-
3~T\
/J8U
Mo. Hill
PYRIMIDINE BIOSYNTHESIS IN THE GENUS Streptomyces.
CHARACTERIZATION OF ASPARTATE TRANSCARBAMOYLASE
AND ITS INTERACTION WITH OTHER
PYRIMIDINE ENZYMES
DISSERTATION
Presented to the Graduate Council of the
University of North Texas in Partial
Fulfillment of the Requirements
for the Degree of
DOCTOR OF PHILOSOPHY
By
Lee E. Hughes, B.A., M.S,
Denton, Texas
May, 1998
-
Hughes, Lee E., Pyrimidine biosynthesis in the crenus
Streptomyces: Characterization of aspartate
transcarbamoylase and its interaction with other pyrimidine
enzymes. Doctor of Philosophy (Microbiology), May, 1998, 188
pp., 16 tables, 43 illustrations, references, 181 titles.
Aspartate transcarbamoylase (ATCase) of Streptomyces
was characterized and its interaction with other pyrimidine
enzymes explored. ATCase and dihydroorotase (DHOase) of S.
griseus were assayed and purified using conventional methods
and HPLC. The two activities were found to co-purify,
suggesting both are found in a complex. The S. griseus
holoenzyme has an estimated molecular mass of 480 kDa.
Examination by SDS-PAGE revealed that the holoenzyme is
composed of two types of subunits. Western blot analysis
using antibody to E. coli PyrB (ATCase catalytic subunit)
showed a cross reaction with the 38 kDa subunit from the S.
griseus ATCase/DHOase complex. Only two other bacterial
ATCases, from Deinococcus and Thermus, have been reported to
contain both ATCase and DHOase catalytically-active
subunits. The similarity of these ATCases to that of S.
griseus is surprising considering the wide divergence
between these organisms and streptomycetes.
The S. griseus ATCase showed typical Michaelis-Menten
kinetics for velocity versus substrate plots for both
aspartate and carbamoylphosphate. These kinetics and the
observed molecular mass place Streptomyces ATCase in the
-
Class A bacterial ATCases. Like other Class A enzymes, S.
grriseus ATCase was inhibited by ATP, CTP, and UTP.
Interactions between the biosynthetic and salvage
pathways were also found. A 5-fluorouracil resistant mutant
of S. griseus was selected which lacked uracil
phosphoribosyltransferase (Upp) activity. This strain was
found to be derepressed 8-10 fold for ATCase and DHOase.
Furthermore, ATCase in this strain was more sensitive to
effector molecules than was the wild-type enzyme.
Considering the positive selection method used, there is
likely to be a mutation in only one gene, upp. Therefore,
the observed loss of Upp activity, derepression of the
pathway, and increased effector sensitivity are linked.
This would be possible only if the same polypeptide were
involved in each.
Genetic studies using PCR showed that the Streptomyces
pyrimidine operon is organized similarly to that of
Mycoba cterium.
-
3~T\
/J8U
Mo. Hill
PYRIMIDINE BIOSYNTHESIS IN THE GENUS Streptomyces.
CHARACTERIZATION OF ASPARTATE TRANSCARBAMOYLASE
AND ITS INTERACTION WITH OTHER
PYRIMIDINE ENZYMES
DISSERTATION
Presented to the Graduate Council of the
University of North Texas in Partial
Fulfillment of the Requirements
for the Degree of
DOCTOR OF PHILOSOPHY
By
Lee E. Hughes, B.A., M.S,
Denton, Texas
May, 1998
-
ACKNOWLEDGMENT
I would like to gratefully acknowledge that this work
was made possible in part by the support of a Grant-in-Aid
of Research from Sigma Xi, The Scientific Research Society.
111
-
TABLE OF CONTENTS
Page
LIST OF TABLES vi
LIST OF ILLUSTRATIONS vii
INTRODUCTION 1
Pyrimidine biosynthetic pathway 4 Genetic organization 6 Aspartate transcarbamoylase 13 Dihydroorotase 25 Streptomyces 35
METHODS 44
Bacterial strains 44 Media and growth conditions 44 Streptomyces spore suspension 46 Selection of 5-fluorouracil resistant strain 49 Harvesting of bacterial cultures 50 Preparation of cell extracts 51 Aspartate transcarbamoylase assay 52 Dihydroorotase assay 55 Growth curve 56 Protein concentration determination 57 Purification strategies 58 Streptomycin sulfate precipitation .59 Ammonium sulfate fractionation 59 Sephadex G-200 column chromatography 61 Sephacryl S-400 column chromatography 62 Concentration of size-exclusion fractions 62 High performance liquid chromatography ion-exchange .63 Nondenaturing polyacrylamide activity gels 63 Sodium dodecyl sulfate denaturing polyacrylamide gel electrophoresis 66
Western blot 68 Isolation of chromosomal DNA 70 Design of oligonucleotides for polymerase chain
reaction 72 Polymerase chain reaction 73
xv
-
TABLE OF CONTENTS (continued)
Page
RESULTS 80
Carbamoylaspartate standard curve 80 Determination of optimum pH 80 Linearity of ATCase activity . .82 Growth curve 82 Stability of Streptomyces enzymes 88 Purification of ATCase/DHOase 92 Kinetics of ATCase 112 Effector response of ATCase 121 DHOase 121 Transcriptional control 138 Polymerase chain reaction products 138
DISCUSSION 145
Pyrimidine enzyme levels during growth phases . . . 145 Enzyme complex 147 Genetic organization 149 Stability of Streptomyces enzymes 153 Kinetics and effector response of ATCase 155 Conclusions 159
