, 75 microliters of 100 mM glycine, pH 10.0,














or 100 mM phosphate buffer, pH 7.0, was placed in a 12 x 75 mm tube.

The tubes were capped with serum stoppers and the gas phase replaced

with helium. Hydrogenase (25 microliters, 12.0 microgram/ml in 10 mM

phosphate buffer, pH 7.0) was added to the tube and incubated at room

temperature for 15 minutes. After 15 minutes, the pH of the assay

mixture was brought to neutrality or maintained at pH 10.0, by adding

100 microliters of 100 mM phosphate buffer, pH 6.5, or 100

microliters of 100 mM glycine buffer, pH 10.0. Tritium exchange

reaction was performed as described in the Materials and Methods

section. For the hydrogen uptake reaction, 500 microliters of either

100 mM glycine buffer, pH 10.0, or phosphate buffer, pH 7.0, was

placed in a 12 x 75 mm tube. The tubes were capped with serum

stoppers and the gas phase replaced with hydrogen. Hydrogenase (100

microliters, 12.0 microgram/ml in 10 mM phosphate buffer, pH 7.0) was

added and incubated at room temperature for 15 minutes. After 15

minutes, the pH of the assay mixture was brought to neutrality or

maintained at pH 10.0 by adding 1.9 ml of either phosphate buffer, pH

7.0, or 100 mM glycine buffer, pH 10.0. The rate of hydrogen uptake

reaction was determined by monitoring the reduction of benzyl

viologen as described in the Materials and Methods section. The

results presented in Table 4-8 show that the enzyme retained 62.5% of

its exchange activity and 76.5% of its hydrogen uptake activity after

incubation at pH 10.0 for 15 minutes. It is also evident that the