1. CHARACTERIZATION OF AN INTRACELLULAR
ESTERASE
FROM BACILLUS SUBTILIS
by
John Franklyn Riefle!, III
A thesis submitted to the faculty of
the Medical University of South
Carolina in partial fulfillment of
the requirements for the degree- o~
Master of Science in the College of
Graduate Studies.
G(W
/2-7. S
B~
RSS'3c
/77S-
c. ;(
Department of Basic and Clinical Immunol~gy and Microbiology
~ 197~5
APprO/~bY: /j.
11'//}~ j-'~,' I _ , ' /
.. -~. jii1"~. /5.~ '"1=Sc
Chairman, Advisory Comm~ttee
UA
2. ABSTRACT
JOHN FRANKLYN RIEFLER, III. Characterization of an
Intracellular Esterase from Bacillus subtilis (Under
the direction 9f Dr. THOMAS B. HIGERD).
Esterase A, obtained by sonic disruption of cells of
Bacillus subtilis SR22 (SpOA12; trpC2), was purified apprcxi~
mately 400 fold with a 50 percent yield, utilizing differ-
ential chemical and heating precipitation, ion-exchange
chromatography and gel filtration. Optimum activity with
ethyl acetate as the substrate occurred over a broad pH
range from pH 7.0 to pH 9.0. The pH stability of the enzyme
followed a similar profile. Sub s t r'at e s pee i f i cit y 0 f the
enzyme appears to reside in the c~rboxylic acid moiety of
the substrate. Esterase A was unable to hydrolyze the
amino acid esters that were tested. Values for Vmax and
l/Km were determined for three c~mmonly used esters and
revealed the following decreasing values: p-nitrophenyl
acetate> a-naphthyl acetate> ethyl acetate. The most
potent inhibitors detected of esterase activity were:
mercuric chloride) DFF, eserine and sodium fluoride.
3. ACKNOWLEDGEMENTS
I would like to thank my advisor, Dr. Thomus B. Higerd
for his patience, constructive criticisms and insistence
upon excellence in all areas of research and teaching. I
am also very grateful to my committee and in particular to
Dr. Ch r i s t ian Sc h wabe, who g e n e r 0 U sly g u v e 0 f his t i 111 e and.
equipment and served as a most valuable consultant through-
out this project. A special word of thanks is owed Miss
Patsy McFadden for her understanding, sense of hUTllor a:cd
encouragement.
4. iii
TABLE OF CONTENTS
Page
ACKN 0 WLED GE~-1EN rr S ..................... .... ....... ii
TABLE OF CONTENTS ... ...... . . . . . . 1;; • • • • • • • • • • • • " .
iii
LIST OF FIGURES • • • • • • • • &; • W • • • • • • • • • • • .. • • • • • • • • • iv
OF TABLES .... ..-. .......................... v
INTRODUC~lION ........................ ............ 1
MATERIALS AND METHODS .......................... 23
RESULTS ... ...................... ................. 32
DISCTJSSION ................................ ...... 55
LIST OF REFERENCES . .. ... ... .......................
7. INTRODUCTION
Sporulation as a Hod~el Differentiation System. For many
years, sporulation of Bacillus ~. has served as a model
system for the study of cell differentiation~ vIi th a
thorough knowledge of the intracellular events that
accompany the process of spore formation, i.~. the means
by which a bacterium is converted from the vegetative
state to a dormant spore state, it may be possible to
partially or completely understand the factors responsible
for forming specialized cells in higher organisms (Hanson
et a1__ ,' 1970; Szulmaj ster, 1973).
There are several important differences between
bacterial and mammalian cells. Eucaryotic cells have a
much greater amount of genetic information, contained in
several chromosomes rather than the limited amount of
deoxyribonucleic acid (DNA) in the single chromosome of
procaryotic cells. Furthermore, each somatic cell in
eucaryotes is diploid which results in two forms (alleles)
of each. gene; whereas bacteria behave as haploid cells.
In addition, non-sporulating bacteria produce more of the
same cell in spite of differences in the environment
(homeostatic regulation) whereas differentiated cells
8. 2
maint~in their differences in an identical environment
(dynamic regulation; Davis et al., 19(2). One factor that
may be responsible for this difference is the presence of
histones in the chromosomes of eucaryotes. Histones are
basic proteins hypothesized to repress DNA transcription
. and thereby control differentiation by forming complexes
with the anionic phosphate groups of DNA (White et a1.,
1968).
Morphological Changes. The general steps in the formation
of a spore have been determined by Young and Fitz-James
(1959) for Bacillus cereus and by Ryter (1965) for ~acillus
The overall process appears to be the same for
both organisms (Kay and Wa!ren, 1968). Eight different
!3tages have been assigned to B. subtilis (Schaeffer et al.,
1965) beginning with the vegetative fbrm containing two or
more DNA copies at the end of exponential growth (stage 0)
(Figure l).
Stage 1 corresponds to the formation of an axial
chromatic filament. The chromosome forms a thread which
lies along the entire length of the cell; the chromosome
may be attached to the membrane by an invagination of the
plasma membrane (mesosome).
Stage II involves the development of the spore Reptum.
One-half of the DNA moves to one end of the cell and the
spore septum (a membraneous structure) begins to be formed.
The septum is attached to a mesosome.
9. FIGURE 1
The general morphological and biochemical events
associated with sporulation in aerobic Bacillus. The
entire process takes about eight hours. In different
sporulation media, the biochemical events may occur at
different times. The diagram represents the compilation
of information obtained by various investigators (Mandel-
starn, 1969).
10. .- -.-. --,----"'f"I..,...--......--- - --.--
!-~;:~,--..--.-"'----
I JJ -'-:0;' ' j • I )'
I
,!if t~',~~,~ '. Ii f ~'I.. If
i: ,'C' ~ f )7; 1II &' "4~
1'1 ,~ . I ;i.
! Hi ;~ II, I I. 1'~ 1I' In .., ': !~. I
~ll (O~
I ,1 ~.~~ , , ~~ji ~
r" STAGE ;-- ' ' STA:E I _ l STAGE II STAGE 111 ': 5T AGE IV • I
~ 11
~J
u 'i
.jJ
Spore protoplast
--1~~'"
/,.~~ ~
!~J{~~)'~·).tI '~ l
t . I
; ;
~V" i
, I
S'r ,G EV srACt VI
CO~t Maturation
I ,~ t ~ V)
I
',Vegct:ltivc ·. Chromatin 'fliament I' Spor~ septum
cell I
I < I
IAlanine 'I' Alk31inc PhO'P.hlt~uc
Cortex form:ltion
(rcfr~ctility) for Mation -1
I
Antibiotic
Exc-prctease
?roi:ein turnover
RitJorwdC'"se
Amylase
t ' i
Aconitase
H'e~t·rcsistant c:aal;uQ
dchydrogcnas~ .
Giucc:i.c dchrd";}t:~r,a::.e
! Ribosid~se . . I~ystc , n(~. I
I " Incorpcr;).tlOn !
! 'Ad(i:ldsitfcCC3.minase 1
~ , .,' . ,.. i O':~! i)ol
IDipicdlnic ;ldd " i rcsj~.i: Hi CC
'. I '
I Upt·~l.l<c of Ca2
.. I
~______~__::3~______.~
I .
i ~"
II
Alanine
r~cemlse
Heat
re~istar.ce
~, '
I
I
t
~,
11. III.
~be formation of the spore protoplast occurs in stage
The septum extends from the point where it is con-
nected to the mother cell membrane and continues toward
the end of the cell; during this process, the fore-spore
becomes unattached from the mother cell and ~loats freely
in the cytoplasm of the sporangium.
The spore cortex is formed during stage IV. Material
begins to be deposited between the double membranes of the
spore protoplast, and is probably the layer in which the
mucopeptide of the spore is 'contained.
4
Stage V is the development of the spore coat. Material
begins to be deposited farther away from the cortex.
The spore undergoes maturation in stage VI. The coat
material becomes deriser and laminated by the laying down
of sheets of material.
In stage VIr, the sporangium disintegrates by the action
of lytic enzymes and results in the release of the completed
spore.
Sporulation in B. subtilis consists of an ordered
sequence o'f structural changes coupled with an ordered
sequence of biochemical events.
correlated by Mandelstam·(1911).
The two events have been
Biochemical Change~. Approximately an hour and a half after
the end of logar1thmic phase (stage I) protein turnover
ensues and exop:(',case, antibiotic, ribonuclease and amylase
12. appear in the medium.
After one and one-half to two and one-half hours (stage
II), alanine dehydrogenase appears.
Stage III (two and one-half to four and one-half hours)
yields enzymes of the glyoxalate and citric acid cycles.
Also, a heat resistant catalase, glucose dehydrogenase and
alkaline phosphatase are formed.
Stage IV (four and one-half to six hours) results in
the production of ribosidase, adenosine deaminase, dipico-
linie acid and the uptake of Ca+ 2 . In addition, sulpholactic
acid is produced.
After six to seven hours (stage V) the incorporation
of cysteine in the spore coat begins and the spore becomes
resistant to octanel und other organic solvents.
During stage VI (seven to eight hours) alanine racemase
is produce~ and heat resistance occurs.
