I 'PRODUCTION, STRUCTUAL ANALYSIS AND FL!JNCTIONAL
DEVELOPMENT OF USEFUL PROTEINSMITSUO YAMASHITA

T ABLE OF CONTENTSPREFACE
Chapter I~.

1Expression of a Gene for Human Protein:

Genetic Design of a Gene for Human Metallothionein II and Its Expression as
an Active Fusion Protein in Escherichia coli
2. Materials and methods

9
10
11

3. Results

15

4. Discussion
5. Summary

24
27

6. References28

1. Introduction

Chapter II

Production and Purification of Monoamine Oxidase:

Production, Purification and Characterization of a new type of Monoamine
Oxidase containing Cu from Klebsiella aerogenes

32

1. Introduction

33

2. Materials and methods

35

3. ResuHs

39

4. Discussion

51

5. Summary

54

6. ReferencE~s

55

Chapter III

Structure and Function of Pullulanase:

Role of Lipid Modification on a Starch-debranching Enzyme, Klebsiella
Pullulanase60

1. Introduction

61

2. Materials and methods

63

3. Results

65

4. Discussion75

5. Summary

77

6. References

78

Chapter IV

Functinal Development of Pullulanase:Random mutagenesis of Pullulanase from Klebsilella aerogenes for Studiesof the Structure and Function of the Enzyme

80

1. Introduction

81

2. Materials and methods

83

3. Results

85

4. Discussion

96

5. Summary

j

•

100

6. References

101

GENERAL DISCUSSION (CONCLUSION)

107

REFERRENCE PUBLICATIONS109

RELATED PUBLICATIONS111ACKNOWLEDGEMENTS

113

PREFACE
The advent of' gene technology in the 1970s has led to the flourishing of theIbiotechnology industry. In the 1980s, the attention was focused on
biotechnology as a result of the enormous potential of recombinant DNA
technology and genetic engineering. This expansion in commercial activity isbased largely on the ability to produce proteins in heterologous cell systems
at industrial scale. Although the conventional mutation and screening
methods are still used to increase the productivity of already existing activities
in microbial cells, recombinant DNA technology allows creation of novel
activities that can be directed toward the synthesis of huge amount of products
normally produced at lower levels by other organisms. The rapid
developments in recombinant DNA techniques have resulted in theidentification and isolation of various novel genes involving several known
and unknovvn functions. There is almost invariably a need to express the genein a heterologous cell system in order to produce : 1) material for structurefunction studies, 2) diagnostic reagents (e .g. monoclonal or polyclonal
antibodies), 3) material for in vivo activity testing .
Several alternative systems for the expression of foreign genes have beendeveloped , an extension of in vivo genetic manipulation or intergenic genetransfer, in mammaliam cells, insect cells, fungal cells, bacterial cells, andtransgenic animals or plants. The choice of expression system for a given
gene depends upon the properties of the encoded protein (e.g . protein
modifications needed for biological activity) and the objective of the study (e.g.
structure-function analysis or production of diagnostic reagents) . Other
important considerations for the investigation are the available facilities, timeand cost involved in generating the amounts of recombinant protein required.
The application of recombinant DNA technology to the production of usefulgene products was focused initially on the research or the commercial level in

Escherichia coli. E. coli has become an industrial microorganism since the1beginning of the modern fermentation industry. How has E. coli been
converted to a producer of useful products? Many examples of the production
I'of heterogenE30us proteins in recombinant microorganisms have beenreported . Ho~/Vever, the productivity of heterogeneous proteins differs
considerably from protein to protein even in the same host, which is E. coli.
The most widely used and convenient system for the production of foreign
proteins is based on the simple prokaryote, E. coli. The advantages of thissystem are : '1) ease of gene manipulation, 2) availability of commercial
products and reagents, including gene expression vectors, 3) ease of
producing quantities of protein (up to a few gram in simple shake-flask
culture) , 4) speed of culture, and 5) adaptability of the system , thereby
allowing solution to expression problems.
Expression of any foreign gene in E. coli begins with the insertion of the gene
into an expression vector. Many forms of expression vector are available andthey usually consist of : 1) a plasmid origin of DNA replication, 2) antibiotic
selectable nlarker, 3) strong promoter and transcriptional terminator separated
by a multicloning site (expression cassette) , and 4) DNA sequences encoding
a ribosomal-binding site.
When plasmids are used as vectors, plasmid stability, copy number, andeffect on growth of the host are very important considerations. Several genetic
menipulations are known to increase plasmid stability. Mutation of the hoststrain to lose both uptake of an amino acid and auxotrophy for that amino acidand insertion of the amino acid biosynthetic genes on the plasmid result to theplasmid-dependent growth of the host strain even in complex medium
containing the amino acid . Plasmid-free segregants die under these
conditions.
There is very little difference between the various strong promoters now
available (pTac, "'pL, ApR, T7) except in the method of transcriptional
regulation . pTac and T7 expression systems are controlled by the chemical2

inducer isopropyl-tl-O-thiogalactoside (IPTG), whereas the ApL and ApR..

promoters b), a temperature shift. Gene expression can alsq be controlled
through the use of inducible copy-number vector. The dual-origin vector relies
upon a temperature switch to increase plasmid copy number from five to 200
copies/cell, ,~hich induces foreign gene expression. Examples showing theuse of these vectors is given below.
The translation of foreign mRNA remains a significant problem associatedwith gene e)cpression in E. coli. The basic rules for achieving translation ofmRNA are simple : 1) complementarity between the Shine-Oalgano (SO)sequence and the 3'end of the 16S ribosomal RNA, 2) a 6-10 bp spacing
between the! SO and the initiation codon (AUG), 3) an AT-rich basecomposition between the SO and the AUG . Regions of internal
complemen1tarity within the mRNA, paticularly those involving the SO andAUG should be avoided, since uncomplementarity in these regions
significantly reduces the frequency of translational initiation. In instanceswhere translation is a problem , a variety of approaches is available toovercome this obstacle.
However, 'we still sometimes encounter problems or limitations in the efficient
expression and production of heterologous protein products in E. coli. This
maybe partly due to the disadvantages of E. coli for foreign gene expression
which are: "1) inability to carry out post-translational modifications typical ofeukaryotic cells and 2) the tendency of foreign proteins to form insoluble
inclusion bodies when expressed intracellularly.
Recent advances have shown that it is possible to engineer the E. coli hostcell genetically to carry out at least a kind of post-translational modification,
which is m),ristoylation of a foreign protein . This result suggests that otherpost-translational modifications can be carried out. The problem of protein
insolubility can be addressed either by refolding the denatured protein in vitro

3and varying the conditions of expression (e.g. temperature), or secreting into
the periplasnl or culture medium .
j'

Accumulation of foreign protein intracellularly may be affected by proteolysis.
This is particularly relevant to small peptides or to protein fragments. Two
approaches have been used to overcome this problem, namely the use of
host strains 'with reduced protease levels and the production of fusion proteinsthrough the ~ n - phase translational fusion of the gene sequence to the Cterminus of :a highly expressed homologous protein . Genetically engineered
proteins can be produced with special tails to facilitate purification . Tails canbe ion-exchange, hydrophobic and covalent, metal chelate, affinity, and foracid precipitation . The fusion protein product is often purified using theproperty of the homologous fusion protein partner and cleaved to release thepeptide or protein fragment of interest. Fusion vectors have been designed to
produce unique chemical or protease cleavage sites between fusion partnerand protein for this purpose. The fusion protein approach has an additional
advantage, in that it also overcomes the problem of translational initiation,
since the N·-terminus of the fusion protein is a highly expressed homologous
protein.
Proteins play central roles in all life processes, catalyzing biochemical
reactions with remarkable specificity and serving as key structural elements inall cells and tissues. As enzymes, they catalyze both the digestion of food
stuffs and the construction of new macromolecules. As collagen, actin, myosin ,and intermE3diate filaments, they control the structure and motion of cells andorganisms. Antibodies protect us against disease. Membrane proteinsregulate ion transport and intercellular recognition . Repressors and activators
regulate gE~ne expression. Polymerases replicate genes, and histones help
package DNA into chromosomes. They are involed in every aspect of life (e.g.
catalysis, structure, motion. recognition , and regulation) and indispensable to
all life.

The properties of many enzymes have been altered by genetic means.
Mutagenesis and random screening of microorganisms over the years haveI 'led to the change in optimum pH, thermostability, feed back inhibition, carbon
source inhibition, substrate specificity, Vmax, Km, and Ki. Such information is
exploited now by the more rational technique of protein engneering. Single
change in amino acid sequences have yielded similar types of changes in alarge variety of enzymes. Today it is no longer necessary to settle for an
enzyme's natural properties since these can be altered to suit the needs of theinvestigator or the process.
The ability to modify some proteins has increased our understanding of avariety of irnportant biologically relevant processes, including protein stability,
catalysis, a.nd substrate binding. We need theoretical approaches to better
understand these properties. With the appearance of fast and affordable
computers" it is now practicable to study complex macromolecular systems
with computer simulation methods.
In recent decades, tremendous progress has been made in understanding
protein structure and function. The amino acid sequence of insuline wasreported in the early 1950s and the three-dimensional structure of myoglobin
was elucidated in 1960s. Genetic engineering was born in 1973. Progress ingene synthesis and genetic engineering have made it possible, in principle, toconstruct any desired amino acid sequence, which started the epoch ofprotein engineering in the early 1980s.
Protein €!ngineering is a more recent development which has important
implications for the biotechnology industry. Clearly, the capability of designing
novel prot,eins broadens the opportunity to identify new products with alteredfunction or improved stability or potency. Since the introduction ofrecombinant DNA methodology, genes can be removed from their nomal
environmE~nt in an intact genome and isolated as DNA fragments on cloning

vectors. The availability of purified genes in vitro in microgram amounts has

5dramatically' expanded the potential for inducing mutations. In controlledenvironment using test tube, it is now possible to alter, efficiently and.'.

systematica.lly, the sequence of nucleotides in a segment of DNA. The new
methods of in vitro mutagenesis are divided into three broad categories: 1)
methods that restructure segments of DNA, 2) localized random mutagenesis,3) site-directed (oligonucleotide-directed) mutagenesis. This classificationemphasizes the practical aspects of each method's application. Analogues
can be produced by recombinant DNA technology: 1) hybrid proteins can beproduced by in vitro recombination of hybrid genes, 2) analogue polypeptides
can be produced by chemical synthesis of DNA that contain triplets encoding
a different amino acid that is present in the natural polypeptides, 3) a cloned
gene can be randomly mutated before insertion into the vector, 4) site-specificmutagenesis can be employed using DNA polymerase to prepare mutant
synthetic oligonucleotide primers on single-stranded DNA templates, 5)deletions or insertion of a few nucleotides in length can be carried out withrestriction lenzymes, and genes can be truncated by insertion of stop codons.
Protein engineering should have tremendous theoretical and practical
implications. It can be used both to explore fundamental questions aboutprotein folding, structure, and function, and to design useful protein for
medical and industrial applications. A flow chart of protein engineering is asfollows: 1) cloning of the gene of an enzyme, 2) determination of thenucleotide, sequence and/or amino acid sequence of the enzyme, 3)
biochemical characterization of the enzyme, 4) determination of threedimensional structure of the enzyme, 5) molecular design of the enzyme, 6)
alteration of the gene of the enzyme by site-directed mutagenesis, 7)
characterization of the mutant enzyme, and back to step 5). Since protein
engineering is an interdisciplinary research field, the cooperation among
biologist, chemist, and physicist is essential for its success.

6

In the area of human therapeutics a number of so-called second-generation..

recombinant products are now in development (e.g. humanized monoclonal.

antibodies and modified tissue plasminogen activators) . Second-generation
peptide products include modifications of current products to alter specificity oftargets, distribution in tissues, pharmacokinetics, side effects, and stability.
These modiifications will be achieved by recombinant DNA technology
through rernoval of proteinase recognition sites and alteration of receptorbinding donnains. Analogues will be especially useful to disturb humanbioregulatory systems in the area of pharmacology. The pharmaceuticalindustry is using recombinant DNA techniques to prepare analogues fortesting as receptor binding inhibitors and as effectors of activity. A commercial
motive is the patentability of such products. Third-generation peptides will fusecoding sequences of different genes. In some cases it remains to be seen
whether thf3 risk of immunogenicity associated with the use of a modifed
protein is offset by improved efficiency, but in general it seems that protein
engineerin9 is destined to have a real impact on drug discovery.
In the near future, it must be possible to alter the specific activity, substratespecificity, and optimum pH of enzymes, increase the thermal stability ofproteins, and design proteins that can be used in nonaqueous solvents. In
addition to modification of existing proteins, the possibilities to design entirely
novel peptides and proteins with useful properties seem endless, since theamino acid sequences found in existing proteins represent an infinitesimal
fraction of all possible amino acid sequence.
In this th~esis, the aspect on the productivity of homologous and heterologous
gene products are discussed in detail. In chapter I, I demonstrated an exampleof the genetic design of chimeric E. coli as an overproducer of mammalianproteins. In chapter II, the other example of overproduction of membranebound protein is shown . In chapter III and chapter IV, I deal with an example of7extracellulalr protein , analysis of structure-function relationships, and theengineering of starch -processing enzymes by recombinant DNA technology .

.

.

T he aim 01r this investigation is to put into a real -world context therecombinant microorganisms used in the area of human therapeutics,
conventional food industry, and industrial processes. I discuss current
problems and how to overcome the limitations associated with the practicaluse of recombinant microorganism , as well as strategies for creating a
superm icrobe.

References
1. RECONIBINANT MICROBES FOR INDUSTRIAL AND AGRICULTURAL
APPLICATIONS. (1994) Murooka, Y. and Imanaka, T. (eds.), Marcel dekker,
New York.

2. Protein Engineering. (1992) Rees, A. R., Sternberg , M. J. E., and Wetzel , R.
(eds.), IRL PRESS, Oxford.
3. DNA cloning Vol I, II , and III. (1985) Glover, D. M. (ed .), IRL PRESS, Oxford.

8

·
'

Chapter I

Expression of a Gene for Human Metallothionein :Genetic Design of a Gene for Human Metallothionein II and Its Expression as
an Active Fusion Protein in Escherichia coli

9

Introduction
Metallothioneins (MTs) are low-molecular-weight, cystein-,rich proteins thatI 'have been found in a broad range of eukaryotic apecies and in many different
tissues (1, 2). These proteins bind a variety of heavy metals, such as Cd, Zn,
Cu, Hg, Co, Pb, Ni, Fe, Bi, Sn, Au, and Ag, with extremely high affinity throughcoordination bonds to their cysteins residues (1, 2). Mammalian MTs arepresent as two major forms, MT-I and -II, and both have been cloned (3-5). The
structure of the MT-II from rat has been determined both by X-ray
crystallography (6,7) and by NMR spectroscopy (8) .
Since MT proteins are small, MT seems to be an ideal model protein withwhich to investigate the nature and function of the binding of heavy metals toproteins. As part of our investigations of the structure and function of MTs, themonkey gene for MT-II was expressed as a native protein and MT protein was
synthesized in Escherichia coli (9). Subsequently, a variety of MT genes fromdifferent sources were cloned and expressed both directly and as fusion
proteins (10-13). However, the rates of production of these MTs in E. coli were
low compared with those of other eukaryotic gene products. The high level ofcysteine residues, which account for one third of the total amino acids in MT,
might be responsible for the low efficiency of the production of MT in E. coli.
Thus, the production of MT in E. coli also provides an excellent model system
for studies IOf the expression of SH-rich proteins.
In this report, we describe the design, expression, and production of an hMTII protein fused to f3-galactosidase. The fusion protien was formed in E. coli
and purified to homogeneity. The fused protein had the ability to bind heavy
metal ions 'when the protein was renatured after extraction from inclusion
bodies. Thus, the fusion protein has the potential to serve as a tool in theengineering of proteins that have binding affinity for the ions of many different
heavy metals.

10Materials and methods
Materials - All reagents were of the higher grade available commerciallyj'and were us€!d witjout further purification. Restriction endonucleases, T 4 DNA
ligase, T 4 polynucleotide kinase, and Klenow enzyme were purchased fromTakara Shuzo Co. Ltd. (Kyoto) or Toyobo Co. Ltd . (Osaka). DNA sequencing
was performed using the sequenase version 2.0 Kit (United States
Biochemical corporation, Cleveland, OH, USA). [a_ 32 P]-labeled dCTP (650Ci/mmol) and [109Cd]CI2 (164 mCi/mg) were obtained from ICN Biochemical
Inc. (Irvine, C;A, USA) and New Egland Nuclear (Boston, MA, USA),
respectively. Metallothionein II from horse and rabbit and pure f3galactosidase from E. coli (native fl-gal) were purchased from Sigma
Chemical Co. Ltd . (St. Louis, MO, USA).
Bacterial strains and plasmids - E. coli JM109 (14) was used in this study.
The cells w€!re cultured in LB medium that contained ampicillin (1 00 ~g/ml).
The vectors constructed in this study were derivatives of pUEX2 (15) which
was purchased from Amersham (Amersham , UK) . pUEX2 is a derivative ofpEX2 (16) and contains a thermolabile repressor, c1857, the lambda PR
promoter, and a cro-lacZ gene. It is basically the same as our previous
construct pF~K1 (17) except that the gene for galactokinase has been replacedby the gene for fl-galactosidase.
lSynthesis of the gene - DNA oligonucleotide were synthesized on a DNA
synthesizer Cyclone TM (Bio-Search, Burlington, MA, USA) and subsequently
purified by HPLC. Phosphorylation and annealing of complementary singlestranded DNA fragments were perfomed by mixing 0.2 ~mol of each strand of
DNA in a 20-~1 reaction mixture that contained 66mM Tris-HCI (pH 7.6) 6mM
MgCI2, 10 ImM dithiothreitol, 1mM adenosine triphosphate and 10 units of T4
polynucle01:ide kinase. The mixture was incubated at 37°C for 60 min, at 65°C
for 10 min and then allowed to cool at 12°C for 4h. Ligation of the fragments

11and manipulations of DNA were performed as described by Maniatis et a/.
(18) .
I 'Gene expre!ssion and preparation of inclusion bodies - rEo coli JM1 09 (100ml) harborin~J an expression vector was cultured in LB medium at 28°C. At themid-Iogarithnnic phase, the cell culture was shifted to 42°C for 1 h and thenincubated 37'oC for a further 16 h. The cells were harvested by centrifugation
at 8,000 x 9 jfor 10 min at 4°C. The cells were washed with 10 ml of 30 mM
Tris-HCI buHer (pH 8.0) that contained 30 mM NaCI and then they weresuspended in 10 ml of the same buffer.
The cell suspension was placed in an ice bath and then disrupted bysonication (rnodel W-385, Heat Systems Ultrasonics, Farmingdale, N. Y.) at 1s intervals for a total time of 10 min at 200 watts output. The supernatant wasremoved by centrigugation at 8,000 x 9 for 10 min at 4°C. The pellet waswashed twice with the same buffer and homogenized with a 2% solution of
Triton X-100 that contained 10 mM EDT A (pH B.O) by sonocation . The
homogenatE~ was incubated at 4°C for 16 h. The supernatant was removed bycentrifugation at 8,000 x 9 for 10 min at 4°C. The pellet was washed twice with
30 mM Tris··HCI buffer (pH 8.0) that contained 30 mM NaCI. Protein
concentrations were determined by using the Bio-Rad protein assa kit (19) or
measuring :absorbance at 280 nm .
Electrophoresis on an SOS-polyacrylamide gel - Sodium dodecyl sulfatepolyacrylanlide gel electrophoresis (SOS-PAGE) was carried out by themethod of Laemmli (20) . A separation gel containing 7.5% (w/v) acrylamide
was used. Samples were dissolved in sample buffer [25 mM Tris-HCI buffer(pH 6.5) that contained 1% SOS, 5% glycerol, 0.01 % bromphenol blue, and
1% 2-mercaptoethanol]. Proteins separated on the gel were stained with
Coomassi€~ Brilliant Blue (CBB) R-250. The densitometric scanning of the

CBB-stained gel was performed with a digital densitimeter (DMU-33C;
Toyokagaku, Tokyo, Japan) .