REFERENCES 161
v
-
LIST OF TABLES
Table Page
1. Linkage map locations of pyrimidine genes in E. coli
and S. typhimurium 6
2. Overview of properties of bacterial ATCase classes . .17
3. Properties of known bacterial Class A ATCases 26
4. Properties of known bacterial Class B ATCases 28
5. Properties of known bacterial Class C ATCases 29
6. Properties of known bacterial ATCases - class not yet described 30
7. Properties of known eukaryotic ATCases - plant and other monofunctional 31
8. Properties of known eukaryotic ATCases -
multifunctional 33
9. Bacterial strains 45
10. Oligonucleotide primers for PCR 74
11. Results of various strategies for purification of S. griseus ATCase 100
12. Results of various strategies for purification of S. griseus DHOase 102
13. Representative purification of S. griseus ATCase and DHOase for SDS-PAGE 104
14. Determination of molecular mass (kDa) of ATCase enzyme in Streptomyces Ill
15. Comparison of percentage of inhibition of ATCase by effector molecules 126
16. Comparison of concentration of effectors necessary for 50% inhibition of ATCase specific activity . . 135
vx
-
LIST OF ILLUSTRATIONS
Figure Page
1. Pyrimidine biosynthetic pathway 2
2. Schematic diagram of bacterial pyrimidine operons... 8
3. Schematic diagram of the S. cerevisiae ura2 locus. . .11
4. Schematic diagram of the Dictyostelium, Drosophila,
and hamster CAD locus 12
5. The reaction catalyzed by ATCase 14
6. Classes of bacterial ATCase 15
7. Model of the domain structure of CAD 22
8. Genetic map of S. coelicolor A3(2) showing relative positions of markers conferring uracil requirement .43
9. Apparatus used for preparation of spore suspensions
10. Standard curve of carbamoylaspartate concentration
11. Effect of pH on ATCase activity of S. griseus 10137
12. Effect of pH on ATCase activity of S. venezuelae .
13. Effect of pH on ATCase activity of S. lividans . .
.47
.81
.83
.84
.85
14. Change in absorbance over time for S. griseus 10137 ATCase activity 86
15. S. griseus 10137 extract concentration and ATCase activity 87
16. Dry weight determination, ATCase specific activity, and DHOase specific activity during growth 89
17. Activity of ATCase of S. griseus 10137 after storage at various conditions 90
18. Activity of DHOase of S. griseus 10137 after storage at various conditions 93
VIX
-
LIST OF ILLUSTRATIONS (continued)
Figure Page
19. Activity of ATCase and DHOase after fractionation on a Sephacryl S-400 column 95
20. Activity of ATCase and DHOase after fractionation on an HPLC ion-exchange column 97
21. (a) SDS-PAGE of partially-purified S. griseus ATCase/DHOase. (b) Western blot of SDS-PAGE using antibody to E. coli PyrB 105
22. Activity of ATCase and DHOase of S. griseus 10137 extract after fractionation on a Sephadex G-200 column 106
23. Activity of ATCase and DHOase of S. griseus 10137 extract, which had been stored for six weeks, after fractionation on a Sephadex G-200 column 108
24. ATCase activity on 4-20% gradient polyacrylamide gel 110
25. Activity of ATCase and DHOase of S. venezuelae extract after fractionation on a Sephadex G-200 column 113
26. Activity of ATCase and DHOase of S. lividans extract after fractionation on a Sephadex G-200 column
27. Velocity versus substrate plot for S. griseus 10137 ATCase as a function of aspartate concentration. . 117
28. Lineweaver-Burk plot for S. griseus 10137 ATCase as a function of aspartate concentration 118
29. Velocity versus substrate plot for S. griseus 10137 ATCase as a function of carbamoylphosphate concentration
30. Lineweaver-Burk plot for S. griseus 10137 ATCase as a function of carbamoylphosphate concentration . . 120
vixi
-
LIST OF ILLUSTRATIONS (continued)
Figure Page
31. Velocity versus substrate plot for S. griseus 10137 ATCase with changing aspartate concentration in the presence and absence of effector molecules . . . . 122
32. Lineweaver-Burk plot for S. griseus 10137 ATCase with changing aspartate concentration in the presence and absence of effector molecules 123
33. Velocity versus substrate plot for S. griseus 10137 ATCase with changing carbamoylphosphate concentration in the presence and absence of effector molecules. 124
34. Lineweaver-Burk plot for S. griseus 10137 ATCase with changing carbamoylphosphate concentration in the presence and absence of effector molecules . . . . 125
35. Effector response of S. griseus ATCases in the presence of changing concentrations of ATP 127
36. Effector response of S. griseus ATCases in the presence of changing concentrations of CTP 129
37. Effector response of S. griseus ATCases in the presence of changing concentrations of GTP 131
38. Effector response of S. griseus ATCases in the presence of changing concentrations of UTP 133
39. Velocity versus substrate plot for S. griseus 10137 DHOase as a function of dihydroorotate concentration 136
40. Lineweaver-Burk plot for S. griseus 10137 DHOase as a
function of dihydroorotate concentration 137
41. Products from PCR of S. coelicolor M145 DNA 139
42. M. tuberculosis operon with PCR primer locations and orientations 140
43. Probable Streptomyces operon with PCR primer locations and orientations 143
xx
-
INTRODUCTION
The role of pyrimidines as building blocks in the
informational macromolecules ribonucleic acid (RNA) and
deoxyribonucleic acid (DNA) makes the study of pyrimidine
synthesis and regulation important. Along with purines, the
pyrimidines are essential for cellular growth and for the
passing of genetic information to subsequent generations.
Pyrimidines are found in all organisms and are six-membered,
aromatic heterocyclic ring compounds. Pyrimidine
nucleosides consist of a pyrimidine base plus a pentose
sugar, generally either ribose or 2'-deoxyribose, while
nucleotides are nucleosides plus one or more phosphate
groups. The pyrimidine ring is a component of coenzymes
such as nicotinamide adenine dinucleotide (NAD) and acetyl
coenzyme A (acetylCoA), and nucleotides are used in the
formation of activated intermediates in carbohydrate and
lipid metabolism.
There are six enzymatic steps in the biosynthesis of
uridine-5'-monophosphate (UMP; Fig. 1), which itself serves
as a precursor for all the pyrimidine nucleotides. The
pathway appears to be universal and follows the same
sequence in all organisms thus far studied (0'Donovan &
Neuhard, 1970; Grogan & Gunsalus, 1993). This pathway has
been studied extensively in bacteria, fungi, plants, and
-
Figure 1. Pyrimidine biosynthetic pathway in Escherichia
coli and Salmonella typhimurium. Gene symbols and the
enzymes they encode are: ndk - nucleoside diphosphate
kinase; pyrA (carAB) - carbamoylphosphate synthetase; pyrBI
- aspartate transcarbamoylase; pyrC - dihydroorotase; pyrD -
dihydroorotate dehydrogenase; pyrE - orotate
phosphoribosyltransferase; pyrF - OMP decarboxylase; pyrG -
CTP synthetase; pyrH - UMP kinase. Adapted from Neuhard &
Nygaard (1987) .
-
CD C "c
as
X +
X Q <
X
X O O O =o
CL Q_
DC O-s
Q_ CL X
M Q <
0=0 Z X I 0=0
1 - Q
x Q -
X o O O X -O,
0 = 0
- Q
~o \ . * 0 \ o co Z X -o O
/ £1 T> O
X y -
/ X O o o X -e>
0 = 0 zx
X Z °x 8
-
animals. Although the sequence is virtually the same in
most organisms, regulation of the pathway and organization
of the enzymes vary in different organisms.
Pyrimidine biosynthetic pathway.
The first step in the synthesis of pyrimidines is
catalyzed by the enzyme carbamoylphosphate synthetase
(CPSase, EC 6.3.5.5). The reaction utilizes bicarbonate,
ammonium ions or glutamine, and two molecules of adenosine-
s' -triphosphate (ATP) in the formation of one molecule of
carbamoylphosphate and adenosine-5'-diphosphate (ADP)
(Anderson & Meister, 1965; Kalman et al., 1966). The
glutamine amidotransferase activity is analogous to that
found in a later reaction of the pathway, CTP synthetase.
Carbamoylphosphate is required for both arginine and
pyrimidine synthesis (Abdelal et al., 1969). While bacteria
other than Bacillus possess only one CPSase, which satisfies
both pathways, eukaryotes possess two CPSase enzymes, one
for each pathway.
The formation of carbamoylaspartate (CAA) by aspartate
transcarbamoylase (ATCase, EC 2.1.3.2) is the first
committed step in pyrimidine biosynthesis. Aspartate is
carbamoylated at the amino group, producing CAA and
releasing inorganic phosphate. The structure and regulation
of ATCase is discussed in detail elsewhere in this paper.
-
The next reaction involves the cyclization of CAA, with
the release of a molecule of water, to produce
dihydroorotate (DHO). This step is catalyzed by the enzyme
dihydroorotase (DHOase, EC 3.5.2.3). This enzyme is
discussed in more detail later in this paper. In the
following step, DHO is oxidized to orotate (OA) in a
reaction catalyzed by dihydroorotate dehydrogenase
(DHOdehase, EC 1.3.3.1). The first pyrimidine nucleotide is
then produced by the transfer of ribose-5'-phosphate from
5'-phosphoribosyl-1'-pyrophosphate (PRPP) to OA to form
orotidine-5'-monophosphate (OMP), a reaction catalyzed by
orotate phosphoribosyltransferase (OPRTase, EC 2.2.4.10).