Importance of Some Sporulation-Specific Events. Some of
the biochemical substances listed above are not necessary
for sporogenesis, others are. There are a number of
categories in which these substances have been placed
(Freese et a1., 1969; Mandelstam, 1969; Hanson et al., 1970) s
namely: 1) byproducts~ such as the pigment melanin found
in spore-forming colonies· of B. subtilis~ 2) repressed
vegetative enzymes that become derepressed. in a poor gr(1'H'th
medium (~.&., glyoxalate and tricarboxylic acid enzyTh"
3) vegetative cell enzymes which continue to be synt .:ized
13. 6
duri~g sporulation (~.£.t enzymes needed for catabolic
reactions, amino acid and nucleotide biosynthesis, 4)
enzymes involved in the formation of spore-specific com-
ponents, but which are not incorporated themselves into
the spore (~.£., dipico1inic acid synthesis), 5) spore-
specific components, such as dipicolinic acid which is
synthesized late in sporulation and does not occur in vege-
tati ve cells.
Aconitase is an example of the second category~because
the enzyme is derepressed. under conditions of glucose ex-
haustion (Hanson and Cox, 1967; Mandelstam, 1969).
Alanine dehydrogenase (Warren, 1968) and alanine race-
mase (Stewart and Halvorson, 1954). are examples of spore
components that may be involved in germination (third
category) •
Dipicolinic acid is produced by the action of two
enzymes: dihydrodipico1inate synthetase and dipico1inic
I
acid synthetase. These enzymes are not necessary for
sporulation (Fukuda et al., 1968; Sebald, 1969) because
mutants unable to synthesize dipico1inic acid produce normal
heat resistant spores when the medium contains dipicolinic
acid (Halvorson and Swanson, 1969); when dipicolinic acid
is excluded, these mutants produce sp~res with a much lower
heat resist~nce (Halvorson and Swanson, 1969; Murrell et al.,
1969) .
Stage I of'sporulation brings about the formation and
release into the medium of antibiotics such as bacitracin
14. (from"B. licheniformis) and bacilysin (from B. subtilis).
It is not known whether or not these products are needed
for sporulation to occur. Hodgson (1970) suggested that
7
the most common characteristic of the sporulation associ-
ated antibiotics is.their ability to modify membrane perme-
ability, functioning as detergents,ion carriers or by
degrading structural components. Hodgson (1970) suggested
that the completion of stages IV and V of sporulation might
be dependent upon the action of such antibiotics via membrane
permeability modification. One of the peptide antibiotics
might be a carrier for dipicolinic acid and Ca+ 2 ions across
the outer fore-spore membrane during cortex' formation.
Basic polypeptides ·such as polymyxin B may contribute to the
removal. of water and contraction of the pre-spore. One of
the surface-active peptides might also be responsible for
the change in the outer fore-spore membrane that allows
bonding with the spore coat.
Also released during stage I are ribonuclease and
amylase. Ribonuclease may be involved in sporulation be-
cause some mutants lacking it have been found to be asporo-
genous as well (Schaeffer, 1967). Mutants have also been
found which lack amylase but continue to sporulate; thus,
this enzyme does not appear to be directly involved in
sporulation (Schaeffer, 1969).
Another extracellular product formed during stage I
is protease. A possible relationship between protease(s)
15. produ~tion and sporulation has been suggested repeatedly
(Spizizen, 1965; Mandelstam et a1., 1967; Schaeffer, 1967;
8
Mandelstam and Waites~ 1968; Schaeffer, 1969). The ability
to produce an extracellular protease seems to be linked
(genetically or fun9tiona1ly) to the ability to sporulate
(Levisohn and Aronson, 1967).
Importance of Protease to Sporulation. The evidence for
the involvement of proteases in sporulation may be summarized
as follows: 1) protease-negative mutants have been found
which are asporogenous or oligosporogenous, don't produce
any antibiotics (characteristically formed by sporulating
cells),and their competence in genetic transformation is
greatly decreased (Schaeffer, 1967); 2) the protease
inhibit?r, L-cysteine, postponed the intracellular breakdown
of protein for a few hours and depressed spore production
to 0.2 percent of the usual level (Mandelstam et al., 1967)·-- ,.
3) mutants that regained protease activity by genetic trans-
formation or transduction showed the wild type rate of
protein degradation, normal spore formation, antibiotic
production and competence (Mandelstam et al., 1967); 4)
vegetative cell protein breakdown and sporulation could be
prevented at the end of exponential growth in B. subtilis
by addition of chloramphenicol (an inhibitor of protein
synthesis); it has been postulated that this effect may be
due to the inability of the inhibited cells to make a
required protease (s) (Kornberg et al., 1968)" It must be
16. 9
pointed out that the production of protease and the ability
to sporulate may be functionally linked or may be acciden-
tal. The two properties are closely related in B. subtilis.
Role of Protease(s). Recently, several theories have been
advanced concerning. the, role of proteolytic enzymes in the
biochemical and morphological alterations that accompany
unicellular differentiation (Mande1stam and Waites, 1968;
Losick et al., 1970; Sadoff and Celikkol, 1971; Sadoff et
a1. ~ 1971; Millet et al., 1972).
During sporulation, protein and nucleic acid turnover
rates are greatly magnified (Kornberg et al., 1968; Mandelstam
and Waites, 1968; Schaeffer, 1969), allowing synthesis of
new kinds of proteins to occur from pre-existing protein~.
One theory for the role of extracellular protease(s) is that
they supply nutrients from any proteins in the environment,
while intracellular protease(s) enable the cell to change
its range of proteins and to synthesize spore structural
components from internal sources at a time when the external
environment would contribute minimally (Mandelstam and
Waites, 1968; Hodgson, 1970).
Limited proteolysis may also be an important process
in sporulation because it,brings about vast phenotypic
changes without the need for gene modification or a second
set of spore genes (Sadoff et al., 1971), i.~., a large
number of macromolecules do not have to be produced for
17. 10
de novo protein synthesis to occur ....
Another role for proteases may be the proteolytic
digestion of inhibitor(s) or repressor(s) which is directly
or indirectly involved in many biochemical pathways in the
vegetative cell. Upon exhaustion of a particular nutrient,
the amount of this repr~ssor is reduced and protease
synthesis is ·:Ie rep res sed (Schaeffer eta1 ., 19 65 ) . It is
known that glucose in the presence of nitrogen inhibits
sporulation by the mechanism of catabolite repression
(Schaef'fer et al., 1965); when glucose becomes depleted from
the medium this repression is released and proteases are
produced.
The sporulation antibiotic of B. cereus can be produced
in vitro by proteolysis of 50 S ribosomes from vegetative
cells; it is a peptide similar to bactitracin. This sug-
gests that protease may exert some kind or translational
control during the course of sporulation (Sado~f and Celikkol,
1971) .
In 1970, Sadoff et ale showed in vitro that a snoru-- ~
lati0n-specific protease from B. cereus converted vege-
tative cell fructose l,6-diphosphate aldolase to spore
e..1 do 1 a s e • These two enzymes were not identical, but
structurally related. The molecular weight of the vegeta-
tive aldolase was 79,000 daltons while that of the spore
aldolase was 44~ooo. They also had dif~erent migration
patterns in acrylamide gel electrophoresls, heat resistance
in the pre3~nce of Ca+2~ ion requirements for catalytic
18. 11
acti~ity and activation energies. Thirty-five percent of
the aldolase activity was destroyed by the protease. The
physical and chemical data presented suggests that the
protease produced a uniform population of enzymes, instead
of randomly acting to produce an average reduction in
activity.
Differences also exist between the vegetative and
spore forms of purine nucleoside phosphorylase of B. cereus,
although they are products Df the same cistron (Engelbrecht
and Sadoff, 1969). It is not known whether limited prote-
olysis occurs with this enzyme.
Escherichia coli cells infected by the bacteriophage
T4 serve as a useful model for studying the regulatory
mechanisms governing sequential gene expression at the
transcriptional level (Bautz et ale ~ 1969; Goff and Webber,
1970; Schachner et a1., 1971; Szulmaj ster, 1973). Three
genetic regions have been discovered on T4 DNA. These g(~nes
code for three different messenger RNA's; they were named
in the order of their expreRsion: immediate-early,
delayed-early and late& The in vitro transcription of
the immediate-early gene was controlled by E. coli sigma
factor (Bau~z et al., 1969). Further m-RNA transcription
was associated with two types of changes in the host RNA
polymerase; namely, alteration and modification (Schachner
~.al., 1971). Alteration involves the sigma factor itseJ.f
or a decrease in affinity for a changed core polymerase.
19. 12
·of
Modi~ication requires protein synthesis and brings about
structural.changes in all the subunits. The a-subunit is
modified by the covalent addition of 5'-adenylate (Goff
and Weber. 1970) and the S:and S~ subunits are also changed
(Travers, 1970). The host a factor is replaced by a new
phage-induced sigma-like factor~ This new a factor is not
detected until 5-15 minutes after infection, then it dis-
appears. These modifications may playa role in shutting
off transcription of the host genes while expressing new
phage genes; this new a factor insures accurate and efficient
initiation of RNA synthesis at a specific promotor site of
the T4 DNA template, in additibn, the E. coli ribosomes
preferentially recognize the T4 m~R~Ats start signals
1Summers and Siegel, 1969; Travers, 1969; Szulmajster, 1973).