12

Blotting of proteins and metal-binding assays - Proteins that had been
separated by SOS-PAGE were transferred electrophoretically to aI 'nitrocellulose! membrane by the method of Towbin et al. (21). In order toreduce proteins on the gel after electrophoresis, the gel was incubated in 25
mM Tris, 19~~ mM glycine buffer (pH 8.3) that contained 0.1 % SOS and 5% 2mercaptoethanol at room temperature for 30 min. The gel was washed withelectrode buffer [20 % (v/v) methanol, 25 mM Tris, and 192 mM glycine (pH8.3)]. Transfjer of the proteins was carried out in a transfer chamber (Marisoru
Co. Ltd. , Tokyo) at a constant current of 120 mA for 18 h at room temperature
with the sarTle electrode buffer.
The assay of the binding of [10ged] to proteins on the nitrocellulose membrane ~~as based on the Aoki et al. (22). The membrane was soaked in
100 ml of 10 mM Tris-HCI buffer (pH 7.4) for 1 to 2 h at room tempetature, andthen it was incubated in 10 ml of 10 mM Tris-HCI buffer (pH 7.4) that contained
1 ~Cilml [109Cd]CI2, 0.1 mM ZnCI2 and 0.1 M KCI for 10 min at room
temperatur€~ . Proteins that had bound the radioactive cadmium were detectedby autoradiography.
Spot transfer estimation of the binding affinity for heavy metals of proteins Proteins, namely fused hMT-1I protein, standard metallothionein II from horse
and rabbit, and pure ~-galactosidase were spotted on a nitrocellulose
membrane and the spots were dried at room temperature. The membrane withdried proteins was incubated in a reducing solution of 25 mM Tris-HCI buffer(pH 8.3), 192 mM glycine, 0.1% SOS, and 5% 2-mercaptoethanol at room
temperature for 3 to 5 min. The membrane was incubated in 20 mM glycineHCI (pH 2.8) for 3 to 5 min and rinsed twice for 3 to 5 min with 10 mM Tris-HCI
buffer (pH 7.4) for 3 to 5 min. The membrane was subsequently soaked in 10
mM Tris-HCI buffer (pH 7.4) that contained 0.1 M KCI, 0.2 J.tCilml [109Cd]C12
and 1mM unlabeled salts of heavy metals for 30 min at room temperature. The
membrane! was rinsed three times with 10 mM Tris-HCI buffer (pH 7.4) that13contained 0.1 M KCI for 5 to 10 min and dried: excess buffer was absorbedwith filter paper, and the membrane was allowed to dry at room temperature .

.
'In order to visualize the transferred protein, the nitrocellulose membrane wassoaked in a 0.1 % solution of amido black in 30% (v/v) methanol and 10%
acetic acid for several minutes and destained with 50% methanol and 10%
acetic acid. Proteins that had bound radioactive cadmium were detected byautoradiography.
Autoradiography - The membrane was completely dried with a hair drier
and exposed to X-ray film (X-Omat AR; Kodak) under an intensifying screen at-80°C for 1E; to 48 h.

1 ·1

Results
Design of the synthetic gene for hMT -II - The hMT -II protein consists of 61j''amino acid nesidues of which 20 are cysteine residues. Of,the 20 cysteine
residues, 18 are encoded by the UGC codon (4, 23). Because of thisunusually large number of UGC codons for cysteine, suitable tailoring of asynthetic gene for hMT might be expected to increase its translation in E. coli.
(17). Thus, I designed a gene for hMT -II in which an equal number of the twocodons, UGC and UGU, for cysteine were present separately, since twocodons, UGC and UGU, were used equally in E. coli. Then I synthesized a
190-bp oligonucleotide for hMT -II. The DNA sequence for the synthesis was
based on the amino acid sequence of human metallothionein (24) and thenucleotide sequence of human metallothionein II (4, 23). The design of the
primary nucleotide sequence also adhered to the criteria suggested by Itakura
et al. (25): first, amino acid codons known to be favored in E. coli were used;second, the fragments were designed to eliminate undesirable inter- andintramolecular pairing; and third, GC-rich followed by AT-rich sequences were
avoided. I designed the gene with the following additional features: I, in orderto generate the hMT -II insert from the plasmid vector, the gene included
suitable sitE~s for restriction endonucleases; II, the initiation codon AUG wasplaced adjacent to the natural NH2-terminus of hMT-II; and "I, two nonsensecodons (U(3A, UAG) were added just after the C-terminal codon. The final
construct of hMT -II after annealing is shown in Fig. 1. The sequence of thesynthetic gene was confirmed by nucleotide sequening using a plasmidbased prirner.
Since the~ accumulation of SH-rich protein, namely, hMT -II, might inhibit cell
growth, thE~ temperature-controlled expression vector pUEX2 was used (15),which includes the gene for the lambda cl857 repressor, that allows inductionof cro-lacZ gene from the lambda PR promoter upon a shift in temperature.
The hMT-11 gene was inserted 27 bp upstream from the C-terminus of lacZ~ hMT-1l

~-gal

I
.
~Iea~age
EFPDKMDP

by formic acid

NCSCAAG DSCTC~- GAATTCCC CGATAAAATGGATCCGAACTGCAGCTGTGCAGCTGGTGACTCCTGCACCTGT--CTTAAGGGGCTt~11 I IACCTAGGCTTGACGTCGACACGTCGACCACTGAGGACGTGGACA
5)'Bam HIPst I
A G S C K C K E C K~C T S C K
GCCGGCTCTIGCAAATGTAAAGAATGCAAATGTACTAGTIGCAAGCGGCCGAGAACGTTTACATTTCTIACGTTTACATGATCAACGTIC®

Nael

K S C C S C C P V G~C A K C A
AAAAGCTGCTGTTCCTGCTGTCCGGTGGGCTGCGCAAAATGTGCGI I I I CGACGACAAGGACGACAGGCCACCCGACGCG I I I I ACACGC71

QGCCKGASDKCSCC~CAGGGCTGCATCTGTAAAGGCGCTAGCGACAAGTGCAGCTGTIGC
GTCCCGACGTAGACATTTCCGCGATCGCTGTICACGTCGACAACG--~

A END END
GCTTGATAGGATCCTCTAGAGCGAACTATCCTAGGAGATCTCAGCTBam HI

SailpZMTOOl
6.90 kb

lac Z

SaIl
Sma I

Fig. 1. Construction of the pZMT001 expression vector. Eight synthetic oligonucleotides
were synthesized corresponding to both strands of the gene for human matallothionein II.
After annealing, the monomer was isolated, subcloned into pUC vector, and then inserted
into the Sma I-SaIl site of pUEX2 by ligation to generate a 6.9-kb plasmid (pZMT001).

16

and, as a result, we expected to synthesize hMT-11 as a f3-gal-hMT-11 fusion.

protein. The Iresultant plasmid was named pZMT001 and w~s used to'transform E. coli JM109 (~(lac-proAB)/F'[/acZ~M15], ApS) to Apr.
Expression of the recombinat hMT -II - E. coli cells cerying plasmid pUEX2
or pZMT001 were incubated at 28°C. At the midlogarithmic phase of growth,
the temperature was shifted to 42°C to inactivate the cl857 repressor and thenkept at 37°C for 16 h. The extracts were analyzed by SOS-PAGE. A band of a
122-kOa pr01tein was observed in the analysis of the cell extract from E. coli
JM109 that carried pZMT001 after SOS-PAGE (Fig. 2a, lane 3) . This protein is
the same size as predicted from the sum of the 116 kOa of f3-galactosidase
and the 6 kDa of hMT-11. The induced cells carrying the plasmid vector pUEX2
only accumulated a 116 kOa protein, the size of which corresponds to that of
f3-galactosidase (Fig. 2a, lane 2). Without either plasmid, neither of these
proteins was formed (Fig. 2a, lane 1).
Densitometric analysis revealed that the ~-gal-hMT-11 fusion protein
accouned for more than 24% of the total protein in the cell extract (Fig. 2c, V).
This value indicates that the recombinant protien is synthesized at a rate ofabout 2 gil culture broth. The fusion protein was present in an insoluble formand could be purifed more than 95% by washing the inclusion bodies with asolution of Triton X-100 only (Fig. 2b). A minor band of a 220-kOa protein wasobserved in the analysis of the crude extract and the purified sample from E.

coli JM 109 that carried pZMT001, but not in the extract of cells carrying
pUEX2. This band may be a dimer of the fusion protien.
Binding of cadmium to the fusion protein - To test the ability of the (3-galhMT-11 fusion protien (95% pure) to bind heavy metals, a blotting method wasused (22). F~adioactive [109Cd] was used to demonstrate the binding by the hMT -/I moiet)f of the fused protein of heavy metals, since it is known that hMT-/I
can bind Cdl 2 + ions (26). After SDS-PAGE (Fig. 3a), the protein wastransferred electrophoretically to a nitrocellulose membrane. The membrane

17

I '

b

a
kDa
212170116 76-

.

-•~y~
-. ..:-.......

~kDa
1 - ~-gal-hMT-11..- ~-gal..- dimer

212 170 -1 - ~-gal-hMT-11

116 -..- ~-gal

...

-.-~

53 -

M 1 2 3M 1 2 3
- - ...-~!-..:.
.- --

76 -

.

~~- .

.a _

53 -

I,

c

v

Fig. 2. SDS-pollyacrylamide gel electrophoresis of recombinant fused matallothionein.
Proteins on the gel were stained with Coomassie Brilliant Blue (CBB) after electrophoresis. (a) Crude extract; (b) samples after washing inclusion bodies with asolution of Triton X-100. Lane M, molecular mass standards: 212 kDa (myosin),
170 kDa (a-macroglobulin), 116 kDa (~-galactosidase), 76 kDa (transferrin), 53 kDa(glutamate dehydrogenase). Lane 1, E. co1iJM109; Lane 2, E. coliJM109 (pUEX2);
Lane 3, E. coli JM1 09 (pZMT001) . (c) Densitometric scanning of 640 nm of the CBBstained gel (a, lane 3) . The ~-gal-hMT-1i fusion protein (indicated by the arrow)represents 24% of the total protein in the transfomed E. coli cells.

18

j

'

a

b
2

1

kOa

..

-

212 170 116 -

1

~

Oimer
~-gal-hMT-11

•

2

kOa

~

~

-gal

212 170 116 -

~

Oimer

~

~-gal-hMT -II
~-gal

~

II

76 -

53 -

76 -

,. \

tt, ""r' .

53 -Fig. 3, Identification of [109Cd 21-binding proteins in vitro. Cultures in E. coli JM1 09
containing the plasmid were grown in the LB medium and the purified proteins wereused for assays (see Materials and methods) . (a) Each protein, 10 mg, was subjectedto SOS-PAGE and the gel was stained with Coomassie Brilliant Blue. (b) An autoradiograph of the nitrocellulose membrane to which proteins were transferred after electrophoresis. A dar.< spot near the ~-gal was an artificial contaminant of radioactive Cd .
Molecular mass standards were the same as those described in the legend of Fig . 2.
Lane 1, E. coli JIM1 09 (pUEX2) . Lane 2, E. coli JM1 09 (pZMT001) .

19

was incubated in a buffer that contained 0.05 JlM [109Cd]CI2 and 0.1 mM
ZnCI2. Addition of Zn 2+ ions into the [109Cd]-containing buffer was found toj'be essential for detection of hMT by this technique because it reduced thenonspecific binding of [109Cd]. When proteins on the nitrocellulose
membrane we~re labeled with [109Cd] in a buffer without divalent metal ions,[109Cd] bound to proteins on the membrane and nonspecific binding of[109Cd] to the membrane caused significant interference in detecting MT (22) .
Autoradiography showed that a major radioactive band of a 122-kDa protein
corresponded to f3-gal-hMT-11 (Fig. 3b, lane 2). A minor band of a protein of
about 220 kDa was also labeled with [109Cd] , which suggested that it was thedimeric form of the f3-gal-hMT-11 fusion protein (Fig. 3, lane 2). However, noradioactive band was observed in the analysis of the extract of cells thatcarried pUX2 . Since the reduction of proteins on the gel was essential for
generation of reproducible results in terms of the radioactive bands of thefusion protien, endogenous Cd 2+ ions were probably removed from hMT-1I
during the reduction process (22) . These results indicate that Cd 2+ ions canbind to the fused hMT -II protein. The fusion protein seemed to be at least
partially renatured and was, thus able to bind Cd 2+ ions.
In order to produce hMT-11 from the fusion protein, the fused protein has to becleaved to remove the f3-galactosidase moiety. Formic acid, acting at theaspartyl-proline bond, has been used for the production of hMT -II (27) .
Samples (2 ~tg of f3-gal-hMT -II and recombinant f3-galactosidase) treated with
70% formic acid and 6M guanidine at 37°C for 48 h were subjected to SDSPAGE, and the metal-binding assay with [109Cd] was performed. Weak andsmeared spots of [109Cd]-binding protein that corresponded to a protein of
about 17 kDa from f3-gal-hMT -II were observed, but no radioactive spot wasobserved from recombinant f3-galactosidase (data not shown) .
Affinity for heavy metal ions by the fusion protein - To determine the extent
of renaturation of the fusion protein, standard metallothionein and the purified

20

~-gal-hMT-11 protein that was 95%

pure were diluted and spotted on two

nitrocellulose membranes. One part of the membrane was stained with amido
4 •black (Fig. 4a) and the other was subjected to labeling with [109Cd] (Fig. 4b).
About 40 J,lg of fl-gal-hMT -II protein (25-fold dilution of about 1000 J,lg ofprotein) were! labeled with [10gCd]. The intensity of the band corresponded tothe intensity of a band ot2 J.tg of standard MT-II from rabbit of horse (25-folddilution of about 10 J,lg of protein) (Fig. 4). These results indicate that, in termsof binding of [10gCd], one mole of ~-gal-hMT -II is equivalent to one mole ofstandard MT -II which binds 7 atoms of Cd. fl-Galactosidase (Sigma) andrecombinant fl-galactosidase from cells that carried pUEX2 did not bind any
radioactivity. Since the percentage of renaturation of the fusion protein was
not known, it was impossible to quantify the specific activity determined in thepresence of Cd 2+ ions.
The affinity of the fused hMT -II protein for heavy metals was qualified. The
membranes spotted with samples of diluted proteins were incubated with 0.01
J.tM [109Cd]C I2 in the presence of various heavy metals ions at 1 mM.
Darkening of the Cd-binding protein after addition of 1 mM Zn 2+ ions to the[109Cd]-containing buffer was observed, but not after a similar addition of 1
mM Cu 2+ or C02+ ions (Fig. 4d-f). The fused hMT -II had a higher affinity for
Cd 2+ ions than for Zn 2+ ions, as did the standard horse MT -II protein (Fig. 4d
and h). By constrast, both the fused hMT-1I and horse MT-II failed to bind Cd 2+ions in the presence of 1 mM of Cu 2+, C02+, or Ag+ ions (Fig. 4e-g). These
results illustrate that the affinities of hMT -II for Cu 2+, C02+ and Ag+ ions arehigher than those for Cd 2+ and Zn 2+ ions, as is the case for horse MT -II (28,29). We observed small black spots of fl-gal-hMT-1I and ~-gal, and highbackground in the presence of Ag+ ions. Ag+ ions do not seem to inhibit nonspecific binding of [10gCd] to proteins and nitrocellulose membrane that is
affected by divalent metal ions (22).

21

x 1 115 1/25 1/125 11625

a~-gal (200 Jl9) ~-gal-IhMT-1l(1000 Jl9) ,~.-rabbit MT-II (10 ~g) horse MT-II (10 Jl9) BSA (10 Jl9) -native ~-gal (10 Jl9) x 1 115 1/25 11125 11625

b~-gal (200 Jl9) -~-gal-'hMT-1l (1 000 ~g)-rabbit MT-II (10 Jl9)horse MT-II (10 Jl9) -

e. ·

••

• •

BSA (10 ~g)­native ~-gal (10 ~g) -chorse MT-II~-gal-hMT- 11

f3-gal

(1 0 ~)(200 1-lQ)(200 1-lQ)

f 1 mM CoCI 2d

1 mM ZnCI 2

9

1 mM AgN03••h

1 mM CdCI 2•

Fig. 4. Effects of heavy metal ions on binding of [109Cd 2i ions to the hMT-11 fusion protein.
The purified fusion protein was dotted on a nitrocellulose membrane as described inMaterials and mE~thods. The membrane was incubated in a solution of 0.2 mCi I ml
[109Cd2+] in the presence of heavy metal ions and then rinsed with 10 mM Tris-HCI buffer.
Panels (a) and (c), protein stained with amido black; panels (b) and (d)-(h), autoradiographafter 16 hand 48 h, respectively. The membrane was labeled with [109Cd 2i in buffer that
contained 1 mM ZnCI 2 (b, d), 1mM CuS04-5H20 (e), 1 mM CoCI 2 (f), 1 mM AgNOa (g), or

1 mM CdCI 2 (h).

22From these results, it can be concluded that hMT -II functions within the fusedprotein that includes f3-galactosidase to bind heavy metal ions the samemanner as standard equine MT -II.

Discussion
The rate of production of gene products that contain large numbers of 8H~.

.

groups in E. coli is often limited because of the toxicity of such proteins to thebacterial cells and the stability of the proteins themselves which sometimes failto undergo the correct folding. In the previous experiments, Murooka et al.
succeeded in the direct expression of the cDNA for monkey MT -II in E. coli (9).
However, thE! yield of protein was less than a few mg per liter of cultured cells.
Several efforts to overproduce MT proteins in E. coli and Saccharomyces
cerevisiae w1ere subsequently made, but no high level of expression of MTsfrom various sources has been reported (10-12, 30). Thus, the production of
MTs in fused form with other proteins has been attempted (13, 31, 32) . The
gene coding for the copper metallothionein of Neurospora crassa wasexpressed as a fusion product at the NH2-terminus of ~-galactosidase and thefused protein was produced as a soluble from. However, the yield of the fusedprotein was 16 times lower than that of the non-fused ~-galactosidase (32).
Moreover, no protein of the size expected for the fused protein was observed
after 80S-PAGE.
Romeyer €~t al. synthesized DNA that encoded for human hepatic
metallothionein and they expressed the gene as a fusion protein with Ara'S.
The fusion protein accounted for approximately 8% of the total protein in E.
coli. The Ara'8-hMT fusion protein produced by E. coli had a half-life of

approximately 50 min (2, 13). Jacobs et al. cloned the gene for hMT in an
expression vector, pUA7, including the promoter-operator and regulatory
sequences of the ara operon of Salmonella typhimurium and part of the 5'coding and all of the 3'-noncoding regions of the Jpp from E. coli. The fusion
protein was produced as a stable protein. The ability of the fused protein tobind heavy metals was characterized, but it was not demonstrated that thehMT -II mOi1ety of the fused protein was renatured to the same degree as thenative MT (31) .