OMP is decarboxylated by the enzyme OMP decarboxylase
(OMPdecase, EC 4.1.1.23) in the final step in the production
of UMP.
The pyrimidine nucleoside triphosphates, uridine-5'-
triphosphate (UTP) and cytidine-5'-triphosphate (CTP), are
ultimately produced from UMP. In sequential steps, UMP is
first phosphorylated to uridine-5'-diphosphate (UDP) by the
highly specific UMP kinase (EC 2.7.4.4). UDP is further
phosphorylated by a non-specific enzyme, nucleoside
diphosphate kinase (NDK, EC 2.7.4.6), to form UTP. UTP is
converted to CTP by the enzyme CTP synthetase (EC 6.3.4.2),
which transfers an amino group from glutamine in a reaction
analogous to that of the first step in the pathway, CPSase.
-
Table 1. Linkage map locations of pyrimidine genes in E. coli and S. typhimurium (Bachmann, 1987; Sanderson & Hurley, 1987; Neuhard & Kelln, 1996).
Gene (enzyme)
Map location (minutes)
Gene (enzyme) E. coli S. typhimurium
carAB/pyrA (CPSase) 0.6, 0.7 1
pyrBI (ATCase) 96.3 98
pyrC (DHOase) 24.2 23
pyrD (DHOdehase) 21.6 20
pyrE (OPRTase) 82 .1 79
pyrF (OMPdecase) 28.8 33
Genetic organization.
In many bacterial systems, the six enzymatic steps of
pyrimidine biosynthesis are encoded by unlinked genes. In
Escherichia coli K-12 and Salmonella typhimurium, the
pyrimidine genes are scattered throughout the genome (Table
1) .
Likewise, the genes in Pseudomonas are found at
different chromosomal locations (Holloway et al., 1990).
However, in both P. putida and P. aeruginosa, the
scaffolding subunit of ATCase is a catalytically-inactive,
DHOase-like protein which is encoded by a gene immediately
downstream of the ATCase catalytic subunit gene, pyrB (Fig.
2). In fact, there is a four base pair overlap of these two
open reading frames (Schurr, 1992; Vickrey, 1993). The
catalytically active DHOase is encoded by a separate pyrC
-
located in another region of the chromosome along with argG
(D. Brichta, personal communication).
In Bacillus, the genes for pyrimidine biosynthesis are
organized into a single operon (Quinn et al., 1991). The
segment of the B. subtilis chromosome containing this gene
cluster has been sequenced and found to contain eight
overlapping cistrons encoding the six enzymes of pyrimidine
biosynthesis, as well as genes for uracil permease (pyrP)
and a regulatory protein (pyrR) (Turner et al., 1994) (Fig.
2) . A similar genetic arrangement has been found in B.
caldolyticus (Ghim et al., 1994; Ghim & Neuhard, 1994).
Other bacterial pyrimidine operons have also been described
(Fig. 2).
The mechanism for control of gene expression has been
reported in two of these bacterial systems. The pyrBI
operon of E. coli is regulated by a rho-independent
attenuator sequence (Roof et al., 1982). Limitation of
pyrimidine triphosphates would cause RNA polymerase to pause
in the pyrimidine-rich tracts in that area. This pause
permits the ribosome to reach this region, coupling
transcription and translation. The ribosome disrupts the
formation of the terminator hairpin and allows read-through
by the RNA polymerase to the pyrBI structural genes.
Likewise, the B. subtilis pyr gene cluster is preceded by
three sets of mutually exclusive antiterminator/terminator
-
Figure 2. Schematic diagram of bacterial pyrimidine operons
(Not drawn to scale). Overlapping genes are shown on
separate lines. Gene symbols and the enzymes they encode:
bbc, orf2, orfl, x - unknown; pyrAA - glutaminase; pyrAB -
carbamoylphosphate synthetase; pyrB - aspartate
transcarbamoylase; pyrC - dihydroorotase; pyrC' - inactive
dihydroorotase-like; pyrD, pyrDa, pyrDb - dihydroorotate
dehydrogenase; pyrE - orotate phosphoribosyltransferase;
pyrF - OMP decarboxylase; pyrl - ATCase regulatory subunit;
pyrP - uracil permease; pyrR - regulatory protein; upp -
uracil phosphoribosyltransferase. Adapted from: Bacillus
subtilis (Turner et al., 1994); B. caldolyticus (Ghim et
al., 1994); Lactobacillus plantarum (Elagoz et al., 1996);
L. leichmannii (Schenk-Groninger et al., 1995); Clostridium
acetobutylicum (Genome Therapeutics Corp., available on the
Internet at http://pandora.cric.com/htdocs/sequences/
clostridium/clospage.html); Pseudomonas aeruginosa (Vickrey,
1993); P. putida (Schurr et al., 1995); Thermus aquaticus
(Van de Casteele et al., 1994); Mycobacterium leprae; and M.
tuberculosis (PhiHipp et al., 1996).
http://pandora.cric.com/htdocs/sequences/
-
B. subtilis
pyrR pyrP pyrB pyrC pyrAA pyrAB or£2 pyrD pyrF pyrE
B. caldolyticus
pyrR pyrP pyrB pyrC pyrAA pyrAB orf2 pyrD pyrF pyrE
L. pi ant arum
pyrR pyrB pyrC pyrAA pyrAB pyrD pyrF pyrE
L. leichmannii
pyrB pyrC
C. acetobutylicum
pyrB pyrl pyrDb pyrDa pyrF
P. aeruginosa and P. putida
pyrR pyrB pyrC'
T. aquaticus
upp pyrB bbc pyrC
M. tuberculosis and M. leprae
pyrR pyrB pyrC orfl pyrAA pyrAB pyrF
-
10
structures which may be regulated by a protein-mediated
equilibrium (Turner et al., 1994).
There are also several known schemes for pyrimidine
gene organization in eukaryotes. In Saccharomyces
cerevisiae, genetic analysis has shown that five independent
genes are involved in the biosynthesis of UMP and expression
of these genes are sequentially induced by intermediates of
the pathway (Lacroute, 1968) . In this yeast, the activities
of the first two enzymes, CPSase and ATCase, are coded by
the same genetic region (ura2) and form a single enzymatic
complex. The four enzymes that follow later in the pathway
are encoded by separate genetic loci and are induced in a
sequential way by the intermediary products. This same
genetic organization has been found for Neurospora crassa
(Caroline, 1969). Sequencing of the ura2 locus in S.
cerevisiae showed that this gene includes a DHOase-like
domain between the CPSase and ATCase domains (Souciet et
al., 1989) (Fig. 3). The same has been found for
Schizosaccharomyces (Lollier et al., 1995).
In the slime mold Dictyostelium discoideum, in
Drosophila melanogaster, and in mammals, a similar genetic
arrangement to that of yeast is found for the first three
enzymatic activities of the pyrimidine pathway (Rawls &
Fristrom, 1975; Freund & Jarry, 1987; Faure et al., 1989)
(Fig. 4). However, in these cases, the CPSase, DHOase and
ATCase domains are all catalytically active (Coleman et al..
-
11
Figure 3. Schematic diagram of the S. cerevisiae ura2
locus. Numbers indicate the number of base pairs from the
start of the gene. White spaces are interdomain linker
regions. Adapted from Souciet et al. (1989).
400 440
ii IlllpllMll 1
1480 1490 1819 1907 2212
GATase domain DHOase-like domain
CPSase domain ATCase domain
-
12
Figure 4. Schematic diagram of the Dictyostelium,
Drosophila, and hamster CAD locus. White spaces are
interdomain linker regions. Adapted from Davidson et al.