By' an analogous mechanism in. B. subtilis, vegetative
RNA synthesis is turned off and new messenger RNA is pro-
duced during sporulation (Doi and Igarashi, 1964; Aronson,
1965). This alteration in gene expression again could be
accounted for by a change in the template specificity of
the RNA polymerase. Such a change is known to take place
early in sporulation in B. subtilis RNA polymerase. Accom-
panying this alteration, is a complete loss of ability to
transcribe virulent phage ~e DNA in sporulating cells
(Losick and Sonenshein~ 1969).
In sporogenesis, the loss of vegetative template
specificity was thought originally to be accompanied by a
20. 13
change in the subunit structure of RNA polymerase (Losick,
1972) • B. subtilis RNA polym~rase can be separated into
two major components: a core enzyme containing S~ and S
subunits o~ 155,000 da1tons, two a-subunits of 42,000
daltons, apd a a-subunit of 55,000 daltons (Losick et al.,
1910). When RNA polymerase from B. subtilis is extracted
from the sporulation phase, the S~-polypeptide is replaced
by a smaller polypeptide of 110,000 daltons. It was
hypothesized by Losick .~1 ale (1970) that the sporulation
RNA polymerase was derived from the vegetative a~-polypeptide
by specific proteolytic cleavage.
Millet et ale ( 1972) pro v ide d e v ide n c e ,f'0 r protea s e s
being responsible, in vivo, for 'the conversion of' B. subtilis
vegetative RNA polymerase to spore form. Purified RNA
polymerase from B. subtilis vegetative cells added to pure
intracellular protease from B. megaterium gave the same
structural modification in one of' the a-subunits, in vitro,
as seen in the RNA polymerase isolated from spores.
Recently~ Linn et ale (1973) have provided evidence
to dispute the aforementioned findings. First, the a1tera-
tion o'f the 8~subunit may be due to proteolysis during
purification and therefore artifactual. Second, the
alteration of the core enzyme does not occur until the
s~aond hour of sporulation~ while the change in templAte
specificity occurs in the first hour after logarithmic
growth. So, the change in core enzyme occurs too late to
21. 14
,of
account ~or the loss of sigma activity. These authors con-
clude that because the alteration of core RNA polymerase
does not account for the loss of sigma activity, then sigma
factor may be destroyed, inactivated or merely removed
from the RNA polymerase in the early sporulation events.
An additional possibility for proteasffi is that they
have no essential function in sporulatio'n and their appear-
ance is merely due to an ordered sequen6e of biochemical
events. The mutants lacking protease(s) therefore would
have been impaired in such a way that the biochemical events
needed for sporulation could not occur and consequently no
protease was produced (Mandelstam, 1969).
Particular Proteases. Three classes· of extracellular pro-·
teases have been isolated and characterized in B. ?ubtilis;
they are classified according to their respective iso-
electric points.
The alkaline protease exhibits high proteolytic
activity but low esterolytic activity has a molecular
weight of 28,000 - 30,700 daltons and a pI> 7.0 (Guntelberg
and Ottesen, 1954; Matsubara et a1., 1958; Rappaport et al.,
1965 ) .
The neutral protease contains zinc as a prosthetic
group~ readily hydrolyzes casein and aminopeptides, but not
esters, has a molecular weight of 33,800 - 44,700 daltons
and a pI ~ 7 (McConn et al., 1964; Keay, i969).
The acid protease has weak proteolytic activity and
22. 15
strOl'lg esterolytic activity (Boyer and Carlton, 1968;
Mande1stam and Waites, 1968; Millet, 1970; Prestidge et al.,
1971) has a molecular weight of 35,000 - 40,000 daltons
and a pI < 7.0 (Boyer and Carlton, 1968).
Very little unfterstanding has been achieved concern-
ing the synthesis of exoenzymes and the processes that lead
to the elaboration of these enzymes into the surrounding
medium (Both et ~l., 1972).
The production of extracellular protease, a-amylase
and ribonuclease in B. amyloliquefaciens are sepa}·ateJ.y
controlled. Ribonuclease synthesis is repressed by
inorganic phosphate, a-amylase is repressed by any medium
which stimulates cellular growth, and protease synthe~is
is rep~eEsed by amino acids; no single acid is effective
by itself, but a mixture of either proline and isoleucine
or o~ glutamic acid, aspartic acid, glutamine and aspara-
gine yields greatest repression of protease (May and
Elliott, 1968). Furthermore, the time courses for enzyme
appearance are different. Ribonuclease is produced
linearly while a-amylase and protease production are
bipbasic (May and Elliott, 196B).
May and Elliott (l96B) hypothesized that protease
is secreted from the B. subtilis cell' as it is synthesized
on ribosomes bound at translational-extrusion sites on
the cell membrane (Both et al., 1972), i.~. there is no
significant intracellular accumulation; this fInding was
23. 16
in agreement with a-amylase and ribonuclease secretion.
Therefore~ these authors speculated that none of these
enzymes are ever present in the completed form inside the
cell membrane, but rather that the nascent polypeptide
is extruded through the membrane as it is synthe-
sized to take up its tertiary structure with enzyme
activity on the outside. The exact location for the pro-
duction of active protease is unknown. Several models
for extrusion of the polypeptide have been proposed
(Fig. 2). The possibilities are as follows: a) the
enzyme may exist as an intermediate form in the cell mem-
brane and assumes its final configuration in the peri-
plasmic spaces b) the enzyme is folded in the cell
membrane, c) the enzyme is active in the periplasmic space,
d) the enzyme is folded only upon reaching the extracellular
medium.
Esterases.
B. subtilis.
Two intracellular esterases have been found in
The first esterase, designated "Aft , has been
identi~ied in the cytoplasm of vegetative and sporulating
cells (Bott, 1971; Higerd and Spizizen, 1973): Extracts
from mature spores, as well as from early blocked
asporogenous mutants demonstrated esterase A activity_
Esterase A had been purified to isoelectric homogeneity and
the molecular weight of the native enzyme had been de-
termined by gel filtration chromatography to be 160,000
daltons. Sodium dodecyl sulfate (SDS) gel electrophoresis
24. FIGURE 2
Four possible locations are presented where the
emerging polypeptide can assume its three-dimensional
form. A) The enzyme exists as an, intermediate in the
cell membrane which assumes its final configuration in
the periplasmic space. B) The enzyme is in its final
configuration in the cell membrane. C) An active enzyme
is produced in the periplasmic space. D) The enzyme is
functional only upon contact with the extracellular
medium (Both et al., 1972).
26. suggested subunits of approximately 31,000 daltons! The
isoelec~ric point was determined-at 6.4 (Higerd and
Spizizen, 1973).
The second intracellular esterase, designa.ted "B n
,
vas not detected duri~g vegetative growth but appeared
after logarithmic growth. ceased (Bott, 1971; Higerd and
Spizizen, 1973). This esterase was not formed by early
blocked asporogenous mutants, but could be detected in
mature spores produced by wild type stains.
An assessment of the molecula..:' weight of a
partially purified prepa~ation of this enzyme resulted in
a determination of 51,000 da1tons. The isoelectric point
was 5.4 (Higerd and Spizizen, 1973).
18
The two intracellular esterases and the extrac~llular
(acid) protease of B. subtilis possess several properties
in ccmm.on (Hall ~ al., 1966; Boyer and Carlton, 1968;
Michel and Millet, 1970; Prestidge et al., 1971; Higerd ane
Spizizen, 1913). All three ex~ibit high estero1ytic
activity and low proteolytic activity. All three enzymes
are capable of hydrolyzing B-naphthyl acetate, are inhibited
by diisopropyltluorophosphate (DFP), possess similar 1so-
electric points and can be partially purified through the
use of similar laboratory techniques.
The time of appearance of acid protease coincides
with the appearance of esterase B (Michel and Millet, 1970;
27. 19
Prestidge et al., 1971; Higerd and Spizizen, 1973). The
enzyme first appears in the growth medium a~ter logarithmic
growth ceases. Early blocked asporogenous mutants fail
to elaborate this enzyme as well as other extracellular
enzymes (Spizizen, .1965; Ionesco et al., 1970).
The intracellular esterase B and the extracellular
(acid) protease are regulated by mechanisms that control
sporulation (Michel and Millet, 1970; Higerd and Spi~izen~
1973) and :~atabolite repression (Levisohn and Aronson,
1967; May and Elliott, 1968; Both et a1., 1971; Sadoff and
Celikkol) 1971); it would appear that~both enzymes are under
similar physiological nnd/or genetic .control. Esterase A,
on the ether hand, does not appear to be controlled by the
same mechanism (Higerd andSpizizen, 1973). One hypothesis
that deserves investigation is that the high molecular
weight esterase gives rise to the lower molecular weight
esterase (by limited proteolysis or dissociation) and that
the regulation, in part, is based 'on the conversion of the
high molecular weight form to the lowt·molecular weight form.
~wo questions which naturally arise are:. 1) Have
esterases been found in mammalian cells?
do they compare structurally and chemically to the bacterial
esterases?
Vertebrate tissues show four types of esterase activity,
viz carboxylesterases (EC 3.1.1.1), ary1esterases (EC 3.1.1.2),
acetylesterases (EC 3.1.1.6) and cholinesterases (EC 3.1.1.8)
28. 20
(Holmes and Masters s 1968).