2 ·1

In the present study, a synthetic DNA encording hMT -II was designed andfused with the ~~ene for a large protein (tl-galactosidase). Moreover, thej'•overproduction of the hMT -II fused protein in E. coli was successfully
achieved. Upon heat induction of the c1857-PR promoter system, the fusion
protein was produced to a level equal to more than 24% of total cellular
protein. Since 1the hMT -II gene contains an usually large number of UGC
codons, for 18 of a total of 20 cysteine residues it was postulated that thesubstitution of codons suitable for expression in E. coli might increase thetranslation of the gene in E. coli. However, the stability of the hMT-11 protein
was so high after fusion with f3-galactosidase that the codon usage was notevaluated. NO\N we are examining the direct expression of hMT -II toinvestigate the! effects of codon changes. The fused hMT -II was stabilized as
an insoluble protein that formed inclusion bodies in E. coli cells. The insoluble
p-gal-hMT -II protein was easily solubilized by incubation treatment with SDS
and 2-mercaptoethanol which genetated the reduced form . Some of the
reduced f3-gal-hMT -II protein formed dimers on SOS-PAGE, as does native
MT (22). The 1fused hMT -II protein was easily detected with standard proteinspecific dyes or by labeling with [10gCd], whereas the native MT obtained bydirect expression is attacked by proteases and has been difficult to detect after
SOS-PAGE because of its small size. Furthermore, we clearly demonstrated
that the hMT -II mOiety of the fusion protein is functional in the same hierarchy
with respect to binding of heavy metals ions (Cu 2+:> Cd 2+> Zn 2+) as native
hMT -II from the horse (33). We also succeeded in regeneration of hMT -II fromthe fusion protein, and the regenerated hMT -II protein migrated in a gel withthe same mobility as that of rabbit liver MT and horse kidney MT (34), andbound the radioactive cadmium (unpublished results). We are presently
studing how l11any atoms of heavy metals the fusion protein binds and whether
the fusion protein binds heavy metals in solution.

25

The present rE~sults suggest that any barriers to the production of a proteinwith many SH ~Jroups in E. coli may not involve the inhibitory effect of any
I'•specific structural feature of the protein, but may rather involve the localization
or stability of the protein within the cells (13, 31). With increased
understanding of structure-function relationships, we should be able to create
novel forms of MT that have an altered specific affinity for given metal ions.

26

Summary
A synthetic DNA coding for human metallothionein II (hMT-II) was designedj'•for efficient expression in Escherichia coli and cloned intora vector, pUEX2.
Upon induction by heating, the gene for hMT-11 was expressed as a fusion
protein accounted for approximately 24% of the total protein in the cells andaccumulated in the cells as an insoluble form . The fused hMT -II protein with amolecular mass of approximately 122 kOa was purified to homogeneity by
washing the inclusion bodies with a solution of TritonX-100. The insoluble
protein was easily solubilized in the presence of 2-mercaptoethanol andsodium dodecyl sulfate without any need for denaturants, such as urea or
guanidine hydrochloride. The fused hMT -II protein bound to radioactive
cadmium . A competition experiment with radioactive cadmium and various
heavy metals showed that the hMT -II fusion protein had a higher affinity for
Cu 2+, C02+ , Ag+ ion than Cd 2+ and Zn 2+ ions. Thus, we secceeded in overexpression of hMT -II as an active form in E. coli.

27

References
1.

Harmer, D. H.: Metallothionein. Annu. Rev. Biochem., 55, 913-951
I '(1986).
2.

Kagi, J. H. R. and Schaffer, A.: Biochemistry of metallothionein.
Biochemistry, 27, 8509-8515 (1988).

3.

Durham , D. M., Perrin, F., Gannon, F., and Palmiter, R. D.: Isolation andcharacterization of mouse metallothionein-I gene. Proc. Natl. Acad. Sci.
USA, 77, 6511-6515 (1980).

4.

Karin, M. and Richards, R. I.: Human metallothionein genes: molecularcloning and sequence analysis of mRNA. Nucleic Acid Res., 10,31653173 (19182).

5.

Schmidt, C. J. and Hamer, D. H.: Cloning and sequence analysis of twomonkey metallothionein cDNAs. Gene, 24, 137-146 (1983) .

6.

Furey, W'. F., Robbins, A. H., Clancy, L. L., Winge, D. R., Wang, B. C., andStout, C. D.: Crystal structure of Cd , Zn metallothionein. Since, 231, 704710 (19B6).

7.

Robbins, A. H., McRee, D. E., Williamson , M., Collett, S. A., Xoung, N. H.,
Furey, \/II. F., Wang, B. C., and Stout, C. D.: Refined crystal structure ofoCd, Zn rnetallothionein at 2.0 A resolution . J. Mol. BioI., 221, 1269-1293(1991 ).
8.

SchultzE!, P., Worgotter, E., Braun, E., Wagner, G., Vasak, M., Kagi, J. H.
R., and Wuthrich, K.: Conformation of Cd7-metallothionein-11 from rat liverin aqueous solution determined by n.m.r. spectroscopy. J . Mol. BioI.,
203,251-268 (1988).

9.

Murooka, Y. and Nagaoka, T.: Expression of cloned monkey
metallothionein in Escherichia coli. Appl. Environ. Microbiol., 53, 204-207(1987).

10.

Berka, T., Shatztman, A., Zimmerman, J., Strickler, J., and Rosenberg,
M.: Efficient expression of yeast metallothionein gene in Escherichia

28coli. J. Bacteriol., 170,21-26 (1988).

11.

Hou, Y. tvt, Kim, R., and Kim, S. H.: Expression of the mouseI 'metallothionein-I gene in Escherichia coli: incresed tolerance to heavy
metals. Biochem. Biophys. Acta, 951,230-234 (1988).
12.

Kille, P., Stephens, P., Cryer, A., and Kay, J.: The expression of asynthetic rainbow trout metallothionein gene. Biochim. Biophys. Acta,
1048, 1'l8-186 (1990).

13.

Romeyer, F. R., Jacobs, F. A., Masson, L., Hanna, Z., and Brousseau, R.:
Bioaccul11ulation of heavy metals in Escherichia coli expressing an
induciblE~ synthetic human metallothionein gene. J. Biotechnol., 8, 207-

220 (19B8).
14.

Yanish-Perron, C., Vierra, J., and Messing, J.: Improved M13 phagecloning vectors and host strains: necleotide sequences of M13mp18
and pUC19 vectors. Gene, 33, 103-119 (1985).

15.

Bressan, G., and Stanley, K.: pUEX, a bacterial expression vector
related to pEX with universal host specificity. Nucleic Acid Res., 15,10056 ('1987).

16.

Keith, K. S. and Luzio, J. P.: Conctruction of a new family of highefficiency bacterial expression vector: identification of cDNA clones
coding for human liver protein. EMBO J., 3, 1429-1434 (1984).

17.

Murooka, Y. and Mitani, I.: Efficient expression of a promoter-controlled
gene: tandem promoters of lambda PR and PL functional in entericbacteria. J. Biotechnol., 2, 303-316 (1985).

18.

Maniatis, T., Fritsch, E. F., and Sambrook, J.: Molecular cloning: A
Laboratory manual. Cold Spring Harbor Laboratory, Cold Spring
Harbor, New York (1982).

19.

Bradford, M. M.: A rapid and sensitive method for quantitation of
micro~Jram quantities of protein utilizing the principle of protein-dye

binding. Anal. Biochem., 72, 248-254 (1976).

29

20.

Laemmli, U. K.: Cleavage of structural proteins during assembly of thehead of bacteriophage T 4. Nature, 224, 680-685 (1970).
j'

21.

,

Towbin, H., Staehelin, T., and Gordon, J.: Electroph(J)retic transfer ofproteins from polyacrylamide gels to nitrocellulose sheets: procedureand some applications. Proc. Natl. Acad. USA, 76, 4350-4354 (1979).

22.

Aoki, Y., Kumamoto, M., Shibata, Y., and Suzuki, K. T.: Detection of
metallotlhionein on nitrocellulose membrane using western blotting
technique and its application to identification of cadmium-binding
proteins. Anal. Biochem., 157, 117-122 (1986).

23.

Karin, M. and Richards, R. I.: Human metallothionein genes primarystructurE~ of the metallothionein-II gene and a related processed gene.

Nature, 299, 797-802 (1982).

24.

Kissling, M. M. and Kagi, J. H. R.: Primary structure of human hepatic
metallothionein. FEBS Lett. 82, 247-250 (1977).

25.

Itakura, 1<., Hirose, T., Crea, R., and Rigga, A. D.: Expression in

Escherichia coli of a chemically synthesized gene for the hormone
somatostatin. Science, 198, 1056-1063 (1977).

26.

Kagi, J. H. R. and Nordeberg, M.: Metallothionein. Birkhauser verlag,
Basel (1979).

27.

Marston, F. A. 0.: The purification of eukaryotic polypeptides expressedin Escherichia coli, p. 59-88. In Glove,D. M. (ed.), DNA cloning, vol. III.
IRL Press, Oxford (1987).

28.

Rupp, H. and Weser, U.: Circular dichroism of metallothioneins. Biochim.
Biophy . Acta, 533, 209-226 (1978).

29.

Suzuki, K. T., Yamamura, M., and Maitani, M.: Chromatography ofmetallothioneins on a gel permeation column: changes induced byreplacE~ment of cadmuim with cuprous and cupric ions. J. Liq.

Chromatogr., 3, 1386-1397 (1980).

30.

Mbikay, M., Brzezinski, R., and Thirion J. P.: Differential expression of30

cloned rnouse metallothionein sequences in Escherichia coli. DNA, 2,
23-30 (1 B83) .

.
'

31.

Jacobs, F. A., Romeyer, F. M., Beauchemin, M., and Brousseu, R.:
Human rnetallothioneinll is synthesized as a stable membrane-localized
fusion protein in Escherichia coli. Gene, 83, 95-113 (1989).

32.

Sugimoto, M., Oikawa, T., Esaki, N., Tanaka, H., and Soda, K.: Chemical
synthesis and expression of copper metallothionein gene of

Neurospora crassa. J. Biochem., 104, 924-926 (1988) .
33.

Suzuki, K. T. and Maitani, T. : Metal-dependent properties ofmetallothionein. Biochem. J., 199, 289-295 (1981) .

34.

Andersen, R. A. and Daae, H. L.: Prepration of metallothionein from rat
liver and studies of its properties with respect to use as a standard in gel
permeation chromatography, polyacrylamide gel systems,
autoradiography and western blotting. Compo Biochem. Physiol. 90B,
59-67 (1 !388).

31

I"

Chapter II

Production and Purification of Monoamine Oxidase :Production, Purification, and Characterization of a new type of Monoamine
Oxidase containing Cu from Klebsiella aerogenes

32Introduction
Monoamine oxidases catalyze the oxidative deamination of monoamines
I '0according to the following reaction:
RCH2NH2 + 02 + H20 --------> RCHO + NH3 + H202
Monoamine oxidase from mitochondria of animal organs (1-5) and plasma (69), from pea seedings (10), from various microorganisms (11, 12), and the cell
membranes of bacteria (13) has been characterized . The enzyme has a broad
substrate specificity and palys a major role in the metabolism of biogenic
amines, such as the neurotransmitters of the central nervous system.
The monoamine oxidase in Klebsiella aerogenes is induced by tyramine andcatecholamines, and it is specific for tyramine and dopamine (14) . Thus, theenzyme from K. aerogenes W70 has been classified as a tyramine oxidase(EC 1.4.3.9) (15). Tyramine oxidase in K aerogenes is encoded by the tynA
gene, and the synthesis of the enzyme is subjected to catabolite andammonium repression (14, 16). Expression of the tynA gene results inderepression of the ats operon, which is closely linked to tynA (17, 18) and iscomposed of the arylsulphatase (atsA) gene and the positive regulator, The

atsB gene (19).
Since the tyramine oxidase from K. aerogenes binds tightly to the cell
membrane and has not been solubilized in an active form (14),characterization of the enzyme and determination of the mechanism of
monoamine oxidization remain to be performed. These studies will require thepure enzyme. During the course of our studies on the genetic regulation of thesynthesis of the tyramine oxidase, we cloned the structural gene for the
tyramine oxidase of K. aerogenes W70 into a multicopy plasmid. This cloning
resulted in overproduction of the enzyme in a soluble form (20), thus
rendering possible both the purification and the characterization of the
enzyme. We found that the pure enzyme was highly specific for dopamine, f3phenylethylamine and tryptamine, in addition to tyramine and contained33

copper but not FAD as a prosthetic group. From these results, we classify the
enzyme from K aerogenes as a monoamine oxidase (EC 1.4.3.6) instead of a:.
~tyramine oxidase (EC 1.4.3.9), and hence the gene encoding this enzyme wasrenamed from tynA to maoA (20).

31Materials and methods
Materials - All reagents were of the highest commercially available gradeI 'and were used without further purification. Horseradish peroxidase (Grade \1)
was purchased from Boehringer Mannheim Yamanouchi Co. Ltd. (Tokyo).
DEAE-Toyopearl 650S, DEAE-Toyopearl 650M, Butyl-Toyopearl 650M , andToyopearl HW55S were purchased from Tosoh Co. Ltd . (Tokyo) . Iproniazid
phosphate salt and tyramine hydrochloride were purchased from Sigma
Chemical Co . Ltd . (St. Louis, MO) . Clorgyline hydrochloride and deprenylhydrochloride were purchased from Research Biochemicals Inc. (Natick, MA) .
[3H]-Tyramine hydrochloride (20 Ci/mmol) was purchased from New England
Nuclear Research/DuPont (Boston, MA).
Bacterial strains and plasmids - The bacterial strain used in this study was
K. aerogenes WlO (prototroph)(21) , which produces monoamine oxidase(14). pT058, a plasmid containing a gene for monoamine oxidase andconferring kanamycin resistance (Kmr) was constructed previously (20) .
Medium and culture conditions - K. aerogenes WlO harboring pT058 was
subcultured in 500 ml of K medium (22) in a 5-liter flask. Xylose (0 .5%) and
1mM NH4CI were used as sources of carbon and nitrogen, respectively.
Tyramine (1 mM) was used as inducer of monoamine oxidase (14) .
Kanamycin (50 f,tg/ml) was used in precultures. The subculture was inoculated
into a jar fermenter that contained 30-liter of K medium. The incubation was
carried out at 28°C for 16 h with aeration. The cells were harvested bycentrifugation at 4,000 x 9 for 30 min, washed with 0.1 M potassium
phosphate buffer (pH 7.4), and frozen at -80°C.
Assay of enzymatic activity and quantitation of protein - Monoamineoxidase activity was determined by a colorimetric assay that was based on acoupled oxidation reaction, with measuremant of the monoamine oxidasedependent formation of hydrogen peroxide. The method was essentially thesame as that employed for assays of amine oxidase in fungi (11) but tyramineand o-dianisidine were used as substrate and indicator, respectively. The
assay mixture was composed of 2.6 ml of 0.1 M citrate-phosphate buffer (pHI '0

7.4) that contained 0.6 mM o-dianisidine, 0.02% (w/v) horse radish
peroxidase, and 5 mM tyramine. No inhibition of the activity of the monoamineoxidase from Klebsiella by o-dianisidine was observed at concentrations up toat least 1.2 mM . The mixture was allowed to stand at 30°C for 5 min and thereaction was started by the addition of O. 1 ml of enzyme solution andincubated at the same temperature for 20 min. The reaction was stopped by
the addition of 0.3 ml of 1 M citric acid, and the mixture was centrifuged when
the preparation of enzyme became impure. The absorbance at 435 nm wasrecorded on a Beckman DU64 spectrometer against a reagent blank without
addition of enzyme solution. One unit of enzymatic activity was defined as theamount that catalyzed the formation of 1 f..lmol of H202 in 1 min at 30°C, which
was caluculated using a molar absorption coefficient of 2230 M-1 cm- 1.
Monoamine oxidase was also assayed by a radiometric procedure with [3H]tyramine (14) . Protein content was quantitated by the method of Lowry et a/.
(23) with bovine serum albumin as a standard. The elution of proteins wasmonitored by measuring absorbance at 280 nm .
Purification of the enzyme - All the purification steps were carried out at
4°C.
Step 1. Preparation of crude extract. The cells (about 177 g wet wt/20 I
culture) of K. aerogenes W70 harboring pT058 were disrupted by sonication.
Intact cells and debris were removed by centrifugation and the supernatant
was used as the crude enzyme solution.
Step 2. Ammonium sulfate fractionation. The precipitate obtained from thesupernatant between 35% to 65% saturation of ammonium sulfate wasdissolved in 50 mM potassium phosphate buffer (pH 7.4), and dialyzed
against the same buffer.

36

Step 3. Column chromatography on DEAE-Toyopearl 650S. The dialyzed
solution (120 ml) was applied to a column (4.4 cm i.d. x 18 cm) of DEAEI '•Toyopearl 650S which had been equilibrated with 50 mM potassium
phosphate buffer (pH 7.4) and eluted with a gradient of NaCI from 0 to 0.5 M inthe same buffer (pH 7.4) . The active fractions were pooled and concentrated
by the addition of solid ammonium sulfate.
Step 4. Column chromatography on Butyl-Toyopearl 650M. The precipitate
obtained by centrifugation was dissolved in 50 mM potassium phosphatebuffer (pH 7.4) and dialyzed against the same buffer. Solid ammonium sulfate
was added to the dialysate to 35% saturation. The concentrated solution of
enzyme (45 ml) was applied to a column (2.3 cm i.d. x 24.5 cm) of ButylToyopearl 650M which had been equilibrated with 0.1 M potassium
phosphate buffer (pH 7.4) that contained solid ammonium sulfate to 35%
saturation. The enzyme was eluted with a gradient of ammonium sulfate from
35% to 0% in the same buffer. The active fractions were pooled andconcentrated by the addition of solid ammonium sulfate.
Step 5. Column chromatography on DEAE-Toyopearl 650M . The precipitate
obtained by centrifugation was dissolved in 50 mM potassium phosphatebuffer (pH 7.4) and dialyzed against the same buffer. The dialyzed enzyme
solution (23 ml) was applied to a column (1 .9 cm i.d . x 35 cm) of DEAEToyopearl 650M which had been equilibrated with same buffer and eluted
with a gradient of NaCI from 0 to 0.5 M in the same buffer. The active fractions
were pooled and concentrated by the addition of solid ammonium sulfate.
Step 6. Gel filtration . The precipitate obtained by centrifugation was
dissolved in 50 mM potassium phosphate buffer (pH 7.4) . The enzyme
solution (1 ml) was loaded to a column (2.6 cm i.d. x 96 cm) of Toyopearl
HW55S which had been equilibrated with same buffer and eluted with same
buffer. The active fractions were collected and used for the characterization of
the enzyme.

3Electrophoresis - SOS-polyacrylamide gel electrophoresis (SOS-PAGE ;7.5% acrylamide) was carried out according to the method of Laemmli (24),.

:1

and the bands of protein were stained with Coomassie Bnilliant Blue R-250.
Molecular weight determination - The molecular weight of the purified
enzyme was determined by gel-filtration on a column (2.6 cm i.d . x 96 cm) of
Toyopearl HW55S by the method of Andrew (25) and SOS-PAGE.
Spectrophotometric determination - Enzymatic reaction and protein
concentrations were measured in a Beckman OU64 spectrophotometer. The
absorption spectra were recorded with an EPSON sp-500 recording
spectrophotom eter.
Metal determination - The matal content of the enzyme was determined
spectrophotometrically. The final sample of enzyme was prepared with metalfree distilled water and reagents. The enzyme was dialyzed for 24 h against
0.01 M potassium phosphate buffer (pH 7.0) that contained 0.1 mM
dithiothreitol and 1 mM EDT A. The dialyzed enzyme at a concentration of 11.5 mg of protein per ml was analyzed with a Nippon Jarrell-Ash Model AA845 atomic absorption spectrophotometer.
Computer analysis - The amino acid sequence were analyzed with
GENETYX programs (SOC Software Development Co., Ltd., Tokyo, Japan) .