(1993).
GATase domain DHOase domain
CPSase domain ATCase domain
-
13
1977; Simmer et al., 1989, 1990a, 1990i>) . This
multifunctional protein is known as CAD or multienzyme
pyrl-3 (Jones, 1980) . A similar situation has also been
shown for the final two steps of UMP biosynthesis in these
organisms. A single polypeptide, UMP synthase or
multienzyme pyr5, 6, contains both OPRTase and OMPdecase
activities (Jones, 1980). Jones (1980) suggests that a
protein similar to multienzyme pyrl-3 may exist in all
animals higher than Diptera.
In other eukaryotes, though, these multienzymic,
pyrimidine proteins are not found. In Chlorella, a green
alga, ATCase and DHOase activities reside on different
proteins (Dunn et al., 1977), while the activities for
CPSase, ATCase and DHOase have also been shown to be on
separate proteins in higher plants (Yon, 1972; Achar et al.,
1974; Doremus, 1986; Bartlett et al., 1994). For parasitic
protozoans, biosynthesis of pyrimidines is catalyzed by six
discrete enzyme activities (Krungkrai et al., 1990).
Aspartate transcarbamoylase.
ATCase catalyzes the transfer of the carbamoyl group of
carbamoylphosphate to the a-amino group of L-aspartate to
form CAA and phosphate (Fig. 5). As the enzyme catalyzing
the first step unique to pyrimidine biosynthesis, the study
of the structure and regulation of ATCase is important in
-
14
COO" COO"
I I CH2 0 CH2 0 I II I II CH2-NH, + C-NH2 CH2-NH2-C-NH2 + P04"
3
I 1 , 1 COO" O-PO3"2 COO"
Aspartate Carbamoyl- Carbamoyl- Phosphate phosphate aspartate
Figure 5. The reaction catalyzed by aspartate transcarbamoylase.
understanding the catalytic mechanism and evolutionary
history of this enzyme in different organisms.
The catalytic activity of ATCase resides in a trimer of
identical subunits. This activity is dependent on the
formation of an active site from half-sites on separate
subunits (Honzatko et al., 1982; Rosenbusch & Weber, 1971;
Stevens et al., 1991). Each trimeric unit forms three
active sites. Regulation of enzyme activity is accomplished
in a different manner in each of the types of ATCase
discussed below.
Three classes of bacterial ATCase have been described
(Fig. 6), with varying molecular weights of the holoenzyme,
quaternary structure, and enzyme kinetics (Bethell & Jones,
1969; Wild et al., 1980) (Table 2).
-
15
Figure 6. Classes of bacterial ATCase. Adapted from Bergh
& Evans (1993).
-
Class A ATCases
Catalytic Trimer
16
45 kDa
34 kDa
Class B ATCases
Catalytic Trimer
Regulatory Dimer
17 kDa
34 kDa
Class C ATCases
Catalytic Trimer • 34 kDa
-
17
Table 2. Overview of properties of bacterial ATCase classes.
Class Properties
A e.g. Pseudomonas aeruginosa
~500 kDa molecular mass Dodecamer, only holoenzyme active Michaelis-Menten kinetics Inhibition by ATP, CTP, UTP
B e.g. Escherichia coli
~300 kDa molecular mass Dodecamer, catalytic trimer also active
Sigmoidal kinetics for dodecamer, Michaelis-Menten for trimer
Allosteric Activation by ATP Inhibition by CTP, UTP
C e.g. Bacillus subtilis
~100 kDa molecular mass Trimer Michaelis-Menten kinetics No effector response
The largest ATCases are found in Class A. One example,
fluorescens ATCase, has a complex with a molecular mass
of 464 kDa. This complex appears to consist of a 1:1 ratio
of 34 kDa and 45 kDa polypeptides (Bergh & Evans, 1993) .
The presence of two polypeptides is consistent with recent
sequencing studies in P. aeruginosa and P. putida which show
two open reading frames encoding polypeptides of
approximately similar sizes (Schurr, 1992; Vickrey, 1993;
Schurr et al., 1995). Based on the mass of the constituent
subunits and the stoichiometry of the complex, the protein
must be a dodecamer (Bergh & Evans, 1993). Similar findings
have also been reported for P. fluorescens and P. syringae
(Shepherdson & McPhail, 1993). The ATCase of P. putida has
-
18
been shown to have a molecular mass of 482 kDa, with the
smaller catalytic polypeptide having a molecular mass of
36.4 kDa. The larger polypeptide, 44.2 kDa, is encoded by
the DHOase-like gene but does not have DHOase activity. The
DHOase-like subunit is required for activity in the
holoenzyme, as no activity has been found solely for the
catalytic trimers (Schurr et al., 1995). Characteristics
for Class A enzymes include hyperbolic substrate saturation
curves and inhibition by ATP, UTP, and CTP which is
competitive with carbamoylphosphate and noncompetitive with
aspartate (Neumann & Jones, 1964; Bethell & Jones, 1969;
Linscott, 1996). The nucleotide effector binding site has
been localized to the catalytic polypeptide, not to the 45
kDa polypeptide (Bergh & Evans, 1993; Schurr et al., 1995).
Class A ATCases have been found to be widespread (Table 3).
The Class B ATCases are smaller and are distinguished
by sigmoidal substrate saturation curves as typified by the
enzyme from E. coli. The enzyme is a dodecamer. Six
catalytic polypeptides, each with a molecular mass of 34
kDa, are grouped into two trimers. The trimers are
connected by three regulatory dimers. Each subunit of the
regulatory dimer has a molecular mass of 17 kDa (Kantrowitz
& Lipscomb, 1988). The holoenzyme molecular mass is 306 kDa
(Fig. 6). Activity of the trimers is retained when they are
separated from the regulatory subunits by chemical means,
such as p-chloromercuribenzoate (Blackburn & Schachman,
-
19
1977; Subramani & Schachman, 1981). The enzyme is under
allosteric control, whereby the activity is influenced
through the binding of effector molecules at a site other
than the active site. The nucleotide binding site is
located between pairs of regulatory subunits. E. coli
ATCase is inhibited by physiological levels of CTP (Yates &
Pardee, 1956; Gerhart & Pardee, 1962, 1964) and activated by
ATP. Subunit interactions are responsible for sigmoidal
concentration curves for both aspartate (Gerhart & Pardee,
1962) and carbamoylphosphate (Bethell et al., 1968). Class
B ATCases have been isolated from several representatives of
the Enterobacteriaceae (Wild et al., 1980), as well as other
Gram negative bacteria and even an archaebacterium,
Pyrococcus abyssi (Purcarea et al., 1994) (Table 4).
The Class C ATCases are characterized by their small
size, insensitivity to pyrimidine nucleotide effectors, and
typical Michaelis-Menten kinetics in the carbamoylphosphate
and the aspartate saturation curves. The B. subtilis enzyme
represents a typical Class C ATCase. The native enzyme is a
trimeric protein with a molecular mass of 102 kDa,
consisting of three 33.5 kDa polypeptides (Brabson &
Switzer, 1975) (Fig. 6). Class C ATCases have been found in
other Gram-positive organisms, Streptococcus faecalis (Chang
et al., 1974) and Staphylococcus epidermidis (Kenny et al.,
1996), as well as in several Gram-negative organisms,
-
20
Stenotrophomonas maltophilia, Xanthomonas campestris, and
Lysobacter enzymogenes (Kenny et al., 1996).