Non-specific esterases (i.~., those which hydrolyze
a variety of small esters) have been found in several
organs of female Aedes aegYpti (Briegel and Freyvogel,
1973), mouse spermatozoa (Bryan and Unnithan, 19(3), guinea
pig liver and kidney (Chow and Ecobichon, 1973), human
gingiva (Cohen, 1967), human urine (Therrien et al., 1971),
human prostate glands (Atanasov, 1973) and human milk
(Kobayashi et al., 1973). They have been associated with
the following functions: active transport (Moule, 1964),
detoxification of endotoxin (Skarnes, 1970), lipid metabolism
(Fruton and Simmonds, 1963), protein synthesis (Bernsohn
et al., 1966) and.proteolysis (Hopsu and Glenner, 1964).
N9n-specific carboxy1esterases are found in animals,
plants and bacteria. They are interesting to compare to
esterase A, because they too are sensitive to organophos-
phate inhibitors (Arndt et al., 1973) and their mechanism
of action involves an acyl enzyme intermediate (as is known
to be the case for serine hydro1ases)(Arndt and Krisch,
1973).
Unfortunately, the vertebrate esterases usually exist
in multiple forms, as charge isomers or as readily formed
aggregates (~'K" 12 electrophoretically different activity
bands w~re found for guinea-pig liver '~nd 14 bands for
guinea pig kidney (Chow and Ecobichon, 1973)); this mak~s
their purification and characterization difficult. The
29. 21
function of mammalian esterases in cellular differentiation
remains undetermined.
Several vertebrate carboxylesterases exist as trimeric
molecules with subunit weight of 61,500, viz pig liver and
kidney esterase (Heymann et al., 1971), bovine liver esterase
(Wynne and Shalitin, 1973), rat kidney esterase (Kleine and
Brebeck, 1972) and rat liver esterase (Arndt et al., 1973).
Crude liver esterases of pig, horse, rat, cow, pidgeon and
rabbit have been shown to exhibit substrate activation with
different derivatives of m-hydroxybenzoic acid as substrate
(Hofstee, 1972). This suggests that the active center of each
enzyme is the same, while the additional binding site (postu-
lated to be involved in substrate activation) is species-
specif~c (Hofstee, 1972).
Reason for Research. The sporulation process and related
biochemical events have been described at length, primarily
to place intracellular esterases in their historical per-
spective and to show the impetus for this project.
While the main thrust of research in bacterial
sporulation has dealt with the target for ttproteases",
few investigations have centered around the individual
enzymes that may be responsible for such alterations. This
area has been neglected primarily because of the lack of
information concerning the properties of these catalysts.
The purpose of this investigation was to purify and
characterize esterase A from B. subtilis, so that some
insight may be gained into its possible relationship to:
30. 22
1) the intracellular esterase B, 2) the extracellular (acid)
protease, and 3) the sporulation process.
Recently, several mutants of B. subtilis have been
isolated which lack either esterase A or B activity (Higerd,
in pr~paration). One mutant, namely~ EA 1, produces
esterase B activity solely and is able to sporulate at wild
type fre~uencies. These results suggest that esterase A is
not a precursor of esterase B, and that esterase A activity
is not required for sporulation.
This work, therefore, has provided physical and chemi-
cal data on the carboxylesterase A of B. subtilis.
31. MATERIALS AND METHODS
Organism. Bacillus subtilis strain" SR22 (SpoA12;
trpC2). A 350 liter culture grown in a 500 liter fermentor
(Department of Biochemistry, University of Georgia, Athens,
Georgia) provided 1,800 grams of cells for the purifica-
tion.
Chemicals. The following reagents were used in this
study: glucose, calcium nitrate, manganese chloride,
ferric sulfate, copper sulfate, sucrose, ethyl acetate,
hydrochloric acid, trichloroacetic acid, mercuric chlQride,
acetone (Mallinckrodt Chemical Works, St. Louis, Missouri);
nutriept broth> tryptose blood agar base (Difco Laboratories,
Detroit, Michigan); sodium chloride, potassium chloride,
magnesium sulfate, pot~ssium hydroxide, B-naphthyl acetate
(recrystallized from dilute ethanol), sodium carbonate",
sodium potassium tartrate, Folin Ciocalteu reagent (2 X),
N,N,N' ,N'-tetramethylethylen~diamine, ammonium persul~ate;
bromophenol blue, mono and dibasic potassium phosphate,
.
glacial acetic acid, ethyl alcohol, sodium fluoride (Fisher
Scientific Co., Fairlawn, New Jersey); tris(hydroxymethyl)
aminomethane (Tris), Fast BB Salt, Coomassie' Brillia~t
Blue R, cetyl bromide, cetylpyridinium chloride, cetY)_t"rl_..-
methyl ammonium bromide, eserine, a,a'-dipyridyl, 1,10-
phenanthroline monohydrate, p-nitrophenyl propionate,
32. 24
p-nitrophenyl butyrate~ p-nitrophenyl caprylate, p-nitro-
phenyl caprate, L-tyrosine ethyl ester hydrochloride,
L-tosylamide-2-phenyl-ethylchloromethyl ketone (TPCK)
(Sig~a Chemical Company, St. Louis, Missouri); p-nitro-
phenyl acetate, p-tosyl-L-arginine methyl ester hydrochloride
(TAME), N-benzoyl-L-tyrosine ethyl ester (BTEE), N-acetyl-L-
tyrosine-methyl ester, N-acetyl-L-phenylalanine methyl ester,
N-benzoyl-L-arginine ethyl ester hydrochloride (BAEE)
(Schwarz/Mann, Orangeburg, New York); crystallized bovine
serum albumin (Miles Laboratories, Kankakee, Illinois);
acrylamide, N~N'-methylenebisacrylamidet a-naphthyl acetate,
.i-butyl acetate, i-propyl acetate, t-butyl ,acetate, n-propyl
acetate, pentyl acetate, ethyl chloroacetate, ethyl bromo-
acetate, ethyl butyrate, ethyl iodoacetate, ethyl propionate,
n-butyl butyrate, Photo-Flo-20D (Eastman Kodak Company,
Rochester, New York); Cellex PAB, Cellex CM, Cellex D,
Cellex T, Bio Gel P-l50 (100-200 mesh), Bio Gel RTP (Bio Rad
Laboratories, Richmond, California).
t
E._~E..nration of Extracts. The culture medium used was
2 X sporulation medium (Greenleaf and Losick), containing
(per'lite.c): glucose, 0.5%; nutrient broth, 16 g; potassium
chloride, 2 g; magnesium sulfate ·7H20, 0.5 g; calcium
nitrate, 2 X IO-3M; manganese chloride, 2 X lO-5M; ferric
If" t 2 X lO-6M.Rua e, The medium was adjusted to pH 7.0 with
10% potassium hydroxide.
Three h1.:ndred fifty liters of sporu~ atio,n medium vas
33. seeded with ten liters of an 18 hour culture of SR22.
Growth of an aliquot was followed turbidimetrically at
6(0 nm in a Spectronic 20 colorimeter (Bausch and Lomb)
eluipped with a red filter. The generation time was
approximately thirty minutes.
25
Cells were refrigerated (4c) at the end of logarithmic
grcwth Lnt: harested by centl'ifugation.
1,800 grams of wet cells were obtained.
Approximately
Following centri-
fugation, the cells were resuspended in 0.05 M Tris-Hel
buf~er, pH 1.5 containing 0.04 M magnesium sulfate, washed
twice, then centrifuged at 21,000 X g for 15 minutes at
4c. The cells were then resuspended in 0.02 M potassiUM
phosphate buff~r, pH 7.5. and centrifuged. Finally, the cells
were resuspended in the latter buffer at a final concentra-
tion of 0.2 g (wet weight)/ml and frozen at -20C ..
Prior to sonication, the frozen sample was thawed ut
room temperature and placed in a rosette flask (Branson
Sonic Power; Danbury, Connecticut) surrounded by an ice-brine
bath. The sample was son1cally disrupted (Bronson Model L)
for fourteen minutes. The sonication time of fourteen minutes
provided approximately 93 percent recovery of enzymatic
activi ""GY. The broken cell suspen3ion was freed from cellular
deb r i sLy c ent r i fu gat ion at 27, 000 X g for 30. min ute >: at 11 C ,
Esterase and. Prot~~~.says" Esterolytic acti. ity W.'lS·
as fj aye L"i by 0 '1 e 0 'f t h r e e mE; the d s . The f 1. r stutil i zed 6- nap h t hy1
acett.te as substrate and "'as performed by a modifj.ca:~ion of
the method of Seligman and Na~hlas (1950). To 1.0 ml of
test solution appropriately. diluted in 0.02 M pot~ssium
34. phosphate buffer~ pH 7.5, preincubated at 30C for five
minutes, 5.0 ml of the prewarmed substrate (0.04 mg!ml)
2G
in the same buffer was added and the reaction mixtll.re
incubated at 30C. After 20 minutes, 1.0 ml of a freshly
prepared solution Qf Fast BB Salt (4 mg/ml) was added,
followed two minutes later by 1.0 ml of 40% trichloroacetic
acid to stop the reaction and precipitate the proteins.
The resulting pigment was extracted into the non-polar
phase by vigorously shaking the reaction mixture with 10 ml
cf ethyl acetate. After settling for several minutes, 10
ml of the top .layer was removed and centrifuged for ten
minutes at 3~000 X g. A portion of the organic layer was
removed and its absorh~ncy measured in a Spectronic 20
spectrophotometer at 540 nm. From a calibration curve of
pure S-naphthol, color density was converted to micrograms
of a-naphthol. The unit of e~terase ~ctivity waF defined
as the number of micrograms of B-naphthol liberated at 3UC
per ml 'of preparation in one minute:.