Results
Purification of monoamine oxidase from K aerogenes W70 (pT058) - The.'.

overall purification procedure is summarized in Table 1. "fter ammonium
sulfate fractionation and chromatography on a series of columns, the enzyme
was purified 10-fold with a yield of 9.1 %. The purity of the enzyme wasanalyzed by 80S-PAGE ana the final preparation of enzyme was found to behomogeneous (Fig. 1, lane 2) .
Molecular weight - The molecular weight of monoam ine oxidase wasdetermined by gel filtration on a column of Toyopearl HW55S and S08PAGE. The enzyme gave a single peak and was eluted at a position near thatof bovine serum albmin (68,000 Oa). Its molecular weight was estimated to be
79,000 (Fig. 2). The purified enzyme gave a single band that corresponded toa molecular weight of 79,000 on SOS-PAGE (Fig. 1, lane 2). Since the value
is agree with that deduced from the predicted amino acid sequence (26) , theenzyme appears though to consist of a single polypeptide chain.
Effects of pH - Enzyme activity was determined in buffers of different pHs.
The enzyme showed maximum activity at pH 6.0 when examined in thepresence of 0.1 M potassium phosphate buffer, and 50% of the maximum
activity was obtained at pH 5.5 and pH 7.0 . The enzyme was stable at pH 4.0,
but lower activity was recorded after a 2-h incubation at pH 10.5 and 4°C.
Effects of temperature - The effects of temperature on the activity andstability of the enzyme were examined. The optimal temperature for activity
was 50°C, and 50% of the maximum activity was observed at 35°C and 55°C.
The enzyme was stable up to 45°C but 50% of the activity was lost after a 20min incubation at 50°C.
Substrate specificity - The activity of purified monoamine oxidase was
examined in the various amines (Table 2). The enzyme exhibited high activity
when tyramine, ~-phenylethylamine, dopamine, and tryptamine were tested
as substrates; moderate activity towards serotonin and histamine. Other

39

Table 1. Purificatibn of monoamine oxidase from Klebsiella aerogenes W70 (pT058)
I

Total
activity

Tatal
protein(U)(mg)

Specific
activity(mU/mg)*

Crude extract

115.2

20160

5.7

100

Ammoniumsulfatefraction

58.9

9480

6.2

51 .5

DEAE-Toyopearl
650S

21 .5

2220

9.7

18.7

Butyl-Toyopearl
650M

14.0

468

29.9

12.2

DEAE-Toyopearl
650M

13.1

264

49.6

11.4

Toyopearl HW55S

10.5

178

59.0

9.1

Fraction* For defination of units, see text.

.0

.

1

'

2

97 kDa67 kDa43 kDa30 kDa•Fig. 1. Results of sodium dodecyl sulfate-polyacrylamide gel electropforesisof monoamine oxidase. Proteins were stained with Coomassie Brilliant Blue.
Lane 1, Molecular weight standards: phosphorylase (94 kDa); bovine serum
albumin (67 kDa); ovalbumin (43 kDa); carbonic anhydrase (30 kDa);
lane 2, fraction from Toyopearl HW55S.

41

I '

-

-..

..c 6.0
0>
'Q5
~
'-

ro

1

:J
()
Q)

0

-E

0

5.0

~

0>
0
..J

4.0

o

0.5

1.0

Relative elution volume

Fig. 2. Relation between elution volumes of proteins and their molecular weights
during gel filtration on Toyopearl HW55S. 1, Aldolase (158 kDa); 2, bovine serum
albumin (67 kOa); 3, chymotrypsinogen A (25 kOa). The symbolll shows the
position of elution of the monoamine oxidase.

~.

Table 2. Substrate specificity of monoamine oxidase from
K. aerogenes W70 (pT058) *Substrate (5 mM)

Relative
activity(%)

Tyramine

100~-Phenylethylamine

89.1
83.7
32.6
<0.1
<0.1
76.6

Dopamine
Octopamine
Adrenaline
Noradrenaline
Tryptamine
Serotonin
Histamine
Benzylamine
Amylamine
Hexylamine
Butylamine
Propylamine

7.6
5.5
<0.1
18.1
15.1
<0.1
<0.1

Substrate (5 mM)

Ethylamineo-Phenylenediami ne
Hexylenediamine
Cadaverine
Putrescine
Propylenediamine
Ethlenediamine
Spermine
Spermidine
Agmatine
L-Tyrosine
L-Histidine
L-DopaKynurenine

Relative
activity
(%)
<0.1
<0.1
<0.1
<0.1
<0.1
<0.1
<0.1
<0.1
<0.1
<0.1
<0.1
<0.1
<0.1
<0.1

* The enzymatic activity was determined under the standard assay conditions
described in Materials and methods. The reaction mixture consisted of 5 mMof a given substrate, 0.1 M citrate-phosphate buffer (pH 5.6) , 0.02% (w/v) horseradish peroxidase, 6.6 mM o-dianisidine and 13.6 mU of the purified monoamineoxidase in a total volume of 3.0 ml. Oxidation of tyramine is taken as 100% ; otheractivities are given relative to that value.

43monoamines, such as benzylamine and kynurenine, which are known as
typical substrates for animal amine oxidases (4, 7), were n~t oxidized at all.
j'

Diamines, polyamines, and amino acids were also not oxidized. Thus, thisenzyme can be classified as a monoam ine oxidase rather than a tyramineoxidase (14, 15). However, the pattern of substrate specificity of the enzyme
from K aerogenes was different from those of other monoamine oxidase (9,11, 27).
Kinetics - The reaction catalyzed by the purified enzyme exhibited
Michaelis-Menten kinetics. The Km and Vm values for tyramine were
calculated to be 0.282 mM and 0.551 (U/mg of protein), reapectively.
Effects of chemical reagents - The effects of various compounds at 1 mM on
the activity of the purified enzyme were investigated, and the results are
summarized in Table 3. Among competitive inhibitors of monoamine oxidase,clorgyline inhibited the activity of the enzyme more markedly than did
deprenyl. From this result, we classify the enzyme as an A type monoamineoxidase (28). The enzyme was also inhibited by isoniazid and iproniazid,
which are specific inhibitors of monoamine oxidase (29). Even thoughclorgyline and deprenyl are substrate analogues, they bind to coenzyme FAD
irreversibly after a long incubation with monoamine oxidase (30). Several
monoamine oxidase have been reported that contain FAD or pyrooloquinoline
quinone (PQQ) as a prosthetic group (1,2). Thus, we examined the absorption
spectra (Fig. 3A) of the purified monoamine oxidase from K aerogenes.
However, no characteristic shoulder was observed, and the enzyme exhibitedan absorption maximum at about 280 nm. The enzyme did not generate any
particular peaks specific for FAD (31,32,33) and PQQ (34), neither in its
absorption spectrum nor in its fluorescence spectrum (34, 35; data not shown).
The enzyem from K. aerogenes was markedly inhibited by carbonyl reagents(Table 3). By contrast, tyramine oxidase from Sarcine lutea (11) is notinhibited by carbonyl reagents, such as hydrazine, hydroxylamine, and44

fit .l '.:., ( )I ;: ) :~ I·llll. : 1..:.

I. " :::, i .. ' i .' i 't..'..

,.. B..

i') 'L~ I ," \".......... ' j

i I I
":

l

IO t · tI~

1: ' :", 1 '! I :

I,' ,

,

I •

! ..

! . Ii.', \ " , I•. ' 1.-. .' ~ I ~.' ., (.!

!
!
.~ :

I

.

I

· .. ',:.:,:,~. ::', ~ ., ' . l,)!,.oj,I . .

··1

,I .

I
I. . .

I

I. ..

,

_ ./ " -' ~ . ) I . ' " \)t··· .

I
.. i

1'.1

I

I

·.J:I

'. Ii) u i :,

I

ii

I

1'1

.. .... 1'1 (! I. ' " <)

.,I

I· ..
I

I

I

I ..................... 1 ,J " .::i)(,i.)

I ·I. ...... 1.......................... 1. ......•...•... I .

.I .

.. ..... I ..................... I. ..

i.> !~·.':I ,(.,,(.

() :>: ("'1""'"

I:..

._.,." r.:. ~:)I· .. ·I I)

1

II

11

' .i'' ,...... .,.' ,Ie)'.'" . .

('j.. ..... .

I

F

A:.

I. .

.'/
."

"

1

I
. ...

I

1

I

I~

1

..

i. .......................... .1 .... ..................... ::.:::1::: ..""......................i.

U" C:IUUU. 1. ........................... .1 ..... .
():I~.:.:.:Ii·l·() (:'. I. ..... .
.'I.,..'" , ' ,'

.L.
,',

",

.:',C,t) l )...... ..... .... j ........ .

...... I ......... .

t.: ) ... J..Ui)(1II

I.::' til.> " I ....

() " ,:: ,() .:)C'

Fig . 3. Absorbance spectra of the monoamine oxidase from K. a erogenes.
Purified enzyme with a specific activity of 59 mU/mg was used at concentrationsof 1.5 (mg/ml) (A) and 259 (mg/ml) (8) in 50 mM potassium phosphate buffer(pH 7.4) . Reduced form I (Red. I) was obtained by the addition of tyramine (final
concentration 0.1 m~) to the purified enzyme under anaerobic conditions.
Reduced from" (R~d . II) was obtained by the addition of sodium d,tionite(final concentration 16 mg/ml) to the purified enzyme under anaerobic conditions.

45

t'l

Table 3. Effects of various reagents on the monoamine oxidasefrom K. aerogenes W70 (pT058)*

Reagent (1 mM)

Clorgyline
Deprenyl
Isoniazid
Iproniazid
1-Ethyl-3-(3-dimethylaminopropyl)-carbodiimide
Aminoguanidine
Hydrazine
Hydroxylamine
Semicarbazide
8-Hydroxylaminea, a'-Dipyridyl
Diethyldithiocarbamate
Cuprizone
Neocuproine
2,4-Dinitrophenol
EDTA
p-Chloromercuribenzoic acid
Monoiodoacetic acid

Inhibition (%)

54.1

10.9
74.7
45.8
< 0.1
58.4
95.5
95.6

94.4
57.4

29.1
52.9
88.4
15.8
7.8
< 0.1
9.5

7.9

* 9 mU of the monoamine oxidase were preincubated for 30 min at 4°C witha given inhibitor at 1 mM before the addition of tyramine under the standard
assay conditions by a radiometric procedure with [3Hl-Tyramine. The values
indicate the amount of the activity remaining after incUibation with eachcompound as a percentage of the activity in absence of the compound .

semicarbazide, which are known to be potent inhibitors of diamine oxidase(27). The moderate inhibition of the enzyme by 8-hydroxyquinoline and othermetal-chelating reagents indicates that the activity of the ~nzyme may be
metal-dependent.
Identification of the metal ion - Effects of metal ions at 1 m M on the
enzymatic were investigated (Table 4). The enzyme was activated by Cu 2 +ions and completely inhibited by Hg2+ ions. These results suggest that the
enzymatic activity is dependent on the Cu 2+ ions. Nara et a/. (3) demonstarted
that copper is a metallic constituent of animal monoamine oxidases. Yamada
et al. (36) also demonstrated that copper is a constituent of the amine oxidasefrom Aspergillus niger. Next, we tested the possibility of involvement of metal
ions in the oxidative processes, using a spectrophotometry revealed that
copper was present in the enzyme. The purified enzyme contained 10 g ofcopper per 79,000 g of protein . The same ammount of copper in the enzyme
was found even when the cells were incubated with 0.3 mM CUS04.
Furthermore the iron, manganase, cobalt, and zinc were also examined.
However, the only metal present in significant quantity was copper. Thus, wereexamined the absorption spectra of the purified enzyme with concentrated
sample. A shoulder at about 480 nm was noticed in the spectrum (Fig. 38).
This absorption band was partially bleached by the addition of tyram ine and ofexcess sodium dithionite under anaerobic conditions.
Search for homology - The amino acid sequence deduced from thenucleotide sequence of the maoA gene, which encodes the structural gene formonoamine oxidase from K. aerogenes W70, revealed that the mature protein
conteins 725 amino acids residues and has a molecular weight of 80,445(20). We searched for homologies between the amino acid sequence of
monoamine oxidase from K aerogenes and those of other amine oxidases.
We found that only peroxisomal amine oxidase from Hansenula polymorpha(37) showed some homology (Fig. 4) . However, the sequences of the amino

47

Table,4. Effects of metal ions on the monoamine oxidasefrom K. aerogenes W70 (pT058)*

Metal (1 mM)

Relative activity (%)

None
CuS04-5HzO
CuClz
CoClz-6HzO
ZnClz
CdS04
NiClz-6HzO
MnClz-4HzO
BaClz-2HzO
LiCI
MgClz-6HzO
CaClz
Pb(N03)Z
HgClz

"100
ll23
ll07
102
100
99
98
1101
97
98
95
99
87
0

CUS04- 5HzO (2mM)
(2mM)
CuClz

183
130

* Monoamine oxidase (12 mU) was preincubated for 30 min at 4°C with agiven metal salt at 1 mM before the addition of tyramine under the standard
assay conditions by a radiometric procedure with [3 H1]-Tyramine. Oxidationin the absence of metals is given as 100% ; other actiivities are given relative
to that value.

48

acids of the two enzymes exhibit 27.9% homology. In neither sequence we
could find the amino acid sequence Gly-X-Gly-X-X or Ser-Gly-Gly-Cys-Tyr,which are highly conserved in the flavin adenine dinucleptide-binding regionof flavoproteins of a number of oxidases and dehydrogenases (38, 39) .

49

MA 0

MA NGL1< FSPit 1< TAL ALAVAVVCAW QS PVFAII GSEA II MVr LDKTL0 EFGAD VOW DDYAQMFAMOMERLRQIASOATAASAAPAMAO

TL I KDGAY V1< V1< PG/\ 1< TAI VNG1< S LDLr Vr vvM1< EGK/\ WVSOT FIN DVF0 SGLDOT F0 VE

AM 0

RPAII P LDr LS TAElK /\ /\ TNT V1< S YFA- G1< 1< I S FNTVTLREP ARK AYlOW KEO GGP L- P P R· . . * * *. . *. * * *. *... * *.

. . . *. . . . *. * *.'. *

,;f

MAO

* *. *. . * *KI1PH P LNS LSAAE I SI<AV T I VKAAPE FOPNTI1FT E I S Lli EADI<AAVWAFALOGT P VOAP RAMO

LAYYVILEAGKPGVKEGLVDLASLSVIETRALETVOPILTVEDLCSTEEVIRNDPAVIEOMAO

TAD - VVMLDGKHVI - EAVVDLONKKILSWTPIKGAIIGMVLLDDFVSVONII - NTSSEFAE

AMO

CVLSGIPANEMHKVYCDPWTIG - YDERWG -- TGKRLOQALVYYRSDEDDSOYSIiPLD - FC

MAO

- VLKKIIGITDPGKVVTTPLTVGFFDGKDGLQQDARLLKVVSYLDTGDGNY - WAHPIENLVAMOPIVDTEEKKVIFIDIPNRRRKVSKHKHANFYPKHMIEKVGAMRPEAPPINVTOPEGVSFKMAOAVVDLEAKKIIKIE - EGPVIPVPMEPRP -- Y----- DGRDRNAPAVKPLEITEPEGI<NYTAMO

MTGNVMEWSNFKFHIGFNYREGIVLSDVSYNDHGNVRPIFHI1ISLSEMIVPYGSPEFPHO

MAO* *... ** .. *.. ***.. ....
**.

. ......... *. *.... * *..... .

** . * *. * . *.. *· . ** *. **. * *.

*. .. ..

. ** ... **.. ** . . .

*.. *... *. *** ...

· ** .. . . * *.. ** .. * * * . **. *. ***. *. *....

** .. ******. *.

ITGDTIHWONWDFHLRLNSRVGPILSTVTYNDNGTKRQVMYEGSLGGMIVPYGDPDVGWYAMO

RKHALDIGEYGAGYMTNPLSLGCDCKGVIHYLDAHFSDRAGDPITVKNAVCIHEEDDGLL

MAO

FKAYLDSGDYGMGTLTSPIVRGKOAPSNAVLLDETIADYTGKPTTIPGAVAIFERYAGPEAMOFKHSDFRDNFATSLVTRATKLVVSQIFTAANYEYCLYWVFMODGAIRLD- --- - - IRLTG

MAO

AMO
MAO

* ** *. ** * ,*. *.

· ** . ...

... *

* *

** ... * . *. *. *.. ** * * . *. ***. * *.. **. * . *** .. *. * . *.. *YKHLE-MGK -- PNVSTERRELVVRWISTVGNYDYIFDWVF~IDNGTIGIDAGATGIEAVKGILNTYILG-DDEEAGPWGTRVYPNVNAIINHOIILFSLRIDPI1IDGDGNSAAACDAKSSPYP.* . . . . . . . *.... ** .. *. . . *** ... . *.* .** .. *.. * *... *.

VLAKTMHDPSAKEDTRYGTLIDHNIVGTTHQHIYNFRLDLDVDGENNTLVAMDPEVKPNTAMOLGSPENMYGNAFYSEKTTFKTVKDSLTNYESATGRSWDIFNPNKVNPYSGKPPSYKLV--MAOAGGPRT - --STMQVNOYTIDSEOKAAOKFDPGTIR --- LLSNTSKENI1MGNPVSYOIIPYAMO--STOCPPLLAK--EGSLV/\KRAPWASIISVNVVPYKDNI1LYPSGDHVPQWSGDGVRGMRE

MAO

AGGTHPAATGAKFAPDEW I YIIR LSFMDKOLWVTRYHPTERYPEGKY -PNRSAHDT - GLGOAMO

WI GDGSEN I DNTD I LFFlfTFG I THFPAPEDFPLMPAEP I TLMLI1PRHFFTENPGLD lOPS

MAO

YAKDDESLTNHDDVVWITT - GTTHVARAEEWPIMPTEWALALLKPWNFFDETPTLGEKKKAMOYAMTTSEAKRAVIIK ETKDK TSHLAF EGSCCGK*. * .

. *. ..

.. *... ...

**. .... * *.... * . .. .. *.. *..

*....... *.... * *. ** .. *.. *. **. *..... ,*. * ** ...

**. *.. *.. *.... *..
. *. *.. **. *. *. *.

Fig. 4. Comparison of amino acid sequences of monoamine oxidasefrom K. aerogenrs (MAO) and peroxisomal amino oxidase fromHansenula po/ymorpha (AMO). Identical amino acid (*) and ,chemically similar ami'no acids (.) are indicated.