Recently, several bacterial ATCases have been described
which do not at present appear to fit within any of the
three classes. Ishihara et al. (1992) cloned the ATCase
gene of Treponema denticola in their studies of antigenic
proteins from this organism. This sequence produces a 55
kDa protein with 33.8% homology to the ATCase of E. coli.
No pyrT-like sequence was found downstream of the gene, and
the enzyme was not inhibited by CTP. Unfortunately, no
holoenzyme size is reported, so it not possible to determine
if the enzyme is a homotrimer or if it contains additional
subunits.
Unusual ATCases have also been reported for Thermatoga
maritima and Thermus aquaticus (Van de Casteele et al.,
1994) . The Thermatoga clone in the study produced a
truncated gene product of approximately 33 kDa, however the
authors suggest that the actual polypeptide is closer in
size to that of T. denticola based on homology of the two
sequences. The gene of T. aquaticus produces a polypeptide
of about the same size as the E. coli or Bacillus catalytic
subunit. The holoenzyme, which is inhibited by UTP, has a
molecular mass of 480 kDa and contains both ATCase and
DHOase activities. This enzyme adds a curious twist to the
evolutionary puzzle when compared to the Class A bacterial
-
21
ATCases with their inactive, DHOase-like subunit and similar
holoenzyme sizes.
In the multienzymic proteins of higher organisms,
ATCase is not under allosteric control. Instead, CPSase is
the site of regulation, with UTP and CTP acting as
inhibitors and ATP as an activator (Jones, 1980) . The yeast
ura2 gene product of Saccharomyces has a predicted molecular
mass of 245 kDa (Souciet et al., 1989), while the CAD
polypeptide has a molecular mass of about 240 kDa (Kelly et
al., 1986). The CAD polypeptide associates as trimers and
hexamers (Coleman et al., 1977). Trimers form by an
association around the ATCase domain (Fig. 7), much like
that seen for bacterial ATCases, while the hexamers are due
to dimeric interactions between DHOase domains of adjacent
trimers (Carrey, 1993; Davidson et al., 1993). It has been
suggested that channeling of substrates is an advantage
conferred by multienzymic proteins (Lue & Kaplan, 1970).
This may occur in effect through a process of facilitated
diffusion between active sites that are close together in
space (Carrey, 1993). Evidence has also been found for
channeling in the yeast enzyme (Penverne & Herve, 1983;
Belkaid et al., 1987) .
In plants, ATCase was found to be a regulatory enzyme.
The ATCase of plants is noteworthy since it combines
regulatory behavior with a comparatively small molecular
mass, 104 kDa in wheat germ (Yon et al., 1982) and 128 kDa
-
22
Figure 7. Model of the domain structure of CAD. ATC =
aspartate transcarbamoylase domain; CPS = carbamoylphosphate
synthetase domains; DHO = dihydroorotase domain; and GLN =
glutaminase domain. Adapted from Carrey (1993) .
-
24
in mung bean (Achar et al., 1974). Yon et al. (1982) showed
that the wheat germ ATCase is also a trimer. Inhibition was
seen with ATP, CTP, and UTP, with the greatest inhibition of
activity by the addition of UMP (Achar et al., 1974). The
CPSase and ATCase enzymes of radish (Raphanus sativus),
spinach (Spinacia oleracea), soybean (Glycine max), and corn
(Zea mays) have been localized in the chloroplasts (Shibata
et al., 1986) .
In the protozoan Crithidia fasciculata, neither
inhibition nor activation of ATCase was seen in the presence
of pyrimidine ribonucleotides (Kidder et al., 1976).
Kurelec (1974) reports that the ATCase of parasitic
platyhelminths is not affected by CTP, but is activated by
ATP. In some invertebrates, evidence suggests that CPSase
and ATCase are separate polypeptides. The single CPSase of
the land snail Strophocheilus oblongus is localized in the
mitochondrion, while ATCase is found in the soluble fraction
of the cell (Tramell & Campbell, 1970). A similar finding
exists for other snails (Otala lactea and Helix aspersa),
earthworm (Lumbricus terrestris), and land planarian
(Bipalium kewense) (Tramell & Campbell, 1971). In some
parasitic platyhelminths, CPSase is apparently absent,
although ATCase is still present (Kurelec, 1972, 1973,
1974) . These organisms require arginine and pyrimidines and
probably derive their carbamoylphosphate from the arginine
-
25
deaminase pathway as is done in Lactobacillus (Hutson &
Downing, 1968).
Properties of reported ATCases are summarized in Tables
3-8. The following organisms have been found to lack
ATCase: Chlamydia psittaci (McClarty & Qin, 1993);
Mycoplasma mycoides (Mitchell & Finch, 1977); and
Trichomonas vaginalis (Heyworth et al., 1984).
Dihydroorotase.
DHOase catalyzes the reversible cyclization of
carbamoylaspartate to dihydroorotate. The enzyme from
Clostridium oroticum has been purified to homogeneity and
shown to be a zinc-metalloenzyme (Taylor et al., 1976;
Pettigrew et al., 1985a). It has also been suggested that
the mammalian dihydroorotase is a zinc-metalloenzyme
(Christopherson & Jones, 1980, Simmer et al., 1990£>) .
The DHOase enzyme in prokaryotes has been found to be a
homodimer, with subunit molecular masses in the range of 40-
50 kDa (Pettigrew et al., 1985b; Ogawa and Shimizu, 1995;
Schenk-Groninger et al., 1995). While all described
prokaryotic DHOases are active in the dimeric form,
Krungkrai et al. (1990) report that the protozoan enzyme,
which is also approximately 40 kDa, is active as a monomer.
The DHOase of a eukaryotic green alga, Chlorella, has an
apparent molecular mass of 88 kDa (Dunn et al. 1977), which
would be consistent with the prokaryotic enzymes if it is
-
26
Table 3. Properties of known bacterial Class A ATCases.
Vmax reported in nmol CAA min"1 (mg protein). KmAsp and KmCP
given in mM of the substrate. References denoted as: 1)
Barron, 1994; 2) Bergh & Evans, 1993; 3) Bethell & Jones,
1969; 4) Burns et al., 1997; 5) Hooshdaran, personal
communication; 6) Kenny et al., 1996; 7) Linscott, 1996; 8)
Masood & Venkitasubramanian, 1988; 9) Mendz et al., 1994;
10) O'Donovan, personal communication; 11) Schurr et al.,
1995; 12) Shepherdson & McPhail, 1993; 13) Shepherdson et
al., 1997; 14) Van de Casteele et al., 1994; 15) Van de
Casteele et al., 1997; 16) Vickrey, 1993; and 17) West,
1994.
-
27
Organism kDa V vraax KmABp Kmcp Ref.
Bacterial Class A ATCases
Acinetobacter calcoaceticus 480 6
Azomonas agilis -450 6
Azotobacter vinelandii -450 3,6
Brevundimonas diminuta -480 139 1.0 1.0 7
Comamonas acidovorans -500 0.6 1.0 7
C. testosteroni -500 1.0 0.7 7
Deinococcus radiophilus 500 6
Helicobacter pylori 23 11.6 0.6 4,9
Leucothrix mucor -450 6
Micrococcus luteus -480 1
Mycobacterium smegmatis 465 34 5,8
Paracoccus denitrificans -450 6
Pseudomonas aeruginosa 484 120 1.5 0.13 3,16
Ps. aureofaciens -480 1.3 1.0 7
Ps. fluorescens 464 40 1.1 0.08 2,3
Ps. mendocina -480 1.0 1.0 7
Ps. pseudoalcaligenes 73 2.7 0.29 17
Ps. putida 482 8 2.2 0.60 11
Ps. syringae 490 31 1.3 0.9 12, 7
Ps. stutzeri -480 1.0 1.0 7
Rhizobium meliloti -480 10
Synechocystis sp. -490 13
Thermus aquaticus 480 20 14,15
-
28
Table 4. Properties of known bacterial Class B ATCases. Vmax reported in nmol CAA min"
1 (mg protein)"1. [S]os for both substrates given in triM. References denoted as: 1) Bethell & Jones, 1969; 2) Jyssum, 1992; 3) Kenny et al., 1996; 4) Purcarea et al., 1994; 5) Purcarea et al., 1997; and 6) Wild et al., 1980.