The second esterase assay utilized p-nitrophenyl
acetate as substrate and employed a modification of the
method of Huggins and Lapides (1947). To 3.0 ml of 0.02 M
potassium phosphate buffer, pH 7.5 in a 3.5 ml quartz
cuvette (Markson Science ~nc.,Del Mar, California) was
added 0.1 ml of 3 X lO-3M p-nitrophenyl acetate in aceto-
nitrile. Fifty ~l of an appropriately diluted enzyme
preparation was added to the sample cuvette and mixed
thoroughly. Hydrolysis relative to the'reference cell was
35. 2'{
followed by the change in the absorbance at 400 nm per unit
time F.. t room temperature on a recording Perkin Elmer double
beam spectrophotometer (Coleman Model 124). From a cali-
bration curve of pure p-nitrophenol, the change in the
absorbance was converted to micromoles of p-nitT~phenol
liberated. The unit of esterase activity was defined as
the micro'lloles of substrate hydrolyzed per minute per ml of
enzyme preparation at room temperature. The derivative~ of
p-nitrophenol were assayed in a similar manner.
The third assay for esterolytic activity on a variety
of substrates measured the amount of acid liberated during
the reaction. The reaction mixture (2.5 ml) contained 0.25 M
of substrate in 0.01 M potassium phosphate buffer, pH 7.55
with 0.1 M ethanol; the r~action vessel was equilibrated to
37C by a Lauda Model K-2/R constant temperature water bath
(Brinkman Instruments, Inc., Westburg, New York). After a
stable baseline was established on a REC 60 Servograph
(Radiometer; Copenhagan, Denmark), 15 ug of an esterase A
preparation was added and hydrolysis was followed for a
minimum of eight minutes. An autotitrator TTT 60 with an
automatic burette (ABU 12; Radiometer) was employed. The
acid liberated upon hydrolys.is was continuously back-titrst0d
with 0.04 M NaJH to pH 7.55, as measured ~y ~ PHM62 3t~ ndard
pH meter (Radiometer). The unit of esterase activity was
defined as the number of millimoles of substrate hy1ro-
lyzed at 37C per minute per ml of preparation.
36. 28
The Lowry (1951) r.ethod :for protein determination was
employed. Bovine serum albumin was used to prepare a
standard curve.
Fractions from columns were monitored :for their
absorbance at 280 ~m, either in a double beam spectrop:lo-
tometer (Cole n Model 124) or by an absorbance monitor
(M)del UA-4; _,;ls1.rumentation Specialties Company, ~nc.,
Lincoln, Nebraska).
Electrophoresis. A disc electrophoresis apparatus
similar to that described by Davis (1964) was used (ISCO
Model 311) connected to a power 'SOlree (ISCO Model 390).
The glass tubes had a5 mm internal diameter and were
76 mm long. The height of the acrylamide gel column was
72 rom. The 7.5% acrylamide gel .system was prepared as .
f 0 11 0 W s . Stoe k sol,.!t ion ( A) con t a i ned: .1 N He1, 48 ml;
Tris, 6.85 gm; and N,N,N',N~tetra~ethylethy1enediamine,
o.4G nl, b r 0 -d. g h t tor; vol u!n e 0 flO a ml wi "1,:, h wa t e 1", pH =
7,5. Stock (B) clntained: ~cryla~ide, ~0 gm~ ;N)N'-
methylenebisacrylamide,
"distilled water. Stock
0.8 gros, brought to 100 ml with
(C) contained: 100 ml distilled
water. Stock (n) contained: ammonium persulfate, 0.07
gms, brought to 100 ml wjth distilled water. The ·working
solution was prepared by mixing stock A, B, C, and Dwell
in the following proportional volumes: 1/2/1/4 ref ;)ect .. veJ .•
Two ml o~ the working solution was placed in each glass tube
end ater was carefully overlaid.
37. 29
acrylamide was complete by 30 minutes at room t'emperature.
Both upper and lower reservoirs of the electrophoresis
apparatus were filled with 0.1 M Tris-Hel buffer, pH 7.5.
The lower electrode served as the anode.
After polymer~zation of the acrylamide, the tubes
were pllced at 4c and 50 ~l of the test solution containing
25 pI 0 30% sucrose and a snaIl amount of bromophenol blue
were carefully layered above the gel. After a loading
amperage of 2 roa/tube for'5 minutes, a constant current of
5 ma/tube was maintained uniJil the tracking dye reached
the end of the tube (approximately 3 hours).
After electrophoresis, the gel was carefully removed
from the electrop~oresis tube where it was stained either
for esterase activity or protein,. The esterase staining
solution consisted of 1 ml of S-naphthyl acetate (20 mg/ml)
in acetone, 50 mg of Fast Blue BB and 99 ml of 0.1 Tris-HCl
buffer, pB 7.5. After 30 minutes of agitation, the gels
were rinsed with water and stored in 7% acetic acid. The
protein fixing solution was 12.5% TeA (W/V) and was
applied for 30 minutes. The protein staining solution
consisted of 0.02% (W/V) Coomassie Brilliant Blue (R 250),
20% methanol (V/V) and 7% acetic acid (V/V). The gels were
stained for protein overnight and the background lestained
in a gel electrophoresis di~fusion destainer (Bio Rad
Model 170) against 7% acetic acid. The gels were stored
in 7% acetic acid.
38. 30
Conductivity Measurements. A standard curve for con-
ductance in the range 0-0.5 molar potassium phosphate, pH
7.5 was made with a conductivity cell (Model 3403, Yellow
Springs Instrument Company, Yellow Springs, Ohio) connected
to a conductivity bridge (YS1 Model 31). Conductivity for
an unknown sample was converted to the corresponding molar
concentration of potassium phosphate.
Inhibitors. A variety of group specific and non-
specific inhibitors were dissolved in 0.5 ml of 0.02 M
potassium phosphate, pH 7.55, distilled water, acetonitile
or anhydrous methanol at a concentration permitting solu-
bilization in the reaction mixture. Each inhibitor solution
had a pH ::; 7.»). None of the four solvents showed inhibi-
tien of'the enzyme in control determinations of activity.
In fact, the anhydrous methanol control sho~ed 118% re-
covery, so values obtained with inhibitors dissolved in this
solvent were adjusted accordingly. Twenty-five microliters
or each inhibitor and 15 ~g of e~terase A in 200-~1 of 0.02 M
potassium phosphate, pH 7.55, were incubated at 37C for 10
minutes, then assayed by measuring the release of p-nitro-
phenol at 400 nm. If inhibition was obtained at a particu-
lar concentration, then ten fold dilutions were made of the
inhibitor until approximately 100% recovery of erlzymatic
activity was obtained.
The effect of 0-1.11 M NaCl and KC1, with no prio!
incubation, on estel'ase A activity -was determined by the
pHs tat met hod wit h e thy 1 :~" c etatea s the sub s t !' ~--. teat pH
39. 31
7 · 55. To 1.9 ml of 0.28 M, 0.56 M, or 1.11 M NaCl or KCl
in 0.01 M potassium phosphate buffer, pH 7.55, was added
67 ruM ethyl acetate. After the baseline was established,
18 ug of esterase A was added to the reaction vessel. The
control sample contained the same buffer without NaCl or
KC1.
pH Stability. ,The folldwing bufrers were prepared:
o.05 M ,p 0 t ass i urn ph 0 s ph at e, pH 6. 5- 8 . 5; o. 0 5 Mbar bit 0 1-
Hel, pH 7.0-9.0; O~05 M citr~te, pH 3.0-6.0; and 0.05 M
carbonate-bicarbonate, pH 8.5~11.O. 180 ul of each buffer
and 12 ug of enzyme were incubated in small culture tubes
(7 X 75 mm) for 16 hours at 4c, then assayed by the mixture's
ability to liberate p-nitrophenol ,from p-nitrophenyl ace-
tate at pH 7.5.
pH-, Opt imum. Potassium phosphate,. barbital-Hel and
citrate buffers, as described above, were used also for
the pH optimum. TD 1.9 ml of each buffer was added 67'
roM ethyl acetate and 18 ug of the esterase preparation.
The velocity was measured by the pH stat method~
..
Km and Vmax.' The substrate.:sused to determirle the
kinetic constants Km and Vmax were assayed at 37C, pH 7.55
by the pH stat; they were dissolved in 0.5 ml acetonitrile
or buffer at a concentration that did not cause precipita-
tion in the reaction mixture. The results'were subjected
to a least squares statistical analysis, from which Km
and Vmax were determined.
40. RESULTS
Enzyme Purification. The growth of B. subtilis SR22
was followed turbidime:tr ically (Figure 3) and the cells
were harvested shortly after logarithmic phase ended.
The cells were washed and disrupted by sonication.
P1.rification of esterase A from cell-free extracts of
B. subtilis was performed at 4c unless otherwise stated.
Manganese chloride (1M) was added slowly to the crude
sonicate to give a final concentration of 3.3% (V/V).
Sodium hydroxide was added until pH 7.5 was obtained. After
30 minutes, the mixture was centrifuged at 27,000 g for 15
minutes. The resulting supernatant was subjected to the
slow addition of acetone to a final concentration of 40%
(V!V) and left at -l5C overnight. The superHatant was
suctioned off and the settled material centrifuged at
27)000 X g for 15 minutes at -10 C. The precipitate W8S
dissolved in 0.5 M potassium phosphate buffer, pH 7.5.