5(lDiscussion
The cloning, expression, and purification system described here has madepossible the preparation of substantial quantities of highly purified
monoamine oxidase in a soluble form. Previously, we determined thecomplete nucleotide sequence of the gene for monoamine oxidase (maoA)from K aerogenes W70 (26). In this sequence, two open reading frames(ORFs), capable of coding for 36-KOa (ORF1) and 84-KDa (ORF2) proteins
were found (26). To identify the product of the maoA gene, we determined the
N-terminal sequence and amino acid composition of the purified monoamineoxidase. The first 11 amino acids from the N-terminus and the amino acidcomposition of the enzyme were identical to those of the predicted mature
protein encoded by ORF2, as deduced from the nucleotide sequence. The
leader peptide extends over 30 amino acids in the precursor, and thetranslated polypeptide is processed to generate the mature enzyme. Thus, thepurified enzyme can be considered homogeneous.
Unlike other monoamine oxidases (1, 3, 27), monoamine oxidase from K.

aerogenes was not inhibited by SH-reagents, such as p-chloromercuribenzoate, which suggests the absence of any role for an essential cystein
residue in the structure and activity of the enzyme. In fact, We found no cystein
residures in our amino acid analysis or in the peptide sequence deduced fromthe nucleotide sequence of maoA gene (26). Patterns of inhibition of the
enzyme by SH-reagents and carbonyl reagents suggest that the mechanismof the reaction catalyzed by the monoamine oxidase of K. aerogenes is likethat of diamine oxidase (7, 27). We showed that the monoamine oxidase from

K aerogenes contains a metal, copper, as an essential cofactor, as do someother amine oxidases (3,36). The activity of the purified enzyme wasstimulated about 20% by Cu 2+ ions but not to any significant extent by othermetals tested. The copper was present in the cupric state. This conclusion isbased on the fact that direct determination of copper in the enzyme accounted

51

for all the copper, and on the fact that the enzyme was inhibited by cuprizone
and diethyldithiocarbamate, which are specific for cupric copper, and not by
neocuproine, which is specific for cuprous copper. Finally" the native enzyme
was shown to contain 10 g of copper per 79,000 g of protein. Since one moleof monoamine oxidase has only 0.16 mole of copper, the enzyme seems to be
not fully saturated with the metal when released from the cell. Metal substitution may occur in this enzyme, but other metal ions did not detect in thisenzyme.
The enzyme from K aerogenes did not give a shoulder of absorbancecentered around 460 nm, even though a FAD-dependent peak of absorbanceat 460 nm was found in the case of the putrescine oxidase from Micrococcus
rubens (33) or hepatic monoamine oxidase (38) . A copper-dependentshoulder of absorbance at 480 nm (43) as seen in copper-proteins was barely
observed when the concentrated samples (259 mg/ml) was used . The faintreddish color of the enzyme could be related to the shoulder around 480 nm .
This absorption band was partially reduced by substrate as well as sodium
dithionite (Fig . 38) . Amine oxidases can be divided into two groups, those with
FAD and those with copper as the prosthetic group. Almost all diamineoxidases, such as the animal and plant diamine oxidases (40, 41) , plasmaamine oxidase (42) , and the amine oxidase from A. niger (36) , and few of
monoamine oxidase from such as that from K. aerogenes, belong to the lattergroup. Amine oxidases of the latter group are inhibited by carbonyl reagents.
The sensitivity to carbonyl reagents of these amine oxidases suggests that a
carbonyl group may be located at the active site of the enzyme. Yamada et al.
(43) reported that the prosthetic group of amine oxidase from bovine plasma
was a copper complex with pyridoxal phosphate. Mondovi et al. (44) foundthat the activity of the highly purified animal diamine oxidase was increasedby the addition of pyridoxal phosphate. Although this activation may beunspecific, it suggests that pyridoxal phosphate ia a coenzyme for the animal

52

diamine oxidase. The monoamine oxidase of K. aerogenes seems to bedifferent from these pyridoxal phosphate-containing enzymes, since noI'phosphate-dependent peak of absorbance was observed, Recently, theamine oxidase from Hansenula polymorpha was isolated and the tyrosine
codon of this enzyme corresponded to 6-hydroxydopa quinone (also known
as topa quinone) at the active site (45) . More recently, the amino oxidase from
Escherichia coli K- 12 grown on 2-phenylethylamine has been shown to havecopper and topa quinone as cofactors (46). The translated nucleotide
sequence of E. coli amine oxidase gene included the consensus sequence of
topa quinone, Asn-Tyr-Asp-Tyr. A comparison of the amino acid sequence ofthe yeast amine oxidase and E. coli amine oxidase with the deduced amineoxidase sequences of the monoamine oxidase from K aerogenes W70 (26)suggests that the enzyme reported here could be a copper- and topa quinonecontaining monoamine oxidase (EC 1.4.3.6) .

Summary
The gene for monoamine oxidase (maoA) from Klebsiella aerogenes W70~.

has been cloned and the enzyme was overproduced in a 'SOluble form . Tne
enzyme was purified approximately 10-fold to homogeneity. The enzyme has
a molecular weight of about 79,000, which is identical to the molecular weight
deduced from the necleotide sequence of the gene for monoamine oxidase,and it consists of individual monomers. The enzyme had maximum activity at
pH 6.0 and 50°C when catalyzing the oxidative deamination of tyramine. The
enzyme catalyzed the deamination of ~- phenylethylamine , dopamine ,tryptamine, and octopamine, but not of diamines, polyamines, or amino acids.
The enzyme was inhibited by clorgyline, isoniazid, and carbonyl reagents, but
not by 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide. The enzyme did notexhibit a typical flavoprotein spectrum , but the enzymatic activity increased
linearly with increasing amounts of added copper. The purified enzyme wasfound to contain 10 g of copper per 79,000 g. The enzymological properties
and the amino acid sequence of the enzyme deduced from the nucleotide
sequence of the maoA gene are different from those of known tyramine ormonoamine oxidase. Thus, monoamine oxidase from K. aerogenes seems to
be a new type of monoamine oxidase.

References1.

Erwin, V. G. and Hellerman, L. : Mitochondrial monoamine oxidase :j'purification and characterization of the bovine kidney enzyme. J. BioI.
Chern ., 242,4230-4238 (1967).
2.

Igaue, I. , Gomes, B., and Yasunobu , K. T .: Beef mitochondrialmonoamine oxidase, a flavin dinucleotide enzyme. Biochem . Biophy.
Res. Comm ., 29, 562-570 (1967).

3.

Nara, S., Gomes, B., and Yasunobu , K. T .: Amine oxidase : beef liver
mitochondrial monoamine oxidase, a copper containing protein. J . BioI.
Chern ., 241 , 2774-2780 (1966) .

4.

Salach , J . I. : Monoamine oxidase from beef liver mitochondrial :simplified isolation procedure, properties, and determination of its
cysteinyl flavin content. Arch . Bioch. Biophy., 192, 128-137 (1979).

5.

Youdium , M. B. H. and Sourkes, T. L.: Properties of purified , solublemonoamine oxidase. Canadian J . Biochem., 44, 1397-1400 (1959) .

6.

Blaschko, H., Friedman , P. J., Hawes, R., and Nilsson, K.: The amineoxidase of mammalian plasma. J. Physio!. , 145, 384-404 (1959) .

7.

McEwen, Jr. C. M.: Human plasma monoamine oxidase. I. Purificationand identification . J . BioI. Chern ., 240, 2003-2010 (1965) .

8.

Tabor, C. W ., Tabor, H., and Rosenthal , S. M.: Purification of amineoxidase from beef plasma. J. BioI. Chern ., 208, 645-661 (1954).

9.

Yamada, H. and Yasunobu , K. T .: Monoamine oxidase. I. Purification ,crystallization, and properties of plasma monoamine oxidase. J . BioI.
Chern ., 237, 1515-1516 (1962) .

10.

McGowan, R. E. and Muir, R. M.: Purification and properties of amineoxidase from epicotyls of Pisum sativum. Plant Physiol ., 47, 644-648(1971 ).

11 .

Yamada, H., Adachi, 0 ., and Ogata, K.: Amine oxidases of
microorganisms. Part I. Formation of amine oxidase by Fungi. Agric. BioI.

Chem., 29,117-123 (1965).
12.

Yamada, H., Uwajima, T., Kumagai, H., Watanaba, M" and Ogata, K.:~.

Bacterial monoamine oxidases. Part I. Purification and crystallization oftyramine oxidase of Sarcina lutea. Agric. BioI. Chem ., 31, 890-896(1967).
13.

Murooka, Y., Doi, N., and Harada, T.: Distribution of membrane-boundmonoamine oxidase in bacteria. Appl. Enviro. Micro., 38, 565-569(1979).

14.

Okamura, H., Murooka, Y., and Harada, T.: Regulation of tyramineoxidase synthesis in Klebsiella aerogenes. J. BacterioL, 127, 24-31(1976).

15.

Yamada, H., Kumagai, H., and Uwajima, T. : Tyramine oxidase (Sarcina

lutea). Method. EnzymoL, 17B, 722-726 (1971).
16.

Oka, M., Murooka, Y., and Harada, T.: Genetic control of tyramineoxidase, which is involved in derepressed synthesis of arylsulfatase inKlebsiella a erogenes. J. BacterioL, 143,321-327 (1980).
17.

Murooka, Y., Adachi, T ., Okamura, H., and Harada, T.: Genetic control ofarylsu Ifatase in Klebsiella aerogenes. J. BacterioL, 130, 74-81 (1977).

18.

Murooka, Y., Higashiura, T., and Harada, T.: Genetic mapping oftyramine oxidase and arylsulfatase genes and their regulation inintergenic hybrids of enteric bacteria. J. BacterioL, 136,714-722 (1978).

19.

Murooka, Y., Ishibashi, K., Yasumoto, M., Sasaki, M., Sugino, H.,
Azakami, H., and Yamashita, M.: A sulfur and tyramine-regulated

Klebsiella aerogenes operon containing the arylsulfatase (atsA) gene
and atsBgene. J. BacterioL, 172,2131-2140 (1990).
20.

Sugino, H., Ishibashi, K., Sakaue, M., Yamashita, M., and Murooka, Y.:
Gene cloning of the maoA gene and overproduction of a solublemonoamine oxidase from Klebsiella aerogenes . Appl. Microbiol.
Biotechnol., 35, 606-610 (1991).
,~

DY'. ,

21 .

MacPhee, O. G., Sutherland, I. N., and Wilkinson, J. F.: Transduction in.

Klebsiella. Nature, 221,475-476 (1969) .
'22.

Adachi, T ., Murooka, Y., and Harada, T. : Catabolite repression andderepression of aryl sulfatase synthesis in Klebsiella aerogenes. J .
Bacteriol., 120, 880-885 (1974) .

23.

Lowry, O. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J.: Protein
measurement with the folin phenol reagent. J. BioI. Chem., 193,
265-275 (1951).

24.

Laemmli, U. K.: Cleavage of structural proteins during the assembly ofthe head of bacteriophage T 4. Nature, 227, 680-685 (1970) .

25.

Andrew, P.: The gel-filtration behaviour of proteins related to theirmolecular weights. Biochem . J ., 96, 595-606 (1965).

26.

Sugino, H., Sasaki, M., Azakami , H" Yamashita, M., and Murooka, Y.: A
monoamine-regulated Klebsiella aerogenes operon containing themonoamine oxidase the monoamine oxidase (maoA) and maoCgenes.
J. BacteriaL, 174,2485-2492 (1991).

27.

Yamada, H., Kumagai, H., Uwajima, T., and Ogata, K.: Bacterialmonoamine oxidase. Part II. Substrate and inhibitor specificities oftyramine oxidase of Sarcina lutea. Agric. BioI. Chem ., 31 , 897-901(1967).

28.

Nagy, J, and Salach, J. I.: Identity of active site flavin-peptide fragmentfrom the human "A"-form and the bovine "B"-form of monoamineoxidase. Arch. Biochem. Biophys., 208,388-394 (1981).

29.

McEwen, Jr. C. M., Sasaki, G., and Jones, D. C.: Human liver
mitocondrial monoamine oxidase. III. Kinetic studies concerning timedependent inhibitions. Biochem ., 8, 3963-3972 (1969).

30.

Singer, T. P.: Monoamine oxidase: structure, functions and alteredfunctions. Academic Press, New York (1979).

31 .

Adachi, 0 ., Yamada, H., and Ogata, K.: Purification and properties of

5'7

putrescine oxidase of Micrococcus rubens. Agric. Bioi . Chem ., 31 , 1202-

1210 (1966).
32.

Desa, R. J.: Putrescine oxidase of Micrococcus rubens. Purification andproperties of the enzyme. J . BioI. Chem ., 247, 5527-5534 (1972) .

33.

Yamada, H., Adachi , 0 ., and Ogata, K. : Putrescine oxidase, a diamineoxidase requiring flavin adenine dinucleotide. Agric . BioI. Chem ., 29,1148-1149 (1965).
34.

Ameyama, M., Hayashi, M., Matsushita, K., Shinagawa, E., and Adach i,
0 .: Microbiol production of pyrroloquinoline quinone. Agric . BioI. Chem .,48, 561-565 (1984).
35.

Fukuyama, M. and Miyaka, Y.: Purification and some properties ofcholesterol oxidase from Schizophyl/um commune with covalently
bound flavin . J . Biochem ., 85, 1183-1193 (1979).

36.

Yamada, H., Adachi , 0 ., and Ogata, K.: Amine oxidase of microorganisms. Part IV. Further properties of amine oxidase of Aspergillus

niger. Agric. BioI. Chem ., 29, 912-917 (1965).

37.

Sruinenberg , P. G., Ever, M. , Waterham , H. A., and Kuipers. J.: Cloning
and sequencing of the peroxisomal amine oxidase gene fromHansenula polymorpha. Biochim . Biophys. Acta., 1008, 157-167 (1989) .

38.

Kearney, E. S., Salach, J. I. , Walker, W. H., Seng, R. I. , Kenny, W.,
Zeszotek, E. , and Singer, T . P.: The convalently bound flavin of hepaticmonoamine oxidase. Eur. J. Biochem ., 24, 321 -327 (1971) .

39.

Untucht-Grau, A., Blatterspiel , A., Krauth -Siegel , A. L. , Salach , M.,
Schiltz, E., Schirmer, R. H., and Liebold , B. W.: The primary structure ofglutathione reductase from human erythrocytes, pp. 38-43. In V. Massey
and C. H. Williams, Jr. (ed .), Flavin and Flavoproteins. Elsevier/NorthHolland Publishing Co., New York (1982) .

40.

Hill, J. M. and Mann, P. J. G.: The inhibition of pea-seeding diamineoxidase by chelating agents. Biochem . J., 85, 198-207 (1962) .

58

41.

Hill, J. M. and Mann, P. J. G. : Further properties of the diamine oxidaseof pea-seedings. Biochem. J., 85, 198-207 (1962).
I

42.

'

Yamada, H. and Yasunobu, K. T.: Monoamine oxidase. II . Copper, oneof the prosthetic groups of plasma monoamine oxidase. J. BioI. Chem.,
237,3077-3082 (1962).

43.

Yamada, H. and Yasunobu, K. T.: Monoamine oxidase. IV. Nature of thesecond prosthetic groups of plasma oxidase. J. BioI. Chem., 238, 26692675 (1962).

44.

Mondovi, B., Rotilio, G., Finazzi, A., and Scioscia-Santoro, A. :Purification of pig-kidney diamine oxidase and its identity withhistaminase. Biochem. J., 91,408-415 (1964).

45.

Mu, D., Janes, S. M., Smith, A. J., Brown, D. E., Dooley, D. M., andKlinman , J. P.: Tyrosine codon corresponds to topa quinone at theactive site of copper amine oxidase. J. BioI. Chem ., 7979-7982 (1992) .

46.

Cooper, R. A., Knowles, P. F., Brown, D. E., McGuirl, M. A., and Dooley,
D.M. : Evidence for copper and 3,4,6-trihydroxyphenylalanine quinone
cafactors in an amine oxidase from the Gram-negative bacterium

Escherichia coli K-12. Biochem. J., 288, 337-340 (1992).

59

~

.

Chapter III

Structure and Function of Pullulanase:

Role of Lipid Modification on a Starch-debranching Enzyme, Klebsiella
Pullulanase60

Introduction
Pullulanase (EC 3.2.1.41) is a maltose-inducible lipid-modified enzyme..

produced by bacteria belonging to the genus Klebsiella (Bender andWallenfels, 1961 ; Konishi et aI., 1979). The enzyme is specifically released
into the medium by Gram-negative Klebsiella aerogenes (Takizawa and
Murooka,1984; 1985) and Klebsiella pneumoniae (d'Enfert et al., 1987).
previously, we cloned the pullulanase gene (pulA) from Klebsiella aerogenes
W70 into vector of pK1212, which is stable maintained in Klebsiella species(Takizawa and Murooka, 1985). Escherichia coli cells carrying therecombinant plasmids producted three to seven-fold more pullulanase than
did the wild-type strain of K aerogenes W70, but most of the pullulanaseremained intracellular, whereas K aerogenes W70 released pullulanase
entracellularly. Transfer to the plasmid into the original producing strain (W70)
resulted in an approximately 40-fold increase in pullulanase production , with
the intracellular enzyme level being about 100-fold higher than that of strain
W70. This accumulation of the enzyme may be caused by titration of 'secretion
factors' responsible for pullulanase release. These factors are encoded by 14
genes, which are linked to the pulA gene (Pugsley and Reyss, 1990; Reyss
and Pugsley, 1990).
Pullulanase is produced as a precusor with a hydrophobic N-terminal signal
peptide. The sequence of K. pneumoniae pullulanase is highly homologous tothat of pullulanase of K. aerogenes. However, K. pneumoniae UNK5023
pullulanase (Kornacker and Pugsley, 1989) lacks a collagen-like sequence
present at N-terminus of the mature K aerogenes W70 enzyme. Pugsley et aI. ,(1986) and Katsuragi et al., (1987) found independently that pullulanase of K.
pneumoniae and K aerogenes are modified by glycerly lipids. The lipids arecovalently attached to the Cys-20 residue in the pentapeptide sequence, LeuLeu-Ser-Gly-Cys, which is similar to the pentapeptides that are adjacent to thesites of post-translational modification and processing of other prolipoproteins61

in E. coli (Inouye et a/., 1977). To investigate this modification, Cys-20 was
replaced by Ser-20. This prevented fatty acylation and caused slow cleavagej'of the signal peptide to be processed at a site within the hydrophobic part ofthe signal peptide. A large amount of the unmodified mutant pullulanase
accumulated in the periplasmim of K aerogenes W70 during the logarithmic
phase of growth, but some was secreted into the medium (Murooka and Ikeda,
1989). Thus, efficient secretion and correct processing of Klebsiella pu"ulanse
require the modification of lipid. In this report, we compare other biochemical
properties of the two forms of the enzyme.

62Materials and methods
Materials, bacterial strains and plasm ids - Pullulan (Mr, 5 x 104 ), G 1- to G5I 'a-cyclodextrins and crystalline pullulanase from K. pneuf170niae IF03321
were provided by Hayashibara Co. Ltd " (Okayaka, Japan) . Pure collagenasefrom Achromobacter iophagus was purchased from Boehringer-Yamamouchi
Co. Ltd., (Tokyo, Japan).