Organism kDa V vmax Asp [S] o s
CP [S] o.s Ref.
Bacterial Class B ATCases
Aeromonas hydrophila 285 18 17 . 0 6
Citrobacter diversus 280 46 5.5 6
C. freundii 290 23 7.5 1,6
Enterobacter aerogenes 295 196 4.8 6
En. liquefaciens 295 23 17 .5 6
Erwinia carnegiana 275 38 2.6 6
Er. herbicola 315 37 2.9 6
Escherichia coli 310 53 5.0 1,6
Neisseria canis 6.7 3.4 2
N. caviae 6.2 1.8 2
N. elongata 5.7 1.8 2
N. gonorrhoeae 9.2 5.4 2
N. meningitidis Ml 295 30.1 8.1 2
Proteus vulgaris 310 9 33 .0 1,6
Pyrococcus abyssi 310 3.0 0.06 4,5
Rhodopseudomonas spheroides 1
Salmonel1 a typhimuriurn 300 51 7.0 6
Serratia marcescens 275 20 19 .5 1,6
Shigella flexneri 285 454 5 . 6 6
Vibrio natriegens 280 3
Yersinia enterocolitica 315 10 4.2 6
-
29
Table 5. Properties of known bacterial Class C ATCases. Vmax reported in nmol CAA min"
1 (mg protein)_1. Km^ and Km^ given in mM of the substrate. References denoted as: 1) Barron, 1994; 2) Bethell & Jones, 1969; 3) Brabson & Switzer, 1975; 4) Chang & Jones, 1974; 5) Ghim et al., 1994; 6) Kenny et al., 1996; 7) Linscott, 1996; and 8) O'Donovan & Shanley, 1995.
Organism kDa V v max ^^•Asp Ktricp Ref.
Bacterial Class C ATCases
Bacillus caldolyticus 10 5
B. subtilis 102 380 7.0 0 .11 2,3
Lactobacillus fermentum -100 1
Lysobacter enzymogenes 120 6
Shewanella putrefaciens -100 2.0 0.5 7,8
Sporosarcina ureae -100 2.5 1
Staphylococcus epidermidis -100 6
Stenotrophomonas maltophilia 112 0.7 0.7 6,7,8
Streptococcus faecalis 128 4
Xanthomonas campestris 126 6
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30
Table 6. Properties of known bacterial ATCases - other, class not yet described. Vmax reported in nmol CAA min"
1 (mg protein) Km, LAsp and Kmcp given in mM of the substrate. References denoted as: 1) Ahonkhai et al., 1989; 2) Currier & Wolk, 1978; 3) Elagoz et al., 1996; 4) Grogan & Gunsalus, 1993; 5) Hutson & Downing, 1968; 6) Ishihara et al., 1992; 7) Li & West, 1995; 8) Linscott, 1996; 9) Makoff & Radford, 1978; 10) Norberg et al., 1973; 11) 0'Donovan & Shanley, 1995; 12) Van de Casteele et al., 1994; 13) Wheeler, 1989; and 14) Wheeler, 1990.
Organism kDa V vraax KmAsp Kiticp Ref.
Bacterial ATCases - Other, Class Not Yet Described
Anabaena variabilis 3 1.0 0.70 2
Burkholderia cepacia -600 74 2.5 2.2 7,8,11
B. pickettii -500 30 3.5 0.9 8
Halobacterium cutirubrum 160 15.0 2.7 9,10
Lactobacillus leichmannii 1.4 30.0 5,9
L. plantarum 3
Mycobacterium avium 0.8 14
M. leprae 13
M. microti 1.2 14
Pseudomonas indigofera -400 9.0 0.7 8, 11
Sulfolobus acidocaldarius .15 4
Thermatoga maritima 400 12
Treponema denticola 55a 6
Vibrio costicola » ̂ i « , — _ i - i
130 1
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31
Table 7. Properties of known eukaryotic ATCases - plant and
other monofunctional. Vmax reported in nmol CAA min"1 (mg
protein)"1. Km^ and Km^ given in mM of the substrate.
References denoted as: 1) Achar et al., 1974; 2) Asai et
al., 1983; 3) Doremus, 1986; 4) Dunn, 1977; 5) Grayson &
Yon, 1979; 6) Hammond & Gutteridge, 1980; 7) Holland et al.,
1983; 8) Kidder et al., 1976; 9) Kurelec, 1974; 10)
Landstein et al., 1996; 11) Lovatt et al., 1979; 12)
Mukherjee et al., 1988; 13) Nasr et al., 1994; 14) Neumann &
Jones, 1964; 15) Ong & Jackson, 1972; 16) Overduin et al.,
1993; 17) Shibata et al., 1986; 18) Tampitag & 0'Sullivan,
1986; 19) Tramell & Campbell, 1970; 20) Williamson & Slocum,
1994; 21) Yon, 1972; and 22) Yon et al., 1982.
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32
Organism kDa V "max Kir̂ Ref.
Eukaryotic ATCases - Plant and other monofur ictional
Arabidopsis thaliana 13
Babesia rodhaini 8 1.9 0.01 7
Chlorella Virus PBCV-1 200 10
Chlorella sorokiniana 160 4
Crithidia fasciculata 100 320 5.0 0.5 8
Cr. luciliae 150 21 2.6 0.03 18
Cucurbita pepo 11
Fasciola hepatica 9
Glycine max 17
Lactuca sativa 14
Leishmania donovani 135 35 7.6 0.3 12
Lycopers icon esculentum 16
Moniezia benedeni 9
Paramphistomum cervi 9
Phaseolus aureus 128 150 4.0 0.09 1,15
Pi sum sativum 110 16 3,20
Raphanus sativus 17
Spinacia oleracea 17
Strophocheilus oblongus 19
Toxoplasma gondii 140 17 17.6 0.03 2
Triticum vulgare 104 4 0.6 0.01 5,21,22
Trypanosoma cruzi 22 6
Vinca rosea 17
Zea mays 17
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33
Table 8. Properties of known eukaryotic ATCases -multifunctional. Vmax reported in nmol CAA min"
1 (mg protein)"1. KmAsp and Km^ given in mM of the substrate. References denoted as: 1) Caroline, 1969; 2) Christopherson & Jones, 1980; 3) Faure et al., 1989; 4) Freund & Jarry, 1987; 5) Hirsch, 1968; 6) Hong et al., 1995; 7) Hoogenraad & Lee, 1974; 8) Jarry, 1976; 9) Kent et al., 1975; 10) Kim et al., 1992; 11) Koskimies et al., 1971; 12) LaCroute, 1968; 13) Lollier et al., 1995; 14) Mally et al., 1980; 15) Penverne & Herve, 1983; and 16) Soderholm et al., 1975.
Organism kDa V vmax KmAsp KmCP Ref.