After heating the enzyme solution at 70 C for 10 minutes,
the sample was immediately cooled to 4c.
Figure 4 shows the effect of heating esterase As
during this stage of purity, in the presence of 0.5 M and
1 • 0 M pot ass i um ph 0 3 Phtt e buffer, p ~ I 7. 5 .
41. FIGURE 3
Growth cutve of B. subtilis SR22 on 2 X sporulation
medium (Gr~en~eaf gnd Losick) supplemented with O.~%
glucose. The time for one gel;.eration was about 30 minutes.
The cells were harvested 4 and 1/4 hours after the inocu-
lation of the culture (arrow). (e-e) = abs6rbance at 660 nm •
.1'·"
43. F, Tf""URE 4.!, J'
The effect of ,heat. on esterase A activity. The
percent act~vity remaining after heating at the indicated
temperature for 5 min .. The prepcration was an aliquot
from step ,h of the purification (Tables 1 and 2). (~---.)
= 0.5 M potassium phosphate, buffer pH 7.5. ( o-r)) :: 1. 'J M
p(, tassium ~phosphate, buffer pH 7.5.
45. 35
The heated preparation was centrifuged at 27,000 X g
for 15 minutes and the supernatant dialyzed against three
changes of 1.2 liters of 0.02 M potassium phosphate buffer,
pH 7.5.
The dialyzed preparation was chromatographed on a
diethylaminoethyJ. (DEAE) - cellulose column which previously
had been equilibrated with 0.02 M potassium phosphate buffer~
pH 7.5. After the es~erase sample was applied to the column~
2.5 liters of the same buffer was washed through the column.
A 3.0 liter gradient of potassium phosphate buffer (0.02-
0.50 M) was applied in a linear manner. Figure 5 depicts
the elution profile obtained. The fractions containing
esterase activity were pooled and mixed with an equal volume
of 2.0 M potassium phosphate buffer, pH 7.5 and heated at
Boc for 10 minutes. The second heating step was initiated
to precipitate additional protein at an elevated temperature
that was not permissible with cruder preparations. Immedi-
ately afterwards, the prep· was cooled in an ice bath and
centrifuged at 27,000 X g for 15 minutes. The supernatant
was dialyzed against two changes of eight liters of cold
distilled water.
The solution w s concentrated by lyophilization. The
lyophilized powdEr was redissolved in 15 ml of distilled
water and 1.0 ml aliquots were applied to a Bia Gel P-150
column (1.5 cm X 90 cm) vhich had been equilibrated ~ith
0.02 M potassium phosphate buffer, pH 7.5. li'igure 6 shows
the elutiori pro~ile.
"lf~ rep 0 0"led •
Fractions containing esterase activity
46. FIGURE 5
Chromatography cf esterase A on a DEAE-cellu1ose
column (5.0 em X 3~.5 cm). The flow rate was 15 ml/rnin.
The enzyme activity was eluted from the column at about
-~_9__M potassium phosphate. (0 - 0) = absorbance at
280 nrn; (X - X) = esterolytic activity; (tJ. -/) ) = 0.02-
0,50 M potassium phosphate' buffer, pH r[ _ 5, gradient.
48. FIGURE 6
Chromatography of esterase A on a Bia Rad P-150
column (1.5 cm X 90 em), equil~brated with 0.02 M potassium
phosphate buffer, pH 7.5. A l~O m1 aliquot of the enzyme
was applied. 1.25 mlfractions were tollected. The void
volume of the column was approximately 45 mI. (0 - 0) =
absorbance at 280 nm; (X - X) =esterolytic activity.
49. CD•
o
• •
ZOI X ~'IAI'OV V 81DJ8,13 10 I'lun
o 0 0 0 0 0
10 • ", N - i
CD ~
• •
o 0
N
d
-0-0- (wu08~) eouDqJOlqy
37
50. 38
The entire procedure is summarized in the flow diagram (Figure
7) as well as in Tables 1 and 2. a-naphthyl acetate and
p-nitrophenyl acetate were used as substrates to monitor the
purification of esterase A (Tables 1 and 2, respectively).
There was a 406 fold purification with a 59% yield for BNA vs.
322 fold and 47% yield with PNPA.
Electrophoresis. ,The afo~mentioned purification
scheme resulted in five protein bands' and one activity 'band
on electrophoresis with 7.5% acrylamide gels (Figure 8).
Other SteEs Attempted. Ammonium sulfate precipitation
was the first treatment attempted to purify and concentrate
the enzyme. Fifty five percent saturation was the lowet
limit, i.~. 92% of the enzyme remained in the supernatant;
while 15% saturation was the upper limit, i~~. ~ 100% of
the enzyme precipitated. However, on several occasions,
there was a large loss (~p to 56%) in the percent yield of
the enzyme on going from 55-75% amm6nium sulfate; conse-
quently, precipitation via organic solvents (~.&. ethanol
and acetone) was attempted. Thirty percent ethanol ,resulted
in a 77% recovery; whereas 30% acetone provided a 94% yield;
therefore, acetone was utilized.
Initially, a linear sodium chloride gr$dient (0-0.5 M)'
was used to elute esterase A from a DEAE-cellulose column •.
Sodium chloride was shown to partially inhibit the enzyme
at the concentration used for elution; consequently, a
gradient or 0.02-0.50 M potassium phosphate proved satis-
factory.
52. Supernatant
Cell-free sonicate
MnC12 precipitation
Centrifuged
40% acetone precipitate
Centrifuged
Supernatant Precipitate
I
Discarded Resuspended in buffer (0.5 M)
Heated at 70C for 10 min.
Centrifuged
Superna~t~a~n~t________-+_________________________P_r..ecipitate
Di!cardedDialyzed
DEAE-cellulose
Chromatography
39
PreC_!Eitate
Di!carded
fo activity Activity
IHscarded
Precipi tate
Di sic arde d
Addition of buffer (1.0 M)
Heated at Boc for 10 min.
Centrifuged
Supernatant
Dialyzed
Lyophilized
Bio-Gel
P-l50 Chroma-
tography
No act"i vi ty_
D
O].•.s-Ic-a...r-d-e-d----.......--At· · t
C 1V1 Y
pooled
Lyophilized
" ' "Pure" prep.
53. - - -
TABLE l
Purification of esterase A with 6-naphthy1 acetate as
VOLUME
Esterase A Protein Specific
Step Before After Units (mg/ml) Activity
(rnl) (ml) (activity/ (u/mg)b
m1!min)
I . Sonicate 0 7,726 8.7 21.3 0.4
II. MnClc> 7,726 ' 8,124 7.1 14.8 0.5
III. Acetone 8,124 1,277 54.4 47.0 1.2
IV. 70C Heat
Step 1,289 1,151 49.6 9.4 5.3
V. DEAE-
Chromatog. 1,604 266 204.2 9.1 22.4
VI. 80c Heat
Step 532 518 97.0 1.4 70.8
VII. Lyophili-
zation 648 23 1,729.6 23.4 13.9
VIII. P-150-
Chromatog. 23 398 99.9 0.6 166.5
a
All values were the average of three determinations
b
u = units of esterase activity
substrate
Purifi-
cation
(fold)
1
1
3
.13
55
173
180
406
a
Yield
( %)
100
86
104
85
81
75
59
59
.f.--
o
54. TABLE 2
Purifjcation of esterase A with p-nitrophenyl acetate as substrate a
VOLUME Esterase A Specific Furifi- Y eld
Units Protein Activity cation %)
Step Before After (activity/ (mg/m1) (u/mg)b (fold)
(m1 ) (ml) ml/min)
I . Sonicate 0 7,726 251.5 21.3 11.8 1 100
II. MnC1
2 7~726 8,124 0 14.8 0 0 0
III. Ac e,tone 8,124 1,277 1,575.8 47.0 33.5 3 104
IV. 70C Heat
Step 1,289 1,151 1,470.2 9.4 156.1 13 87
v. DEAE-
Chromatog. 1,604 266 5,481.0 9.1 602.3 51 75
VI. 80c Heat
Step 532 518 2,409.9 1.4 1,759.1 149 61+
VII. Lyophili-
zation 6}~ 8 23 )~ 5 ,600 . a 23.4 1,947.1 165 54
VIII. P-150-
Chromatog. 23 398 2,282.8 0.6 3,804.7 322 41
a
AJI values vlere the average of three determinations
b
u ::;: units of esterase activity
I-
/--
55. FIGURE 8
Densitometer tracing of an acrylamide gel after
1 t h . f " . f· d U
1 t· f t Ae ec rop oreSlS 0 a purl le so U lon a es erase .
The gel was stained for protein with Coomassie Brilliant
Blue. The arrow indicates the position of esterase A as
determined in a duplicate acrylamide gel stained for
esterase activity_ No protein bands were visible in the
lower half of the gel.
57. 43
In the first few experiments, 0.1 M tris-Hel buffer,
pH 7.5 was used. Tris caused a significant loss in
enzymatic activity (~ 40%) when the same prep~was stored
at 4c over a period of several days.
phosphate, pH 7.5 was finally used.
Thus, 0.02 M potassium
Chromatography with hydroxylapatite, PAB-cellulose,
TEAE-cellulose and eM-cellulose was attempted but the results
were discouraging.