A" other reagents were standard commercial

preparations. The bacterial strains used in this study were K aerogenes W70
and E. coli C600 (leu, thr, thl). pMPK1, a plasmid contains the pulAgene, andplasmid pMPK151, in which the pulA gene was mutagenized (Cys_20 to Ser20), were constructed by Murooka and Ikeda (1989).
Production and assay of pullulanase - K. aerogenes W70 carrying pMPK1
or E. coli C600 carrying pMPK151 were cultured for 3day in minimal medium(Takizawa and Murooka, 1985). E. coli C600 cells without the plasmid did not
produce any pullulanase (Takizawa and Murooka, 1985). The medium used
for production of pullulanase contained 0,5 % soluble starch for K aerogenes
W70 carrying pMPK1 and 0.5% pullulan for E. coli C600 carrying pMPK151 asa sole source of carbon. Pullulanase activity was measured as descreived by
Takizawa and Murooka (1984). The release of aldehyde groups from pululan
was measured by the Somogyi method as modified by Nelson (1944) . One
unit of activity was defined as the amount of enzyme causing liberation of
11-lmole of aldehyde per min. Proteins were measured by the method of Lowry
et al. (1951) with crystalline bovine albumin as a standard. The absorbance at

280 nm was used to monitor protein in the column effluent.
Purification of pullulanase - All the steps were carried out at 4°C.
Step 1. Preparation of subcellular fractions. Three days culture broth (about 6I) was used . Pullulanases from the extracellular, periplasmic, and cytoplasmic
fractions were perpared as described previously (Murooka and Ikeda, 1989).
Step 2. Fractination with ammonium sulfate. The crude enzyme wasfractionated with 20 to 800/osaturation of ammonium sulfate. The precipitate63was dissolved in a small volume of 0.02 M acetate buffer (pH 6.0) anddialyzed against the same buffer.
~.

Step 3-1. DEAE-Toyopearl 650S column chromatography. The dialyzate wasapplied on DEAE-Toyopearl 650S column equilibrated wih 0.02 M acetate
buffer (pH 6.0) containing 0.1 M NaC/. In the extracellular wild-type enzyme,
two peaks of pullulanase activity were appeared (Fig. 1A) . The two peaks (Iand II) were concentrated and purified individually.
Step 3-2. Butyl-Toyopearl 650M column chromatography. For the purificationof periplasmic enzyme, this step was added. The dialyzate was brought to 1.5
M saturation by the addition of ammonium sulfate and the mixture was applied
onto a Butyl-Toyopearl 650M column equilibrated with 0.02 M acatate buffer(pH 6.0) containing 1.5 M saturation of ammonium sulfate. The proteinsabsorbed were eluted with a decreasing gradient of ammonium sulfate (1.5 too M) in the same buffer.
Step 4. Gel filtration with Toyopearl HW55S. The concentrated solution wasapplied onto a Toyopearl HW55S column equilibrated with 0.02 M acetate
buffer (pH 6.0). Elution was performed with the same buffer. The fractions withenzyme activity were collected and the solution was concentrated with amembrane filter.
Step 5. High-performance liquid chromatography. Final purification of
enzyme was carried out by a high-performance liquid chromatography on a
DEAE-NPR column (Toso Co. Ltd., Tokyo,Japan).

64

Results
Purification of the pullulanases - The wild-type and mutant pullulanases..

were purified about 150-fold with specific activity of 132.1,Umg-1 protein andabout 110-fold with specific activity of 139.9 Umg-1 protein, respectively (Table1). The wild-type enzyme chromatographed as two active peaks, eluting at
0.15 M (I) and 0.3 M (II) NaGI, on OEAE-Toyopearl (Fig. 1A), whereas themutant enzyme chromatographed a single active peak eluting at 0.3 M NaGI(Fig . 18). The two active fractions of the wild-type pullulanase wereindistiguishable on sodium dodecyl sulphate polyacrylamide gelelectrophoresis (SOS-PAGE) and in enzymeatic properties (data not shown)(Table 1 shows data only for peak I).
Both purified enzymes migrated a single protein band (on SOS-PAGE) (Fig.
2). More than 90% of the extracellular wild-type pullulanase was modified withglyceryl lipids, but none of the mutant pullulanase from any cell fractions
contained lipid (Murooka and Ikeda, 1989). The molecular masses of the wildtype and mutant enzymes were both determined to be 120 kOa by SOSPAGE. However, the wild-type enzyme usually appeared as two active peaks
on gel filtration (appromimately 350 kOa and 110 kOa). Since the protein withthe higher molecular mass behaved as a single protein of 120 kOa on SOSPAGE, we concluded that it probably coresponds to a trimer reported for the K.
neumoniae FG9 pullulanase (Gharalambous et al., 1988). The other peak ispresumably monomer, for which a mass of 117 kDa is predicted from DNA
sequence data (Katsuragi et aI., 1987). In contrast to the wild -type
pullulanase, the mutant pullulanase produced a single active peak on gel
filtration. These results suggest that lipid palys a key role in facilitating
trimerization of pullulanase.
Properties of the unmodified enzyme - The effects of pH and tamperetureon active and stability of the mutant pullulanases were compared with those of
the wild-type and the K. pneumoniae (Aerobacter aerogenes) IF03321

65

Table 1. Purification of the extracellular wild-type and the periplasmic
mutant pullulanases

Fraction

Total
activity(U)

Total
protein(mg)

Specific
activity(U/mg)

Yield(%)

Crude extract

1345

1520

0.9

100

(1971)(1247)

(1.3)

(100)

1088

87.4

12.4

89

(2063)(148)(13.9)(123)(1 597)(31 .7)(50.4)(96)

Ammoniumsulfatefraction(20-80%)Butyl-T oyopearl
650M
DEAE-T oyopearl
650S

455

13.0

35.1

34

(1236)(9.6)(129.0)(74)

Toyopearl HW55S

261

3.2

81.3

19

DEAE-NPR

230
(854)

1.7
(6.0)

132.1
(139.9)17(51)

The extracellular wild-type and the peri plasmic mutant pullulanases wereprepared from K. aerogenes W70 (pMPK1) and E. coli C80a (pMPK1),
respectively. Values in the parenthese represent the peri plasmic enzyme,
Butyl-Toyopearl 850M colume was used, whereas Toyopearl HW55S wasused for the extracellular enzyme. In the extracellular wild-type enzyme,
purification steps from DEAE-Toyopearl 8508 show only peak (I) (Fig. 1A) .

66

2 0 ------------------------------------~

A

II

o

,,-....,

E

I

c

o

U

ro

00
N

Z

...........
Q)

0.5

u

c:

ro

.0
L-

o

(/)

.0

«

0.1
o~~~--~--~~~~~~~~--~L-~

o

10

20

40

30

Fraction number
5

B
4
0

I 3

,,-....,~...........

,,-....,

u

c

O.S

00
N
...........

roZ,,-....,

E

0..........

:::>...........

E

Q)

u

c

2

ro

~
+J

.0
L0
(/)
.0

:~
+J
u

«

«

0.1
o+----+~~r=~~~~----~~~--~~~

o

10

20

30

40

Fraction number
Fig . 1. Elution pattern of the extracellular wild-type pullulanase (A) and mutant-type
pullulanase (8) on DEAE-ToyopearI650S. Elution was done with a linear gradientof 0.1 to 0.5 M NaCI. The two active fractions, I and II were purified individually.

67

..
B

A

M 1 2 3170kd
116kd iM 1 2 3

170kd

4 5

6

7

8

9

10

. -- .. ... . . .
"

116kd

76kd
76kd
53kd
53kd

-- - -- -- - ----- - -Fig . 2. SOS-PAGE of purified pullulanase from the culture medium of the wild-type
W70 cells carrying pMPK1 (A) and the peri plasmic fraction of E. coli cells carrying
pMPK151 (B) . Polyacrylamide gel electrophoresis with and without 0.1% SOS was
carried out by the method of Laemmli (1910) . The gel were stained fro proteins with
Coomassie Brilliant Blue. A. Lane 1, ammonium sulfate fraction ; lane 2, fraction of
OEAE-Toyopearl 650S ; lane 3, fraction of Toyopearl HW55S. B. Lanes 1, 4, 7,
ammonium sulfate fraction; lanes 2,5,8, fraction of Butyl-ToyopearI650M ; lanes 3,6, 9, fraction of OEAE-Toyopearl 650S ; lane 10, fraction of OEAE-NPR . Lanes 1, 2,
3, extracellular fraction ; lanes 4, 5,6, periplasmic fraction ; lanes 7, 8, 9, 10, cytoplasmic fraction .

68

pullulanse, which is though to be identical to that of the W70 enzyme (Ohbaand Ueda, 1973). The enzyme activities of these three pullulanase were..

maximal at pH 6.0. They were stable at lower pH but comJ>letely inactivated at
pH 12.5, although the IF03321 pululnase showed lower activity at pH 7.0 or
more alkaline pH (Fig. 3). Activities of the wild-type and mutant pullulanases
were maximal at 60°C (when stabilized by the addition of the substrate
pullulan), whereas the optimal temperature for the IF03321 enzyme was 50°C
(Fig. 4A). Nearly half the activities of both the wild-type and the mutant
pullulanases were lost at 50°C when incubated for 30min in the absence ofpullulan (Fig. 48).
The substrate specificities of three pullulanase were also examined (Table
2). All enzyme showed high activities towards pullulan , moderate activity
towards amylopectine and soluble starch but little or no activity towards oyster
glycogen, a- or ,:l-cyclodextrins, or glycosyl-(a- 1-6)-cyclodextrin . Maltosyl-,
maltotoriosyl-, maltotetrasyl-, and maltopentasyl-(a -1-6)-cyclodextrins werehydrolyzed to similar degrees by the three enzymes. The Km and Vmax valuesof both the Wild-type and mutant pullulanases for pullulan were 1.7 x 10-3
gml- 1 and 556 ~moles min- 1 mg- 1 of protein, respectively.
The effects of various compounds on the activities of the wild-type, mutant
and IF03321 enzymes were examined. Ca2+, 8a2+, Fe 2+, C02+, Zn 2+,
Mn 2+, U 2+, and Mg2+ had no effect, but 0.1 mM HgCI2 inhibited the enzyme
activity completely and 0.1 mM CUCI2 caused about 60% inhibition. One mM
SOS and 1 m M ethylendiaminetetraacetic acid (EDT A) were not inhiibitory.
Since 2-mercaptoethanol, dithiothreitol, and p-chloromercuribenzoate were
not inhibitory, an accessible sulphydryl residue seems nonessential for theactivity of pullulanase. Similar values were obtained for the three enzymes.
Physical difference between lipid-modified and unmodified pullulanases It is known that a lipoprotein of the outer membrane of E. coli changes its
apparent molecular weight upon heating in solutions containing SOS

69

120

A
100
;--..

~

9~1

...........
~

~

'5

I '

.~

rn

>

"

""
",'. ,
, ,

.,.
,

~

60

----.',

I,

\1

U

Q)

~-- -- Q

~P

80

.~

.

..,..
'

•

rn
Q)

0:::

40

..,

20

.'.b,

..
h

0
4

2

6

pH

10

8

12

120

B
100
;--..

~
...........
~

~

'5
u

80

.~

rn
Q)

>

60

.~

rn
Q)

0:::

40

20

O+-~--~--~~--~~--~--~~--~~~

2

4

6

8

10

12

14

pH
Fig. 3. Effect of pH on enzyme activity (A) and stability (8) of pullulanase.
The enzyme activity was assayed with various buffer. ( - -), acetate buffer;( - ) , phosphate buffer; (•••••• ), glycine-NaOH buffer. (0 ), wild-type
pullulanase; (.), mutant pullulanase; ( 6), crystaline IF03321 pullulanase.
For the pH stability, the enzyme solution was incubated at 4°C for 30 min.

70

120

A
100
.-

':!!.

0
.........

~

80

'5

·u«1
<1>

>

60

.~

<1>

a::

40

20

0
10

20

30

40

50

70

60

Temperature (OC)
120

8
100
.-

':!!.

0
.........

~

80

'5

·u«1
<1>

>

60

.~

Q5

a::

40

20

O+-~--~~--~~--~~--~~~~~--~

10

20

30

40

50

60

70

Temperature (OC)Fig. 4. Effect of temperature on enzyme activity (A) and stability of pullulanase (8).
(O),Wild-type pullulanase; (e), mutant pullulanase; (~), crystalline IF03321
pullulanase. For the thermstability, the enzyme solution in 0.1M acetate buffer,
pH 6.0, was kept at various temperature for 30 min.

71

,

"

Table 2. Comparison of substrate specificity for pullulanase of the wild-type
W70, mutant and IF03321.
Relative activity (%)
Substrate
Wild-type

MutantIF03321

Pullulan
Amylopectin
Soluble starch
Oyster glycogen

100
30
20
2

100
28
18
2

100
34
26
5

a-Cyclodextrin

2

0

0

~-Cyclodextrin

2
1
13

0
2
14

0

25

32
32
27

Gl-a-Cyclodextrin
G2 -a-Cyclodextrin
G3-a-Cyclodextrin
G4-a-Cyclodextrin
GS-a-Cyclodextrin

27
31

0
0
31
30
28

Enzyme activities were calculated from the amount of reduing sugars to thatof total sugars of each molecules. The values indicated the relative percentage of the activity of each substrate to that of pullulan.

72

(Schanitman, 1973). Thus, we tested the migrations of the lipid-modified andunmodified pullulanases were heatd in SOS-PAGE. When these pullulanases~.

were heated in SOS solution, they showed the same mobility (Mrapproximately 120,000) on SOS-PAGE (Fig. 5). However,when thesepullulanases were not heated, they migrated more faster and have theapparent Mr of about 60,000. The mutant pullulanase migrated more slowly
than the the band of the wild-type pullulanase or the crystalline IF03321
enzyme, which showed rather broad band.
To test whether the difference of migration rates between the wild-type andmutant pullulanases was caused by binding of lipids to the protein, bothpullulanases were digested with collagenase. The pullulanase from W70 has
a collagenase recognition sequence six repeated of the tripeptide (Gly-X-Pro)
between Gly_11 and Pro- 28 from the N-terminus of mature enzyme and isdigested by bacterial collagenase in this sequence, as reported byCharalambous et a/. (1988) for the FG pullulanase. Collagenase digestion
caused unheated wild-type enzyme to migrate with the unmodified (mutant)
enzyme (Fig. 5) These results suggest that the difference in migration rates
between the wild-type and mutant pullulanases is caused by binding of lipid
and/or the sequence of the extreme N-terminal region of the mature enzymes.

73

I '

::-•

II _
crm.
+--+-- + - - •--+ --+ - - +
n·
,.

-

N

-170"

.. -116 ..

....

...

- 16kd\ : - 53kd

1 2 J • I • J • I 10

Fig. 5. Effects of heating and collagenase digestion of pullulanase on the migrations
on SOS-PAGE. Lanes 1-3, wild-type pullulanase; lanes 4-6, mutant pullulanase,lanes 7-9, IF03321 pullulanase. Lanes 1,4,7, heating for 5 min in the Laemmli buffer(Laemmli, 1970); lanes 2,5,8, without heating; lanes 3,6,9, collagenase digestion(0.01 U mr 1 of 0.02 M phosphate buffer, pH 7.2) without heating; lane 10, molecularmarker.

·1

Discussion
We have examined the effects of lipid modification of the secreted enzymeI'pullulanase on enzymatic properties and physical stabilitY'. No distinct
differences in various properties (optimal pH and temperature, substratespecificity and affinities, and pH and tempareture stabilities) were observed
between the lipid-modified and the un-modified pullulanases. However, wefound that pullulanase from K pneumoniae IF03321 differs from W70 enzyme
in optimal temperature and thermal stability, implying that these enzymes are
not identical.
Extracellular and intracellular wild-type pullulanases were usually separable
by DEAE-Toyopearl column chromatography, into two active fraction (Fig. 1A) .
Since the mutant pullulanase chromatographed as a single peak (Fig. 1B),
this phenomenon may be due to the lipid molecule in the wild-type
pullulanase. Pugsley et al. (1991) also observed aberrant migration of wildtype pullulanase in gel when the crude samples were not heated to 100°C.
We now report that the lipid-modified and unmodified pullulanases migratedifferently in SOS-PAGE when the enzymes are not heated prior to loading on
the gel. The presence or absence of reducing agents such as fl-m ercaptoethanol did not affect electrophoretic mobility. It is possible that heating
causes unfolding of pullulanase with consequent increased binding of SOS .
This alteration of migration in SOS-PAGE probably accounts for the variationin estimates of Mr of pullulanase (143,000 according to Eisel et al., (1972) and
70,000 to 80,000 according to Ohba and Ueda (1973)). Furthermore, we also
confirmed the finding by Charalambous et al. (1988) that pullulanase exists as
non-covalently associated homotrimers. Three monomers of pullulanase may
associate to form a collagen triple-helical structure that serves to stabilize thetrimer in a manner analogous to the structure of a number of mammalian'collagen-like' proteins (Charalambous et al., 1988) However, since themature enzyme, which retains the collagen-like sequences but lacks the fatty

75acids, migrates as a monomer upon gel filtration without collagenase
traetment, both the collagen-like sequence of the pullulanase polypeptide andj'

the N-terminal fatty acids must be required to stabilized the trimeric form .

76

Summary

Klebsiella pullulanase is a lipoprotein synthesized as a precursor with aI 'signal peptide, which is processed by lipoprotein signal peptidase. To clarify
the role of lipid modification of pulullanase, we purified lipid-modified wildtype and the unmodified (mutant) pulullanases and compared thier properties.
The Km and Vmax values of both pulullanases for pullulan were the same. The
optimal pH and temparature, the stabilities over pH and temparature ranges,
the specificity of substrates, and the patterns of inhibition of the lipid-modified
and the unmodified pulullanases were also the same. However, we found thatthe wild-type pullulanase formed trimers whereas the unmodified enzyme did
not, and that the migrations of the two enzymes on sodium dodecyl sulphateelectrophoresis were different when the samples were applied on the gel
without heating . The results presented in this paper and in previous work
show that the correct processing and translocation of pullulanase in K.

aerogenes require modification of lipid . However, the enzymatic properties
and physical stabilities of pullulanase were not affected by the lipid
modification .

77

Refarences
Bender, H., and Wallenfels, K. (1961) Untersuchungen an Pullulan , II.
~.

Spezifisher Abbou durch ein Bacterielles. Enzym Biochem Z 334 : 79-95.
Charalambous, B., Kenn, j. N., and McPherson, M. J. (1988) Collagen-likesequences stabilize homotrimers of a bacterial hydrolase. EMBO J 7 : 29032909.
d'Enfert, C., Chapon, C., and Pugsley, A. P. (1987) Export and secretion of
lipoprotein pullulanase by Klebsiella pneumoniae. Mol Microbiol 1: 107116.
Eisel, E., Roshed , I. R., and Wallenfels, K. 1972 Molecular chracterization ofpullulanase from Aerobactor aerogenes. Eur J Biochem 26 : 62-67.
Inouye, S., Wang , S., Sekizawa, J., Halegoua, S., and Inouye, M. (1977)
Amino acid sequence of peptide extension on the lipoprotein ofEscherichia coli outer membrane. Proc Natl Acad Sci USA 74 : 1004-1008.
Katsuragi, N., Takizawa, N. , and Murooka, Y. (1987) Entire nucleotide
sequence of pullulanase gene of Klebsiella aerogenes W70. J Bacteriol
169: 2301-2306.
Konishi , Y., Amemura, A., Tanabe, S., and Harada, T . (1979) Imunologicalstudy of pullulanase from Klebsiella strains and the occurence of thisenzyme in the Enterobacteriaceae. Int J Syst Bacteriol 29 : 13-18.
Kornacker, M. G., and Pugsley, A. P. (1989) Molecularchracterization and its
product, pullulanase, a secreted enzyme of Klebsiella pneumoniae
UNF5023. Mol Microbiol 4 : 73-85.
Laemmli, U. K. (1970) Cleavage of structural proteins during assembly of thehead of bacteriophage T 4. Nature 224: 680-685.
Lowry, O. H., Rosenbrough, N. J., Farr A. L., and Randall, R. J. (1951) Protein
measurement with the Folin phenol reagent. J Bioi Chem 193: 265-275.
Murooka, Y. and Ikeda, R. (1989) Biosyhthesis and secretion of pullulanase, alipoprotein from Klebsiella aerogenes. J Bioi Chem 264: 17524-17531 .