Eukaryotic ATCases - Multifunctional
Chicken 900 11
Coprinus radiatus 800 5
Dictyostelium discoideum 3
Drosophila melanogaster 800 5.4 0.44 4,8,16
Neurospora crassa 46 1
Hamster 243a 240 44
9.05 .004 .02
2,10,14
Mouse 900 0.7 2,11
Rana catesbeiana 900 30 9
Rat 900 10 7, 11
Saccharomyces cerevisiae 600 16.6 1.18 12, 15
Schizosaccharomyces pombe 13
Squalus acanthias 6
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34
also a dimer. The catalytically-active yeast DHOase is
similar to those described previously for prokaryotes, while
this activity is found within the CAD multienzyme for higher
eukaryotes.
Dendograms of DHOases show that they fall into two
distinct groups, with the multifunctional hamster,
Dictyostelium, Drosophila, and yeast inactive domain forming
one group while the monofunctional proteins from yeast, E.
coli, and Salmonella typhimurium constitute a second class
(Simmer et al., 1990Jb) . Even so, a 44 kDa proteolytic
fragment of the CAD protein, which forms a dimer, has been
shown to catalyze only the dihydroorotase reaction (Simmer
et al., 1990jb) . Comparison of this domain with well
characterized prokaryotic dihydroorotases indicated that
many features of the common ancestral protein had been
conserved (Kelly et al., 1986).
More recent analysis has shown that additional DHOase
sequences confirm the division into two diverged types
(Shepherdson et al., 1997). Active DHOases from T.
aquaticus, Synechocystis species, and P. aeruginosa align
with the E. coli group described above. This group is
designated the family C type DHOases by Shepherdson et al.
(1997) and generally has a molecular mass of 40 kDa. The
other group, designated family C', includes the CAD and CAD-
like DHOases as shown by Simmer et al. (1990jb) as well as
those from two archaeas, Methanococcus jannaschii and
-
35
Sulfolobus solfataricus. The family C' also contains the
DHOase of Lactobacillus leichmanni and the DHOase-like PyrC'
polypeptides of P. putida and Synechocystis sp., hence the
name of the family.
The existence of divergent families of DHOase adds to
the debate on the evolution of the CAD polypeptide. The
discovery of bacterial complexes containing ATCase and
either active or inactive DHOase subunits suggested that
these complexes could represent a CAD precursor (0'Donovan &
Shanley, 1995). This theory is further supported by the
presence of both CAD and these bacterial proteins in the
same DHOase family.
Streptomyces.
Pyrimidine metabolism has not been studied extensively
in one important order, the Actinomycetes. This order
contains a wide range of genera and species which are
generally divided into eight broad families based on partial
sequencing of 16S rRNA. Of these families, only one, the
family Streptomycetaceae, does not contain diverse taxa
(Goodfellow, 1989). The members of this family are
morphologically complex and have the ability to form spores
in or on the mycelium. The family contains a number of
genera, one of the most widely known being Streptomyces.
The genus Streptomyces was proposed in 1943 by Waksman
& Henrici for aerobic, spore-forming actinomycetes which had
-
36
been previously termed Actinomyces. The streptomycetes are
found within the high G+C Gram-positive subdivision based on
rRNA homology (Woese, 1987). They are most notable for
their large genome size, 10.5 x 106bp {Benigni et al.,
1975), their DNA base composition of 69 to 78 mol % guanine
(G) plus cytosine (C) (Korn-Wendisch & Kutzner, 1992), and
the wide range of antibiotics, vitamins, enzymes and enzyme
inhibitors they have been found to produce (Goodfellow &
Cross, 1983). As a result of these useful secondary
metabolites, thousands of strains have been isolated from
soils and sediments around the world. In addition,
Streptomyces also stands out as one of only a few
prokaryotes in which a linear chromosome is present
(Hinnebusch & Tilly, 1993; Lin et al., 1993; Lezhava et al.,
1995) .
Streptomyces and related genera occupy a unique
evolutionary position, being filamentous, spore-forming
prokaryotes. The spores are hyphal in origin, developing as
a result of septation and fragmentation of pre-existing
hyphal elements. These spores are, at maturity, relatively
unspecialized compartments of hyphae. They are bounded by
walls of true peptidoglycan which are normally about 30-50
nm thick, compared to 10-20 nm for vegetative cells (Locci &
Sharpies, 1983).
Following germination of the spore, the first type of
tissue to develop is the substrate mycelium. This fungus-
-
37
like growth is a branching network of multinucleate hyphae,
which are occasionally interrupted by septa (Chater, 1993).
Substrate mycelial growth continues until, presumably, the
nutrient supply is exhausted. At such a time,
differentiation takes place.
Aerial mycelia, upon which the spores will develop,
begin to grow on the colony. The aerial hyphae form a dense
lawn and grow perpendicular to the substrate mycelium.
These hyphae escape the aqueous environment of the colony
surface in order to grow into the air. This gives a fuzzy
white appearance to the surface of the colony. During this
stage, lysis occurs in the substrate mycelium, releasing
nutrients which can be utilized by the aerial mycelium for
growth (Mendez et al., 1985). The aerial mycelium then
undergoes septation into uninucleate units, which develop
into chains of gray-pigmented, hydrophobic spores. The
aerial mycelium is a device for the dispersal of the spores
(Kaiser & Losick, 1993). The use of mutants and molecular
genetics has begun to reveal how differentiation is brought
about in the multicellular, mycelial Streptomyces spp.
(Chater, 1993).
Classification within the genus has been complicated by
the proliferation of named species. The number of species
names that have been used for a streptomycete in the
scientific and patent literature rose from 41 in 1920 to 153
in 1957 to now more than 3,000. This explosion in species
-
38
number is certainly due in part to the many isolates
investigated in the course of screening for antibiotics and
other useful industrial compounds. This resulted in the
discovery of thousands of different secondary metabolites,
each one suggesting the existence of a distinct producer or
"special type", which were often considered as a "new
species" (Korn-Wendisch & Kutzner, 1992).
Initially, classification was based purely on
morphological features. This led, however, to many new
genera and families being created, as well as many species
and genera being rearranged. Overall, a very complex
situation was created. Chemical composition of the cell
wall provided a tool for distinguishing streptomycetes from
other actinomycetes. Lechevalier and colleagues (Becker et
al., 1964, 1965; Lechevalier & Lechevalier, 1970)
demonstrated that Streptomyces and other genera of the
family Streptomycetaceae contained LL-diaminopimelic acid
(LL-A2pm) in its peptidoglycan. Most other actinomycetes
described thus far contain meso-A2pm. Streptomycete
peptidoglycan is also characterized by an interpeptide
bridge composed of a glycine residue (Schleifer & Kandler,
1972) . A number of other biochemical criteria have been
used to examine Streptomyces. The Streptomycetaceae form a
homogeneous family in regard to these, including the pattern
of sugars in whole-cell hydrolysates (Lechevalier &
Lechevalier, 1970), phospholipids (Lechevalier et al.,
-
39
1977), fatty acids (Kroppenstedt, 1985), menaquinones
(Alderson et al., 1985; Kroppenstedt, 1985), and acetylated
muramic acid residues (Uchida & Aida, 1977).