.Eli Stability. The stability of esterase A in 0.05 M
buffe~of different pH's, after 16 hours incubation at 4c
is shown in Figure 9 A. The enzyme is relatively stable
above pH 6.0, but rapidly loses activity below pH 5.0.
.Eli Optimum. The optimal pH o£ esterase A in 0.05 M
buffers was determined with the pH stat utilizing ethyl
ace tat e as the sub s t rat e (F i g u r e 9B ). A bra ad pH 0 P tim um
within the alkaline range tested was observed. A sharp
decrease in hydrolytic activity was observed below pH 6.0.
Substrate Specificity. Esterase activity on a variety
of aliphatic, aromatic and amino acid esters was determined.
The esters were checked at the concentrations stated.
Table 3 shows that as the acid moiety of ethanol esters
increases in carbon length, enzyme hydrolysis diminishes,
eliciting no observable hydrolytic activity when the side
chain contains four carbons. This same specifici"ty for the
carboxyl group of esters was shown with p-nitropbcnyl de-
rivatives, i.e. the enzyme's hydrolytic rate is only about
58. FIGURE 9
~ The pH stability of esterase A in 0.05 M buffers
( 0 - 0 ), pot ass i um ph 0 s p hat e ( X - X ), barbitol-
~. -. ), and carbonate-bicarbonate (0 - 0), assayed
at p5 7.5 with p-nitropbenyl acetate as the substrate,
after 16 hours incubation at 4c. The hydrolysis was
relative to a reference cell without enzyme. Each point
is an average of two determinations.
B. The pH optimum of esterase A with ethyl acetate
as the substrate in the presence of 0.05 M buffers (citrate
o - 0 ), potassium phosphate (X - X), and barbitol-Hel
.- .). The activity of the preparation was established
with subtraction of the hydrolysis rate without the additi6n
of enzyme preparation.
60. 45
TABLE 3
Influence of the chain length of the acyl group of
p-nitrophenyl and ethyl esters. Concentrations of
p-nitrophenyl and ethyl esters was 0.1 and 60 mM respect-
ively. Enzymatic activity for p-nitrophenyl derivatives
was determined spectrophotometrically and ethyl deriva-
tives determined by the pH stat method, and compared with
values obtained for p-nitrophenyl acetate and ethyl acetate
respectively.
Substrate
p-nitrophenyl acetate
p-nitrophenyl propionate
p-nitrophenyl butyrate
p-nitrophenYl caproate
p-nitrophenyl caprylate
p-nitrophenyl caprate
Ethyl acetate
Ethyl propionate
Ethyl butyrate
%Activity
(100)
13
a
o
o
o
(100)
11
o
61. one - e i g h th a s ra s t f n r p - nit r 0 ph e n y1. pro p ion ate (t h r e e c a. .1.' bon s )
a sit i s for p - nit r 0 ph en y 1 ace tat e (t woe arb () n s )) V;' ['~ i 1 e p:r p-
derivatives with four or more carbon atoms are not hydrolyzed
app~eciably (Table 3).
In order to determine the influence of the alkyl resi-
due upon esterase activity, the acyl group was kept constant
and the alcohol moiety varied from C
2
to C
ll
- The results
are given in Table 4. Ethyl acetate and phenyl acetate are
the substrates of choice,as well as p-nitrophenyl acetate
and 8-naphthyl acetate, not shown on the table. In general
increasing the alkyl chain length, decreases the activity
until negligible activity was demonstrated beyond Cg. Con-
versely, the variation of the acyl group appears to be of
more importance to maximum activity than the length of the
alkyl chain. The n-isomers or the alkyl group appeared to
be preferred over the branched isomers of propyl and butyl
acetate esters.
The highest enzyme activity was shown with the aromatic
acetate esters, p-nitrophenyl acetate and B-naphthyl acetate,
with activity toward indoxyl acetate being significant~y
lower (Table 5).
A number of amino acids esters were used as substrates,
in an effort to determine if esterase A possessed esterolytic
prnperties similar to those found with trypsin and chym:-
trypsin. Trypsin catalyzes hydrolysis of peptide, amide
or ester bonds of arginine or lysine residues; while 2hym~-
tr:vpsin acts only at bonds of tryptuphan, phenylalan.ine ::r'
62. Influence of the chain length of the alkyl group of acetate
esters. The concentration of esters was O.25M. Enzymatic
activity was measured by the pH stat method and compared
with values obtained for ethyl acetate.
*
Substrate
Ethyl acetate
n-propyl acetate
i-propyl acetate
n-butyl acetate
i-butyl acetate
t-butyl acetate
n-pentyl acetate
n-hexyl acetate
n-heptyl acetate
n-octyl acetate*
n-nanyl acetate*
n-decyl acetate*
n-undecyl acetate*
phenyl acetate*
indoxy1 acetate*
%Activity
100
71
50
66
.64
1
43
24
16
17
a
o
o
94
74
These esters were not fully solubilized urider the conditions
of the assay.
63. TABLE 5
Hydrolysis of aromatic and amino acid esters by esterase A
Ester
p-nitropheny1 acetate
S-naphthyl acetate
Indoxyl acetate
Tyrosine ethyl ester
N-acetyl-L-phenylalanine
methyl ester
N-acetyl-tyrosine methyl
ester
N-benzoyl-L-arginine ethyl
ester
N-benzoyl-L-tyrosine ethyl
ester
p-tosyl-L-arginine methyl
ester ·Hel
*
Concentration
(mM)
1.99
1.33
1.33
1.33
1.33
1.33
1.33
1 .. 33
Values extrapolated from Lineweaver-Burk Plot.
Specific
Activity
(ul mg )
72.5*
35.1*
2.6
o
o
o
o
o
o
64. 49
tyrosine residues (White etal., 1968). None of the amino
acid esters listed were hydrolyzed by esterase A (Table 5).
Km and Vmax. Three esterolytic assays, based on dif-
ferent principles, were used throughout this study. There-
fore, in order to make comparisons between the three assays,
it was necessary to determine the apparent affinity constant
(Km) of esterase A for each of these substrates and also the
amount of substrate turnover or lability of the ester linkage
to enzymatic attack (apparent Vmax). Esterase A showed the
following order in l/Km and Vmax: p-nitrophenyl acetate>
B-naphthyl acetate> ethyl acetate (Figure 10). The
Lineweaver-Burk plots were drawn according to the method of
least squares.
Inhibitors. Tables 6 and 7 show the effect of various
inhibitors on esterase A activity. The most potent inhibi-
tors were: mercuric chloride, diisopropylfluorophosphate,
eserine and sodium fluoride. The other inhibitors did not
show any significant inhibition at the concentrations tested
(Table 7).
Figure 11 shows that sodium chloride and potassium
chloride (0-1.0 M) have a considerable inhibitory effect on
esterase A activity, without prior incubation, at pH 7.55
with ethyl acetate as substrate. The same inhibitory pattern
could also be demonstrated with B-naphthyl acetate as the
substrate. The lack of inhibition of IO-1M sodium chloride
and potassium chloride (Table 7) is presumably due to a
dilution of inhibitor during assay; no such dilution occul"red
66. FIGURE 10
Lineweaver-Burk plots of esterase A with p-nitrophenyl
a~etate, 6-naphthyl acetate and ethyl acetate as substrates.
pn?A (0 - 0), SNA (0-0) and ethyl acetate (X - X).
67. Substrate
p- ni trophenyl aceta te
a-naphthyl acetate
Ethyl acetate
- .c
E
)(
I xGt
E
)(
-,
•0
E
E
-I>
- 800 - 600 - 400 - 200 o 200 400 600 800
1 M-1
[sf , m
Vmax
(Units/mg)
83.9
41.7
34.1
0
1000 1200 1400
Km
(mM)
.0013
.0021
.0435
0
1600 1800 2000
VI
f-J
68. 52
TABLE 6
Effect of esterase inhibitors on esterase A. Enzyme and
inhibitor incubated together at 37C for ten minutes~ then
assayed at room temperature via PNPA. Percent activity
remaining was based on the enzymatic hydrolysis of the
substrate under identical conditions but in the absence of
inhiliitor.
% Activity
Inhibitor Co nc . (l-1) Remaining
Mercuric Chloride 10- 3 0
10-4 49
10- 5 86
10-6
99
Diisopropy1f1uorophosphate 10- 3 3
10- 4 69
10- 5 99
Pheny1methy1 Sulfonylfluoride 10- 2
85
10- 3 94
10- 4 96
Eserine (Physostigmine) 10- 2
7
10- 3 61
10-1;. 90
Sodium Fluoride 10-1 36
10- 2 83
10- 3 101
71. DISCUSSION
One of the objectives of this study was to compare
este~ase A with known physical and chemical properties of
ot~er a~ety1 esterases and the extracellular proteases of
B. subtilis in an effort to reveal possible relationships.
The broad pH optimum for esterase A is not uncommon,
~.£. the acidic and basic proteases of B. subtilis show a
~ide pH range of activity (between pH 6.5 and 8.5 on BTEE
and casein; Boyer and Carlton, 1968), similarly undecyl
acetate esterase from Pseudomonas cepacia (Shum and
~a~kovetz, 1974a) exhibits a wide plateau from pH 7.0 to
8.5 f8T undecyl acetate and finally rat liver esterase
(Arndt and Krisch, 1973) has a broad enzyme activity from
pH 5.5 to 10.0 for methyl butyrate.