78

Nelson., N. (1944) A photometric adaptation of the Somogi method for thedetermination of glucose. J Bioi Chem 153: 375-380.
Ohba, R., and Ueda, S. (1973) Purification, cystallization and some properties
of intracellular pullulanase from Aerobacter aerogenes. Agri Bioi Chem 37 :2821-2926.
Pugsley, A. P., Chapon, C., and Schwartz, M. (1986) Extracellular pullulanaseof Klebsiella pneumoniae is a lipoprotein. J Bacteriol 166: 1083-1088.
Pugsley, A. P., Poquet, I., and Kornacker, M. G. (1991) Two distinct steps inpullulanase secretion by Escherichia coli K12. Mol Microbial 5 : 865-873 .
Pugsley, A. P., and Reyss, I. (1990) Five genes at the 3' end of the Klebsiellapneumoniae pulC operon are required for pullulanase secretion . Mol
Microbiol 4 : 365-379.
Reyss, I., and Pugsley, A. P. (1990) Five additional genes in the pulC-O
operon of the Gram-negative bacterium Klebsiella oxytoca UNF5023 which
are required for pullulanase secretion . Mol Gen Genet 222 : 176-184.
Schnaitman, C. A. (1973) Outer membrane proteins of Escherichia coli. Arch
Biochem Biophys 157: 541-552.
Takizawa, N., and Muraoka, Y. (1984) Intergeneric transfer of the pullulanasegene between Klebsiella aerogenes and Escherichia coli by in vivo genetic
manipulation . Agri Bioi Chem 48 : 1451-1458.
Takizawa, N., and Murooka, Y. 1985 Cloning of the pullulanase gene andoverproduction of pullulanase in Escherichia coli and Klebsiellaaerogenes. Appl Environ Microbiol 48 : 294-298.

79

·
'

Chapter IV

Functinal Development of Pullulanase :Random mutagenesis of Pullulanase from Klebsilella aerogenes for Studies of
the Structure and Function of the Enzyme

Abbreviations: a-CD a-cyclodextrin (cyclohexaamylose); tl-CD, tl-cyclodextrin
I(cyclo heptaam ylose).

80Introduction
Pullulanase [EG 3.2.1.41], which was first isolated by Bender and Wallenfels (1)4 •from Klebsiella aerogenes (formerly Aerobacter aerogenes), is one of the starchdebranching enzymes that are widely distributed in plants (2) and bacteria (3-6).
This enzyme hydrolyzes a-(1-6)-glucosidic linkages and is commonly used in the
starch industry for the production of dextrose and high-fructose corn syrup at hightemperature. Two initial steps are involved in the process: first, liquefaction, which
requires a very thermostable a-amylase; and second, saccharification, in which
pullulanase in addition to glucoamylase or ~-amylase generates dextrose with ayield of greater than 94%. For the latter step, pullulanase should be compatible
with the other enzymes in terms of pH optimum and thermostability. Thisrequirement for the adaptability of pullulanase attracted our interest to thestructure and function of this enzyme.
Extensive studies have been reported of pullulanases from K aerogenes (7-11)
and Klebsiella pneumoniae (12, 13). The genes for pullulanase (pulA) from K.

aerogenes W70 (14, 15) and K. pneumoniae (12) have been cloned andsequenced, but most studies have focused on the secretory pathway in Gramnegative bacteria (16, 17). Little information about the structure and function ofpullulanases has been reported (13). Additional pullulan-hydrolyzing enzymes(a-amylase-pullulanases) that hydrolyze both a-(1-4) and a-(1-6)-glucosidic
linkages have recently been described (18-22). Plant et al. (23) suggested thatthe same active center might be involved in the dual activity toward a-( 1-4)- anda-(1-6)-glucosidic linkages. Kuriki and Imanaka (24) reported the existence offour highly conserved regions in a-amylases, pullulanases, isoamylases [EG 3. 2.
1. 68], cyclodextrin glucanotransferases [EG 2. 4. 1. 19], and neopullulanases thathydrolyze not only the a-( 1-4 )-glucosidic linkages but also the a-( 1-6)-glucosidic
linkages of several branched oligosaccharides. These four conserved regions
most likely constitute the active center of each enzyme. Kuriki et al. (25) also
reported

that

one

activecenterinneopullulanasefrom

Bacillus81

stearothermophilus participated in the dual hydrolytic activity toward a-(1 -4)- and.

a-( 1-6)-glucosidic linkages .
.

To date, two eukaryotic a-amylases, namely, Taka-a~ylase A of Aspergillus
oryzae (26) and a-amylase from pig pancreas (27), have been studied by X-ray
crystallography. However, no three-dimensional structure of any pullulanase has
been reported . A rational approach to enzyme design involving site-directedmutagenesis is inefficient in the absence of detailed structural information orwhen the molecular basis for the property of interest is poorly understood. In suchcases, random mutagenesis combined with selection or screening can be auseful alternative for generating both the desired improvements and a data basefor future rational approaches to protein design. Random mutagenesis has beenused to enhance or alter various features of enzymes, such as thermal stability(28-33), stability to alkali (34), and substrate specificity (35) , as well as to restore
the catalytic activity of an enzyme damaged by site-directed mutagenesis (36).
Here we report seven variants of pullulanase that were generated by randommutagenesis and selected by screening at high temperature. Characterizationand localization of the amino acid substitutions provided information useful forfurther improvements in the enzyme's function .

82

Material and methods
Materials and Chemicals - Restriction endonucleases, T 4 DNA ligase, AMVj'reverse transcriptase, and M13 primer were purchased from Takara Shuzo Co.
Ltd . (Kyoto) or Toyobo Co. Ltd. (Osaka). Pullulan (average molecular mass, 99,8
kDa) was provided by Hayashibara Co. Ltd. (Okayama) . a-CD and ~-CD were
gifts from Dr. J . Abe (University of Kagoshima). Reactive Red 120 was purchasedfrom Sigma Chemical Co. (St. Louis, MO, USA). Reactive Red 120-derivatizedpullulan (PRR) was prepared by the method of Rinderknecht et a/. (37). All otherreagents were standard commercial preparations.
Bacterial Strain, Plasmid, and Phages - The following bacterial strain, plasmid ,and phages were used; Escherichia co1iJM109, pUC18,and M13mp18 and mp19
(38) .

Manipulation of DNA - Procedures for purification of DNA, restriction enzyme
digestion, ligation, agarose-gel electrophoresis, and transformation of competent
Escherichia coli cells were carried out as described by Sambrook et al. (39). DNA

sequencing was carried out by the dideoxy chain -termination method (40).
Random Mutagenesis - A convenient expression vector for pullulanase, pMP10
was constructed with a 2.3-kb EcoRl-Aat II fragment that consisted of the origin of
replication and the gene for ampicillin resistance from pUC18, and a 5.5-kb
EcoRI-Aat II fragment that contained the structural gene for pullulanase (pulA)

from pMPK1 (16). The pulA gene was divided into four fragments of similar length(Fig. 4), and each fragment was subcloned into phage vectors M13mp18 (coding
strand) and mp19 (noncoding strand). Random mutagenesis of the cloned DNA
was performed by the method of Myers et al. (41). In brief, single-stranded DNA ofthe hybrid plasmid was treated at 20°C with either hydrazine (for 5-10 min),formic acid (for 5-10 min) or sodium nitrite (for 1-1.5 h). The mutagenized singlestranded DNA was annealed with the M13 primer and a duplex was generated byreverse transcriptase with dNTPs. The duplex DNA was digested with appropriate
restriction endonucleases. The resulting fragments that contained part of pulA

83

were inserted into pMP10. E. coli JM 109 was transformed with the hybrid
pfasmids and selection was performed after plating on LB medium (39) that~.

contained agar and ampicillin (100 f..lg/ml).
Screening for Mutant Pullulanases - Fresh transform ants were replicated on
MM agar medium to monitor production of pullulanase (42). Cells were grown at
37°G for two days and incubated at 700G for 1 h until the wild-type pullulanase
activity was lost. The plates were overlaid with PRR (0.5 %) soft agar. Expressionof mutated pulA genes that were functional in E. coli was detected as haloes
around individual colonies, whereas colonies of cells with an inactivatedpullulanase do not form haloes (43). The positive clones were purified byrepeated streaking on M M agar plates to confirm the expression of active
pullulanase. Mutations generated by random mutagenesis were identified by
DNA sequencing.
Production and Purification of Pullufanases - E. coli JM 109 cells carrying aplasmid that included a wild-type or mutated pulA gene were cultured in MM
medium . Wild-type and mutant pullulanases were purified as described by
Yamashita et a/. (44).
Assay of Pullulanase Activity - Pullulanase activity was measured by amodification of the method of Takizawa and Murooka (42). A 1.4-ml reaction
mixture consisting of enzyme and a 0.5% solution of pullulan in 0.1 M acetate
buffer (pH 6.0) was incubated for 10 min at 50 0G and the release of aldehydegroups from pullulan was measured by the method of Somogyi (45) and Nelson(46). One unit of activity was defined as the amount of enzyme that caused theliberation of 1 f..lmol of aldehyde groups per min. For the kinetic analysis ofhydrolysis of pullulan, initial rates were determined by linear regression analysis,
and the kinetic parameters were calculated from Eadie-Hofstee plots (47). Protein
contents were measured by the method of Lowry et al. (48) with crystalline bovine
serum albumin as the standard. The absorbance at 280 nm was used to monitor
protein in column effluents.

81

Results
Screening for and Isolation of Mutant Pullulanases - In the present experiment,,.

when individual transformants (a total of about 50,000 colonies) were analyzed on
the plates, seven clones were positive for pullulanase activity after incubation at
70°C for 1 h. All of them were obtained from the formic acid treatment (about
20,000 transformants). Plasmid DNAs were isolated from these clones.
The DNA sequences of both strands of each fragment from the seven plasm ids
were determined. Table 1 summarizes the base substitutions in the gene forpullulanase that were caused by chemical mutagenesis. Among 17 basesubstitutions in seven kinds of mutant pullulanase, one was a transition and 16
were transversions. Of the 16 transversions, 14 were transversion from a purine toa pyrimidine. In this experiment, we mutagenized both the coding and thenoncoding strands of the fragment using M13 mp18 and mp19, respectively. This
procedure may have increased the variety of mutant genes. Of 17 base
SUbstitutions generated during mutagenesis, 13 resulted in amino acidreplacements, while the other four did not (Table 1). None of the seven mutant
pullulanases showed any change in secondary structure calculated according toChou and Fasman (49) (data not shown).
Amino Acid Substitutions Clustered in a Single Region - The clones thatformed haloes on PRR plates were isolated and cells were cultured in MM
medium. Crude preparations of enzymes from these cultures were prepared andassayed for pullulanase activity at 70°C. No activity was found in cells with either
the wild-type or the mutant enzymes at 70°C. The increased thermostability of themutant enzymes was confirmed by measuring the residual activity after heat
treatment of solutions of crude enzyme at 50°C for 30 min (Table 1). Under these
conditions, the residual activity of the wild-type enzyme was 18.0% of the initialactivity. Most mutant pullulanases had higher residual activity than that of thewild-type enzyme, whereas other mutant enzymes had the same or slightly lower
activity than that of the wild-type enzyme, although they had been selected for

85

~

00

Pro 403 Gin
Leu 482 Gin
Val 492 Val
Ala 493 AlaAla 159 Glu
Ala 165 Ala
Ser 332 Arg
Gin 317 Leu
Thr 484 Ser

GGC to GTC
GAC to GTC
CAG to CAT
CGG to CGT

CGC to CTC
CGG to CGT
TCA to TCT

CAG to CTG
ACG to TCG

GCC to GCA
TIC to ATC

M13mp19M13mp19M13mp18

M13mp18

1-62

1-75

1-92

1-97

32.7

37.4

15.9

11. 6

23.9

22.4

33.4

109

23.7

44.0

67.5

Specific activity (U/mg)

Nucleotide changes are shown in the strand used for chemical mutagenesis with the mutagens, as described in the text.
M13mp18 and mp19 phages carried the coding and noncoding strands of the pulA gene, respectively; the cod ing strand
is the DNA strand whose nucleotide sequence is the same as that of the mRNA for the enzyme. Amino acid sequences
run from the NH 2-terminus. The values indicate the amount of activity remaining after heating at 50°C for 30 min as apercentage of the activity after incubation at 4°C for 30 min. The activities were measured in duplicate and the two determinations were in agreement to within 200/0 under the conditions described in Materials and methods.

Ala 265 Ala
Phe 562 lie

43.7

Leu 409 Gin
Phe 562 lie

GAC to GTC
AAG to TAG

M13mp19

I-56
23.4

19.4

Ala 515 Ser
Thr 617 lie

GCG to TCG
ACC to ATC

M13mp18

1-35

Val 441 Leulie 600 Phe

GTG to TIG
ATI to TIT

31 .3

Residual activity (%)

M13mp18

Amino acid substitution

1-27

Nucleotide change
18.5

Phage

Wild type

StrainTable 1. Base changes induced by chemical mutagenesis and deduced amino acid substitutions encoded by the mutant genes
for pullulanase and specific activities of the purified wild-type and mutant pullulanases.

thermal stability of the enzyme on PRR plates. The amino acid substitutions were
all located within only one of four fragments of the pulA gene and no substitutions,.
~

were detected in any other fragment. Among 12 kinds of amino acid substitution,
10 were substitutions of noncharged amino acids. Substitutions of two amino acidresidues in mutant 1-75 resulted in introduction of charged amino acids. In the
mutant I-56, Phe562 was replaced by lIe562 as in mutant 1-97, in addition to thereplacement of Leu409 by Gln409.
Characterization of Mutant Pullulanases - Wild-type and mutant pullulanases in
the cytoplasmic space were purified about 100-fold. Each purified enzyme
migrated as a single protein band during SOS-PAGE by the method of Laemmli(50). The molecular mass of each enzyme was determined to be 120 kOa (datanot shown). The specific activities of the purified enzymes were measured (Table1). The following analyses were performed with purified enzyme samples at a
final concentration of pullulanase of 1 U/ml .
The optimum pH and pH stability of the seven mutant pullulanases werecompared with those of the wild-type enzyme. The optimum pH of the Wild-type
and all the mutant pullulanases was pH 6.0 (data not shown). The mutant
enzymes also displayed pH stability similar to that of the Wild-type enzyme (44).
Although six of the seven mutant pullulanases had the same optimum
temperature (55°C) for activity as that of the wild-type enzyme, the 1-97 mutant
pullulanase had maximum activity at 60°C (Fig. 1). To test the thermal stability ofthe mutant enzymes, residual activities Were measured after incubation at various
temperatures for 30 min. The residual activity of the 1-27, I-56, 1-92, and 1-97
mutant pullulanases ranged from 4 to 100/0 after incubation at 55°C for 30 min, but
no activity was detected in the case of the wild-type and the 1-35, 1-62, and 1-75
mutant pullulanases (Fig. 2). Since the kinetics of irreversible deactivation of an
enzyme reflect its thermodynamic stability (51), the time courses of irreversible
thermal inactivation of the mutant pullulanases were determined (Fig. 3). The
rates of thermal inactivation of the 1-27, I-56, 1-92, and 1-97 mutant pullulanases

87

00
00

Q)

<U

cr:

30

oI

20

40

40

50

60

[ 11--..,....---;

80
30

o I

20

ca 40

Q)

cr:

40

60

Temperature (OC)50~

Fig. 1. Temperature-activity profiles of wild-type and mutant pullulanases.
(a) 0 , Wild type; • , 1-27; 11 , 1-35;' , I-56; 0 , 1-62.
(b) 0 , Wild type; . , 1-75; 11 , 1-92;' , 1-97.
Data points represent the means of three experiments.

Temperature (OC)

70

Q)

>

.';:::;

Q).';:::;>·u<U 60·s"-·u<U 60

"-

~
.....

lOa

.....~
·s

(a)

~
80
0

D..

120

~80
0

100 ~

120

70

80

, )~-..,....----1

(b)

~

00

~

-

a:

Q)

'w

20

30

40

so

:"{ ]t--_____--,,...---I

70

10

20

40

so
Temperature (OC)

30

60

70

~)~~,.~--~--~

Fig. 2. Thermal stability of the enzymatic activity of wild-type pullulanase and mutant pullulanases.
The enzyme in 0.1 M acetate buffer (pH 6.0) was incubated at various temperatures for 30 min.
The residual activity was measured as described in the text. (a) 0, Wild type; e, 1-27; 11 , 1-35;" I-56; tI, 1-62. (b) 0 , Wild type;e , 1-75; 11, 1-92; , , 1-97.
Data points represent the means of three experiments.

Temperature (OC)

60

o I

10

oI

40

20

a:

'w
Q)

20

40

::J

"0

"0

::J

co 60
co

0

'';::;

~

-

'5

co 60
co

'13

'5

0

~ 80

-

100

100

~ 80

0

-

120------------------------------------~

120~----------------------------------------~

i'120------------------------------------~

100

--~
0

-

80

>.

.s:
t=s«S

60

«S

:::J

U

'w
Q)
a:

40

20

O+---~--~----~--~--~--~----~--~

o

10

20
30
Incubation time (min)40

Fig. 3. Kinetics of thermal inactivation of wild-type and mutant pullulanases.
The pullulanases were incubated at 50°C in 0.1 M acetate buffer (pH 6.0)
for the indicated times and residual activities were determined as describedin the text.
0, Wild type; ., 1-27; Il, I-56; ., 1-92; 0, 1-97.
Data points represent the means of three experiments.

9()were found to be two to four times lower than that of the wild-type enzyme. These
results show that these mutants are more stable than the wild-type enzyme and,:Ithat the 1-97 mutant pullulanase has an altered optimum temperature andenhanced thermal stability (Fig. 1-3).
The substrate specificities of the wild-type and mutant pullulanases were
examined (Table 2). The 1-35 mutant pullulanase was more effective against
amylopectin than the wild-type enzyme, whereas the 1-27 and 1-75 mutant
pullulanases were less effective against amylopectin. The activities of 1-35, 1-62,
and 1-75 mutant pullulanases against soluble starch were lower than that of thewild-type enzyme. Neither the Wild-type nor the mutant pullulanases hydrolyzed
amylose, suggesting that none of the mutant pullulanases could hydrolyze a-( 14)-glucosidic linkages.
The effects of a-CD and ~-CD as competitive inhibitors of pullulanase (47) on
the activities of the Wild-type and mutant pullulanases were examined (Table 3).
Concentrations (w/v) of a-CD and ~-CD for 50% inhibition of the Wild-type
pullulanase activity were determined to be 0.01 % and 0.0004%, respectively. At aconcentration of 0.01 % a-C~, five of the mutant pullulanases, but not I-56 and 197, were inhibited about 8 to 15% more than the Wild-type enzyme. These mutant
pullulanases were also inhibited about 15 to 37% more than the Wild-type
enzyme by 0.0004%~-CO. Since the 1-35 and 1-75 mutant pullulanases weredifferent from the wild-type enzyme in the substrate specificity and inhibition ratio,these mutant pullulanases appear to have altered affinity for the substrate andinhibitors.
Effects of Mutations on the Catalytic Activity of Pullulanase - Kinetic parameters
were determined for the purified wild-type and mutant pullulanases (Table 4). No
significant difference in the catalytic rate constant (kcat) between the mutant andwild-type enzymes was observed. The kcat values were all within a range of ±50%. On the other hand, the Michaelis constants (Km) of the 1-27, 1-35, 1-62, 1-75,
and 1-92 mutant pullulanases were two to seven times greater than that of the

9.1

4 .