The stability of esterase A, between pH 6.0 and 10.0,
was previously shown for the alkaline and acid prot eases
of B. subtilis (Boyer and Carlton, 1968); these two enzymes
retained approximately 80% of their original activity after
40 hours incub~tion in this pH range.
~he alkaline protease (subtilisin) is proteolytic and
o~ly weakly esterolytic (Boyer and Carlton, 1968); it is
~~~e to hydrolyze such amino acid esters as N-toluenesulfonyl-
~-2~ginin2 methyl ester (TAME; Glazer, 1967) and N-benzoyltyro-
72. sine ethyl ester (BTEE; Ottesen and Spector, 1960) while only
,{eakly hydrolyzing ethyl acetate (Matsubara et al., 1958).
The neutral protease, containing zinc as a p.rcsthetic
group (McConn et al., 1964), possesses proteolytic and
aminopeptidase activity (Ray and Wagner, 1972) but is not
esterolytic (Feder, 1967).
The acid protease displays high estero1ytic activity
but low proteolytic activity (Millet, 1970)~ It exhibits
strong esterolytic activity on BTEE (Boyer and Carlton,
1968), which is used to detect chymotrypsin endopeptidase
activity (Hummel, 1959). It also attacks p-nitrophenyl
acetate (PNPA; Millet, 1970), naphthyl acetate (NA; Millet;
1970), benzoyl-L-arginine ethyl ester (BAEE; Hageman and
Car1ton, 1970; Prestidge et al., 1971), acetYI-L-tyrosine
ethyl ester (ATEE; Hageman and Carlton, 1970; Prestidge
et al., 1971).
The inability of esterase A to hydrolyze amino acid
esters, the lack of inhibition with L-l-(p-Toluenesulfonyl)
amino-2-phenylethylchloromethyl ketone (TPCK), specific for
chymotrypsin (White et al., 1968), and finally the high
specificity for short acyl groups suggested that a search
for other protein substrates would have been futile;
therefore, no attempt was made to further characterize
esterase A with respect to protein substrates.
73. The differences in substrate specificity of the three
extracellular proteases and esterase A indicate that all
four enzymes are different.
57
Other serine hydrolases are: a-chymotrypsin, trypsin,
elastase, thrombin and plasmin (Bender, Kezdy and Wedler,
1967) ..One of the best characterized is a-chymotrypsin. It
hydrolyzes p-nitropheny1 derivatives in the order: acetate <
butyrate < valerat e (Stoops et ·al., 1969). In addition , it
displays high esterolytic activity with amino acid esters, ~.&.
acetyl-L-tyrosine ethyl ester (ATEE) and acetyl-L-tyrosinamide
(White et al., 1968).
Chow and Ecobichon (1972) isolated twelve guinea pig
liver esterases and fourteen renal esterases; these were
identified as carboxylesterases. The hepatic esterases hydro-
lyzed p-nitrophenyl esters in the decreasing order of PNP-
butyrate> PNP-propionate > PNP-acetate. The renal esterases
attacked PNP-butyrate and PNP-propionate at the same rate
and much faster than PNP~acetate; this specificity was in the
reverse order of that found for esterase A. Two distinct
liver enzymes were detected (mol. wt. : 240,000 and 56,000);
the larger molecule is believed to be a tetramer, as is
thought to be the case for esterase A. The renal enzyme had
a molecular weight of 56,000.
Arndt and Krisch (1973) determined the effect of alkyl
and acyl groups on substrate affinity and hydrolytic rate
74. 58
of rat liver esterase. Two trends were observed: 1) Vmax
inc rea sed (ab 0 u t f i v -:~ f 0 1 d) a s the a c y 1 g r 0 up e 1 0 n gat e d
from C2 to C4, then dropped abruptly for C 5 and C6; the
length of the acyl group ha~ no effect on Km, 2) As the
alcohol chain lengthened from Cl to C3 , Vmax increased then
sharply decreased for C4, while Km increased and plateaued
No studies were undertaken to determine the
effects of the acyl and alkyl groups on Km and Vmax for
esterase A; however, rat liver esterase appears to share
esterase A's specificity for substrates with acyl and alkyl
groups of four carbons .or le-ss.
An esterase was partially puri~ied (54 fold) from an
adult rat brain .. The Vmax for a-naphthyl ester derivatives
d e'c r e 9. sed wit han inc rea s e in the c h a in 1 eng tho f the a. c y 1
group. The acetate est~r was hydrolyzed thirty-four times
faster than butyrate and approximately seven times as fast
as propionate (Dabich et al., 1968). Although the hydrolysis
of a-naphthyl ester derivatives was not determined for
esterase A, the enzyme hydrolyzed PNP-acetate about eight
times faster than PNP-propionate. Hence an increase in
length of the acyl group caused a similar decrease in
hydrolytic rate for rat brain esterase and esterase A.
Hofstee (l971) investigated the hydrolysis of n-fatty
acid esters of m-hy-droxybenzoic acid by liver esterases
from: pig, horse, cow, rat, rabbit, pig~on and chicken.
As substrate concentration increased, each enzyme's
75. reaction rate increased more rapidly than Michaelis-Menten
theory predicted (i.~. substrate activation occurred).
Purified esterase A showed no deviations from typical
Michaelis-Menten kinetics (Figure 10).
The most potent inhibitor tested on esterase A was
mercuric chloride, which serves as a non-specific protein
precipitating agent (White et a1., 1968) Table 6r
59
As expected, Esterase A appears to possess a serine
hydroxyl group at its active site because or its inhibition
by DFP. Another inhibitor specific for serine hydroxyl
groups, PMSF, showed much less inhibition, possibly because
it has less ability to penetrate the enzyme's active site;
alternatively, the ten minute incubation time may have been
inadequate for PMSF to exert its full inhibitory effect.
Sodium fluoride, at high concentrations, showed a
considerable degree of inhibition on esterase A. The
inhibition with DFP apparently was not caused by the flu-
oride ions because at a concentration of IO-3M, DFP was
g~eat1y inhibitory while sodium fluoride showed no inhibition
(Shum and Markovetz, 1974b)~
Physostigmine (eserine) completely inhibits cholines-
terases at a concentration of lO-5M, while requiring higher
concentrations to inhibit other esterases (Holmes and
Masters, 1969; Krisch, 1971). Hence~ esterase A does not
appe2.r to be 8.. cholinesterase.
The sulfhydryl specific inhibitors peME and iodoaceta-
mide, did not inhibit the enzyme significantly. Like-r..rise,
76. 2-mercaptoethanol, which breaks disulfide bonds, did not
inhibit esterase A. This result was expected, for it is
known that SH-groups are not essential for the activity of
other esterases (Krisch, 1973).
60
The detergents, chelating agents and other salts were
ineffective inhibitors at the concentrations tested (Table 7 ).
The partial inhibition of esterase A when assayed in
the presence of NaCl or KCl (0-1.0 M), at a constant pH
(Figure 11) is presumably due to a chloride effect, because
no such inhibition was observed with potassium phosphate at
the same concentrations.
The inhibition by organophosphates and the wide
specificity for aliphatic and aromatic esters~suggests that
esterase A is similar to the non-specific carboxy1esterases
(EC 3.1.1.1) or B-esterases (Whittaker, 1972).
The characteristics of esterase A that are shared by
the undecyl acetate esterase from Pseudomonas cepacia (Shum
and Markovetz, 1974b) are as follows: both enzymes have a
broad pH optimum, there are similarities·in inhibition (with
the exception of PCME which strongly inhibits undecyl acetate
esterase), the two enzymes exhibit high enzymatic activity,
with aromatic acetate esters (p-nitrophenyl acetate and
naphthyl acetate) and do not hydrolyze BTEE. However, the
specificity of the two enzymes for the alkyl group of
substrates appears to be different, i.e. undecyl acetate
esterase shows increasing activity as the chain lertgthens~
77. 61
while esterase A decreases in reactivity.
At present, the predominant organisms known to utilize
long chain methyl ketones are Gram-negative rod shaped
bacteria. The induction of an esterase in a Bacillus species
during methyl ketone oxidation would strengthen the proposed
catabolic pathway for Pseudomonas (Forney and Markovetz~
1969) as a general phenomenon for r~cycling these organic
molecules and would explain the role of intracellular
esterases.
Attempts to inhibit esterase A activity with unbranched
alcohols from C1 to C1l(not listed) have been unsuccessful.
In addition, cultures of B. subti"lis 168 and a mutant, EB-l,
lacking esterase B activity could not be induced to produce
higher levels of esterase activity when grown in the presence
of 2-tridecanone.
The goal of this study was to compare esterase A to the
extracellular (acid) protease and tbe intracellular esterase
B of B. subtilis; the underlying premiss being that one or
more of these enzymes might be involved in the sporulation
process.
It now seems clear that esterase A and the acid protease
are different enzymes because of, their differences in
suostrate specificity. In addition, it appears that esterase
A is not a precursor of esterase B and is not necessary for
the sporulation process (Uigerd, in preparation). The
function of esterase A and B and the acid protease of B.
subtilis awaits future investigations. It may be that one
78. 62
or more of' these enzymes is invo1ved l~.: re:~ylcing methyl
ketones, as proposed for the u~decyl acetate esteras~ from
p ~~ t.! udc uo na s ceuacia
-,.<'--- (Forney and Markovetz, 1969).
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