Table 2. Comparison of the substrate specificities of wild-type and mutant pullulanases.
Relative activity (%)
Substrate
Pullulan
Amylopectin
Soluble starch
AmyloseWild type

1-27

1-35

I-56

1-62

1-75

1-92

1-97

100
42.9
19.8

100
33.9
19.8
0

100
52.3
16.5
0

100
44.8
22.1

100
40.2
16.4

100
38.7
16.9

100
43 .8
25.4

100
41 .1
23.3

0

0

0

0

0

0

Enzymatic activities were calculated from the amount of released reducing sugars relative
to that of total sugars in each molecule. The activities were measured in duplicate and thetwo determinations were in agreement to within 15% under the conditions as described inMaterials and methods. The values indicate the relative activity as a percentage
with each substrate compared to that with pullulan .

~.

Table 3.

Effects of a- and (3-CDs on pullulanase activityof the wild-type and mutant enzymes.
Residual activity (%)
EnzymeWild type
1-27
1-35
I-56
1-62
1-75
1-92
1-97

ABC
100
100
100
100
100
100
100
100

50.0
39.2
37.8
53.8
37.8
34.7
42.2
48.6

50.0
34.8
16.4
53.1
25.5
12.7
18.9
57.6

A, Pullulan 0.5% ; B, pullulan 0.5% + a-CD 0.01 % ;C, pullulan 0.5% + (3-CD 0.0004% .
The activities were measured in duplicate and the twodeterminations were in agreement to within 200/0 under
the conditions described in Materials and methods.The
values indicate the activity remaining in the presenceof inhibitor as a percentage of the activity without any
inhibitor.

Table 4. Comparison of kinetic constants for the reactions of wild -type andmutant pullulanases with pullulan .

..
kcat I Km(M- 1• s-1.1 0-6)

fl.fl. G **(kcal • mol- 1)45(100)a0.73(100)a

0

47
(142)

97
(216)

0.48
(66)

0.27

1-35

18
(55)

313
(696)

0.06
(8.2)

1.60

I-56

31
(94)

35
(78)

0.89
(122)

- 0.13

1-62

49
(148)

130
(289)

0.38
(52)

0.42

1-75

35
(106)

167
(371)

0.21
(29)

0.80

1-92

32
(97)

87
(193)

0.37
(51 )

0.44

1-97

34
(103)

31
(69)

1.10
(151 )

-0.26

kcat
(S-1)

Km
(JiM)

Wild type33(100)a1-27

EnzymeResults are means of results obtained with 10 concentrations of substrate. Kinetic parameters, namely, kcat, Km, were determined in duplicate and the two determinations werein agreement to within 100/0 under the conditions described in Materials and methods.
a The values in parentheses are percentages based on value for the wild-type enzyme.

94

Wild-type enzyme, whereas those of the I-56 and 1-97 mutant pullulanases werelower than that of the wild-type enzyme. The higher Km values of the 1-35 and 1-75,.

mutant pullulanases were probably the cause of the chanme in substratespecificity, as seen in Table 2, and the extent of inhibition by a-CD and ~-CD, asseen in Table 3.
The specific activities of mutant pullulanases showed similar behavior to thepercentages of catalytic efficiency (kcat I Km) that based on the value for the wildtype enzyme, except for the 1-35 and 1-97 mutant enzymes (Tables 1 and 4).The
specific activities of the 1-35 and 1-97 mutant enzymes were four times higher andnine times lower than that of the wild-type enzyme, respectively. The I-56 and 1-97
mutant pullulanases showed 1.2-fold and 1.5-fold increases in catalytic efficiency(kcat I Km), respectively, compared to the wild-type enzyme (Table 4). Such
improvements were a result of their lower Km values (decreased by a factor of 1.3
and 1.5 compared to the wild type) but not due to the change in the kcat values(decreased by a factor of 1.1 and 1.0).
The effects of amino acid sUbstitutions on the catalytic efficiency can also bequantified in terms of the changes in activation energy (L\L\G**) for pullulanhydrolysis (52). The changes in activation energy calculated from the relationship,{L\L\G**=- RTln [(kcat I Km)mutant I (kcat I Km)wild-type]), are shown in Table 4. The

effect of the amino acid substitutions in I-56 and 1-97 was to decrease theactivation energy (relative to the wild type) by 0.13 and 0.26 kcall mol (1 cal =4.184 J), respectively, whereas the mutant 1-35 pullulanase was associated with amarked increase in L\L\G**.

Discussion
In this report we have described a method of random mutagenesis for creating
functional pullulanases with altered enzymatic properties. Key features of theapproach are the incorporation of random amino acid substitutions bymutagenesis and a technique for appropriate selection of or screening for thedesired variants. We tried to identify mutations that might increase the thermal
stability of pullulanase. Temperature-sensitive or -resistant mutations provide one
way of identifying residues and interactions that contribute to the folding andstability of a protein . The screen for mutants described here is based on a plate
enzyme assay for the thermal stability of pullulanase. A large number of enzymes
including protease, amylase and protein can be assayed on Petri plates. The
susceptibility of a protein to thermal stability might be detected by using asensitive colored substrate. Consequently, the method that we have outlined here
may be applicable for obtaining useful variants of other enzymes. After we had
analyzed about 50, 000 mutagenized colonies, we obtained seven kinds of mutant
gene. Five mutant pullulanases had altered catalytic properties. The amino acidsubstitutions caused an increase in Km values while having a less significant
effect on kcat values. The amino acid sUbstitutions of the five mutants were
distributed on the NH2-terminal side of the four conserved regions (Fig . 4) . A
significant change in activity with substrates, such as amylopectin and soluble
starch, and in response to competitive inhibitors, such as a- and ~-CD , wasobserved for the 1-35 and 1-75 mutant pullulanases, and may also be associatedwith the increased Km value. In the 1-75 mutant pullulanase, amino acids werechanged from hydrophobic or neutral residues to hydrophilic residues. The
mechanism of the effect on catalytic function of these amino acid sUbstitutions
may involve hydrophobic interactions.
Iwamoto et al. (47 and personal communication) suggested that the pullulanaseof K. aerogenes has an additional substrate-binding site, with low affinity for
pullulan, over and above the conserved binding regions found in a -amylases. A

96

·

.

Asp602 HIs607""~________~D~D~D~D~__~lcOOH
EcoRI _~~-lttn- Sphl

,

,
,,

1-27

Kpnl_~______lttn_Smal_ _ _.-..lttns.......
Hind"lHind"l~Smal ___- - Sphl.........

-.... ........ -- ......

....

aJ:41

_- ........

V
lIe~o
Leu - - - - - P h e

1---------1----------1-------- "

0 --f

AI~15

1-35

Ser

Leu409

I-56

Th~17

Phe562"

Gin - - - - - l I e Pro403
Leu482"

1-62
Ala159

"

0

lIe-l

0 ---I"G l n - - G l n - - - - - D ---I
Ser332"1-75 t---Glu-Arg - - - - - - - - Gln317
Thr484""

1-92 I-----Leu - - - - - Ser - - - - Phe562"

1-97 1 - - - - - - - - - - - - - l I e -0 ---I

0 ---I
0 ---I

Fig. 4. Positions of amino acid substitutions in mutant pullulanases. Amino acidsequences are shown from the mature NH 2 -terminus. Each rectangle indicates
one of four conserved regions in members of the amylase family. Amino acidon the left rectangle are numbered from the mature NH 2 -terminus of pullulanse.
Arrows indicate restriction sites and the fragment of the pulA gene that was
mutagenized . The wild-type enzyme consists of a total of 1,083 amino acidresidues.

97

deletion pullulanase, with 122 amino acids deleted from the NH2-terminus, had
the same activity as the wild-type enzyme (Takizawa, N., personal communi~.

cation). In our preliminary experiments, a deletion pullular;lase, with 133 aminoacids deleted from the NH2-terminus, had lower activity than that of the Wild-type
enzyme. The amino acid substitutions caused by the present random mutation
started at the 159th amino acid from the NH2-terminus in the mutant 1-75 andended at the 617th amino acid in the mutant 1-35. These results suggest that the
NH2-terminal region of the pullulanase of K. aerogenes is important for thecatalytic function.
Nakajima et al. (53) found that the amino acid sequences of a-amylases,
irrespective of origin, have four highly conserved regions. Each a-amylase
contains an eight fold (f3/a)s barrel-supersecondary structure, that is, a barrel ofeight parallel f3-strands surrounded by eight a-helices (26). Although all aamylases have the same catalytic function, their amino acid sequences are quite
varied. We found that the pullulanase of K. aerogenes also has a conserved
eight fold (f3/a)s barrel structure as indicated by its amino acid sequence (15). The
molecular mass of pullulanase is at least twice those of a-amylases, and thepullulanase of K. aerogenes has a particularly long NH2-terminal region beyond
the four conserved regions. No amino acid sequences or secondary structure in
pullulanase and a-amylases were found to be homologous except for those in thefour conserved regions. Effective amino acid sUbstitutions generated by randommutagenesis were located on the NH2-terminal side of the four conserved
regions. The colonies with enzymes containing these four conserved regions (Fig.
4, Kpnl-Smal) that had been randomly mutated could not grow in MM medium(data not shown). These colonies may produce an inactive pullulanase. Twoamino acid residues (His607 and AspS77) of pullulanase of K aerogenes in thefour conserved regions were individually replaced by site-directed mutagenesis
to probe their roles in catalysis (54). None of the mutant pullulanases had any
activiy, although the enzymes were detected on SDS-PAGE. The mutations 1600F

9H

in mutant 1-27 and T6171 in 1-35 are very close to the predicted substrate-binding..

site of pullulanase (Fig . 4). The Km values of the 1-27 and 1-35 mutant
pullulanases were higher than that of the wild-type enzyme. These results seemto support the hypothesis that the pullulanase of K. aerogenes has active sites
similar to those of a-amylases with the four conserved regions.
One type of mutant pullulanase exhibited increased thermal stability. Although
the screening process after random mutation was designed to enhance thethermal stability of the enzyme, not all of the seven mutant pullulanases exhibitedenhanced thermal stability. Since E. coli does not secrete pullulanase into themedium, bacterial colonies selected on PRR plates probably protected
pullulanase from thermal inactivation . A distinct enhancement of the thermostability of this pullulanase was seen upon the replacement of Phe562 by lie in
the 1-97 mutant pullulanase. In general, a globular protein is stabilized byhydrophobic residues in its interior (55) . Further work is in progress to define thecatalytic sites of pullulanase. To investigate the effect of particular amino acidresidues at position 562 on the thermostability of the enzyme, we plan to replace
the Phe residue at position 562 of pullulanase with all possible amino acidresidues by site-directed mutagenesis.

9f)

Summary..

To study the structure and function of pullulanase from Klebsiella aerogenes, amethod involving random mutagenesis of the entire gene 'for pullulanase wasused. Out of 50,000 clones screened at high temperature, seven genes for mutant
proteins were identified by DNA sequencing. The amino acid substitutions in theseven mutant proteins were clustered on the NH2-terminal side of the four
conserved regions found in a-amylases. These mutant pullulanases wereclassified into two types: those whose catalytic activity was altered and those
whose thermal stability was increased . The results presented here and inprevious reports suggest that pullulanase from Klebsiella aerogenes has similar
active sites to those of a-amylases with the four conserved regions, as well as
another substrate-binding site closer to the NH2-terminus. The plate assay
method used for isolation of thermostable variants may be applicable to thegeneration of useful variants of other enzymes.

10()

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Biochem. J. 246, 537-541
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44. Yamashita, M., Nakagawa, A., Katuragi, N., and Murooka, Y. (1992) Role oflipid modification on a starch-debranching, Klebsiella pullulanase-comparisonof properties of lipid-modified and unmodified pullulanase. Mol. Microbiol. 6,389-394
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Takahashi, K., Hashimoto, H., and Sakai , S. (1993) Interaction between
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10S

GENERAL DISCUSSION (CONCLUSION)
In this thesis,· the author presented the results of the study 'on the productivity,of homologous and heterologous gene products, analysis of structure-function
relationships, and functional development. The conclusions of this work aresum marized as follows.
(1) An example of heterologous expression was chosen as a project toproduce a sulfhydryl (SH)-rich protein, human metallothionein II (hMT -II) andillustrate approaches in the genetic design of Escherichia coli that can act asoverproducer. The gene of hMT -II was cloned into a vector, pUEX2 andexpressed as a fusion protein between ~-galactosidase and hMT-11. The
fusion protein accounted for approximately 24% of the total protein in the cells.
The fused hMT -II protein was bound to cadmium. Thus, the successful
development of chimeric cells that overproduced an active human protein wasachieved.
(2) The practical aspect was focused on growing a recombinant Klebsiellaaerogenes strain in fermentation culture and purifying the recombinant protein
product. The gene for membrane-bound monoamine oxidase (maoA) from K.

aerogenes W70 was cloned and the enzyme overproduced in a soluble form .
The enzymological properties and the amino acid sequence of the enzyme
deduced from the nucleotide sequence of the maoA gene were different fromthose of known tyramine or monoamine oxidase. The enzyme is a new type of
monoamine oxidase that requires Cu and Topaquinone as cofactors. The
purified enzyme was crystallized and the structure analyzed by X-ray(3) To analyze secretion and structure-function relationships, an extracellular protein and starch-processing enzyme, pullulanase of K aerogenes,
was used. First, to clarify the role of lipid modification of pullulanase, lipidmodified wild-type and the unmodified pullulanases were purified and theirproperties compared. The enzymatic properties and physical stabilities ofpullulanase were not affected by the lipid modification. The modification oflipid on pullulanase in K. aerogenes was a signal of correct processing andtranslocation similar to glycosylation typical of eukaryotic cells.
~.

(4) To analyse the structure and alter the function of pullulanase, a method
involving random mutagenesis was used. Seven genes for mutant proteins
were identified . These mutant pullulanases were classified into two types :those with altered catalytic activity and those with increased thermal stability.
The pullulanase from K aerogenes has active sites similar to those of a amylases with four conserved regions, as well as other substrate-binding site
closer to the NH2-terminus.

10 8

RELATED PUBLICATIONS
1. Genetic Design of a Gene for Human Metallothionein II and Its expression,

as an Active Fusion Protein in Escherichia coli.
M. Yamashita, H. Kuwata, H. Murakami, and Y. Murooka.
J. Ferment. Bioeng.,77, 113-118, (1994).
2. Purification, Characterization and Crystallization of Monoamine Oxidasefrom Escherichia coli K-12.
J. H. Roh, H. Suzuki, H. Azakami, M. Yamashita, Y. Murooka, and H.
Kumagai.
Biosci. Biotech. Biochem., 58, 1652-1656, (1994) .
3. Crystallization and Preliminary X-ray Analysis of Copper Amine Oxidasefrom Escherichia coli K-12.

J. H. Roh, H. Suzuki, H. Kumagai, M. Yamashita, H. Azakami, Y. Murooka,
and B. Mikami.
J. Mol. BioI., 238, 635-637, (1994) .
4. Purification and Characterization of Monoamine Oxidase from Klebsiellaaerogenes.
M. Yamashita, M. Sakaue, N. Iwata, H. Sugino, and Y. Murooka.

J. Ferment. Bioeng ., 76, 289-295, (1993).
5. Fundamental Aspects of Secretory Enzyme Production by Recombinant
Microbes.
N. Takizawa, M. Yamashita, and Y. Murooka.
RECOMBINANT MICROBES FOR INDUSTRIAL AND AGRICULTURAL
APPLICATIONS. Marcel dekker Inc., 271-290, (1994).
6. Role of lipid modification on a starch-debranching enzyme, Klebsiellapullulanase; comparison of properties lipid-modified and unmodified
pullulanase.
M. Yamashita, N. Nakagawa, N. Katsuragi, and Y. Murooka.
Mol. Microbiol., 6, 389-394, (1992).

109

7. Random mutagenesis of Pullulanase from Klebsiella aerogenes for Studiesof the Structure and Function of the Enzyme.
M. Yamashita, T. Kinoshita, M. Ihara, T. Mikawa, and Y. fv1urooka.
J. Biochemistry, 116, 1233-1240, (1994) .

110

REFERRENCE PUBLICATIONS* Necleotide S~equence of the Gene for Monoamine Oxidase (maoA) from

Escherichia coli. H. Azakami, M. Yamashita, J. H. Roh, H. Suzuki, H. Kumagai,
and Y. Murooka., J. Ferment. Bioeng.,77, 315-319, (1994).

* Cloning Vectors Commonly Used for Recombinat Microorganisms. M.
Yamashita., in RECOMBINANT MICROBES FOR INDUSTRIAL AND
AGRICULTURAL APPLICATIONS. Marcel dekker Inc., 853-860, (1994).
* Bacterial cholesterol oxidase are able to act as flavoprotein-linked steroid

monooxygeneses that catalyse the hydroxylation of cholesterol to 4cholesten-6-01-3-one. I, Molnar, N. Hayashi, K. P. Choi, H. Yamamoto, M.
Yamashita and Y. Murooka., Molecular Microbiology, 7,419-428, (1993).
• A monoamine-Regulated Klebsiella aerogenes Operon Containing the
Monoamine Oxidase Structural Gene (maoA) and the maoe gene. H. Sugino,
M. Sasaki, H. Azakami, M. Yamashita and Y. Murooka. J. Bacteriol. 174,24852492 (1992).
* Gene cloning of the maoA and overproduction of a soluble monoamineoxidase from Klebsiella aerogenes. H. Sugino, K Ishibashi, M. Sakaue, M.
Yamashita, and Y. Murooka. Appl. Microbiol. Biotechnology 35, 606-610(1991 ).
* Chemical Synthesis, Cloning, and Expression of Genes for Human

Somatomedin C (Insulin-like Growth Factor I) and 59Val-Somatomedin C. M.
Niwa, S. Sato, Y. Saito, F. Uchiyama, H. Ono, M, Yamashita, T. Kitaguchi, Y.
Shiga, J. Notani, H. Yamada, Y. Ishii, I. Ueda and Y. Takagi. Ann. N. Y. Acad.
Sci. 469, 31-52 (1986).
·Use of lac gene Fusion to Study regulation of Tyramine Oxidase, Which is
Involved in Depression of Latent Arylsulfatase in Escherchia coli.
M. Yamashita, and Y. Murooka. Agri. BioI. Chem. 48, 1459-1470 (1984).

* In vivo cloning of Arylsulfatase-Tyramine oxidase genes in ree strains ofKlebsiella aerogenes. Y. Muraoka, M. Oka, M. Yamashita, M. Sugiyama, andT. Harada. Agric. BioI. Chern . 47,1807-1815 (1983) .

11 ~~ACKNOWLEDGEMENTS..
I would like to express my deep gratitude to Professor Ph. D. Yoshikatu
Murooka for his expert guidance throught my research work.
My profound gratitude and thanks are due to Professor Ph . D. Shirou Nagai,
Professor Ph. D. Tokichi Miyakawa, Professor Ph. D. Naomichi Nishio,
Associate Professor Ph. D. Tsutomu Morinaga, and Associate Professor Ph. D.
Nobonu Takizawa for their advices.
I am deeply indebted to all staff members of Biotechnology Department of
Fujisawa Pharmacetical Co. Ltd ., for their suggestions and discussions
throughtout my basic studies.
Thanks are also due to all staff members of Laboratory of Industrial
Microbiology of Hiroshima University for their kind support of this work.
My sincere thanks are due to my wife, Yoshiko Yamashita for her love andpatience.

11 ~