Williams et al. (1983) studied 475 strains, which
included 394 type cultures of genus Streptomyces and
representatives of 14 other actinomycete genera. Overall
similarities of these strains for 139 unit characters were
determined. The results of this study provided a basis for
the reduction of the large number of Streptomyces species
which have been described. It demonstrated that the
previous use of a limited number of subjectively chosen
characters to define species-groups or species had resulted
in artificial classifications. Their data, together with
those from previous diverse studies, indicated that the
genera Actinopycnidium, Actinosporangium, Chainia,
Elytrosporangium, Kitasatoa, and Microellobosporia should be
reduced to synonyms of Streptomyces. The status of
Streptomyces clusters defined by Williams et al. have been
supported by chemical (Saddler et al., 1987; Manchester et
al., 1990), serological (Ridell & Williams, 1983), and
nucleic acid sequencing methods (Mordarski et al., 1986;
Witt & Stackebrandt, 1990; Labeda & Lyons, 1991; Labeda,
1992) .
Species classification within the genus has been
attempted using a number of characters. Streptomyces phage-
typing has shown that the host range pattern of a set of
-
40
polyvalent phages does not cross genus-boundaries and can
also be used in taxonomic studies for identification at the
species level (Korn-Wendisch & Schneider, 1992).
The application of polymerase chain reaction (PCR) to
obtain rDNA sequences has dramatically extended the
potential to use these sequences to elucidate natural
relationships between organisms (Stackebrandt et al., 1992).
In the genus Streptomyces, Stackebrandt and colleagues
observed that the 16S rRNA of too many species contains too
similar sequences that the analysis of the complete
sequences seems to be "unpractical and unnecessary".
Instead, restricting analysis to two regions of the
sequence, one for species-specific signatures and another
for intragenic classification, appears to be sufficient for
the purposes of streptomycete classification (Stackebrandt
et al., 1992; Kim et al., 1993). Stackebrandt et al. also
note that, given the limitations of highly similar 16S rRNA
sequences, analysis of this molecule cannot be hailed as the
solution to all problems of streptomycete taxonomy.
Analysis of rRNA is only one piece on the jigsaw puzzle that
eventually may lead to a more complete picture about
streptomycete evolution (Stackebrandt et al., 1992).
Analysis of ATCases could be another.
While much work has been done to identify the medically
and industrially important secondary metabolites produced by
the members of this genus, there are many basic questions of
-
41
primary metabolism which have yet to be answered. These
studies are not only necessary to help further understand
the events leading to and controlling secondary metabolite
production, but may also be useful as additional taxonomic
markers in the classification of species within the genus.
As a basic and universal pathway, the pyrimidine
biosynthetic pathway provides an opportunity for such study.
Enzymes of the pathway in other organisms have been used as
taxonomic and evolutionary markers with success (Neumann &
Jones, 1964; Bethell & Jones, 1969; Wild et al., 1980;
Foltermann et al., 1981; Beck et al., 1989; Major et al.,
1989; Tricot et al., 1989; Wild & Wales, 1990; Jyssum, 1992;
Kern et al., 1992; Kimsey & Kaiser, 1992; Nagy et al., 1992;
Davidson et al., 1993; Schofield, 1993; van den Hoff et al.,
1995; Kenny et al., 1996; Lawson et al., 1996; Linscott,
1996; 0'Donovan & Shanley, 1996).
A representative species, Streptomyces griseus, was
chosen for study. S. griseus was one of the original
antibiotic-producing organisms discovered by Waksman. The
genetically well-characterized species S. coelicolor was
also used in these experiments as several pyrimidine mutants
were available.
Genetic mapping studies of S. coelicolor A3(2) have
identified four uracil-requiring strains which have been
kindly supplied for this study by Dr. D. A. Hopwood. One of
these loci, designated uraA, maps in a separate location
-
42
from the others. However, the remaining three, uraB, uraC,
and uraD, are found in close proximity of each other and are
likely placed in the operon discovered in this research
(Hopwood & Kieser, 1990)(Fig. 8). The enzymes encoded by
these loci have not been identified, though the uraD strain
requires both uracil and arginine, suggesting that this is
the gene for CPSase. Since these mutations are found in the
same region of the chromosome, they could represent
mutations in different genes of an operon or within
different domains of a multienzymatic polypeptide.
In this research, I sought to determine the nature of
the enzyme ATCase, its structure and regulation, in
Streptomyces. These properties as well as the relationship,
both enzymatically and at the genetic level, of ATCase to
other enzymes of the pyrimidine pathway are important in
examining the classification of this enzyme relative to
other known ATCases. This study may also serve as a basis
for determining the type and distribution of ATCases within
the genus and in relation to other genera in the high G+C
Gram-positive subdivision.
-
43
Figure 8. Genetic map of S. coelicolor A3(2) showing
relative positions of markers conferring uracil requirement.
Adapted from Hopwood & Kieser (1990).
uraB,C,D
uraA
-
METHODS
Bacterial strains.
The wild-type strain Streptomyces griseus (ATCC 10137)
and S. lividans (ATCC 19844) were obtained from the American
Type Culture Collection (ATCC). S. griseus LU2 was selected
from the wild-type for 5-fluorouracil (5FU) resistance. S.
coelicolor M145 was kindly provided by Dr. D. A. Hopwood,
John Innes Institute, Norwich, England, while S. venezuelae
was obtained from Ward's Biological Supply. Table 9
includes a list of the strains utilized and their
genotype/phenotype.
Media and growth conditions.
Streptomyces spore suspensions were obtained from
growth on sporulation agar plates (Hopwood et al., 1985) or
from nutrient agar slants. Sporulation agar contained
1 g l"1 yeast extract, 1 g l"1 beef extract, 2 g l"1 tryptose,
trace FeS04, 10 g l"1 glucose, and 15 g l"1 agar in ddH20.
Difco nutrient agar contained 3 g l"1 beef extract, 5 g l"1
peptone, and 15 g l"1 agar in H20.
Two different defined media were used to grow the
Streptomyces strains. The solid medium used was
Streptomyces minimal agar medium (Hopwood, 1967), modified
for alternate carbon source. This medium contained 2 g l"1
44
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45
Table 9. Bacterial strains
Organism Source Genotype and/or Phenotype
S. griseus 10137 ATCC wild-type
S. griseus LU2 This study 5FU resistant
S. coelicolor M145 Hopwood wild-type
S. lividans 19844 ATCC wild-type
S. venezuelae Ward's Biological wild-type
B. cepacia 25416 ATCC wild-type
E. coli K12 Bachmann, B. wild-type
E. coli TB2 Roof, W. D. pyrB argF requires uracil and arginine
P. aeruginosa PAOl Holloway, B. W. wild-type
(NH4)2S04, 0.5 g l'1 K2HP04, 0.2 g l'
1 MgS04'7H20, O.Olgl"1
FeS04'7H20, and 15 g l"1 agar in ddH20. Succinate, pH7.0, was
added after autoclaving to a final concentration of 20 TOM.
Growth in liquid medium was done in Streptomyces minimal
liquid medium (Hopwood et al., 1985), which contained 2 g l"1
(NH4)2S04, 5 g l"1 Difco Casamino acids, 0.6 g I"1 MgS04'7H20,
and 50 g l"1 polyethyleneglycol (PEG) 8000 in 800 ml ddH20.
To this, 1ml minor elements solution (per liter: lg
ZnS047H20, lg FeS04'7H20, lg MnCl24H20, and lg CaCl2,
anhydrous) was added. After autoclaving, added 150 ml final
volume of 0.1M NaH2P04/K2HP04 buffer, pH6.8, and carbon
source, succinate, pH 7.0, to 20 mM final concentration.
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46
Required growth factors were added at the time of
inoculation.
For isolation of total DNA, Streptomyces cultures were
grown in nutrient broth with 34% (w/v) sucrose. Difco
nutrient broth contained 3 g l"1 beef extract and 5 g l"1
peptone in water to which 340 g l"1 sucrose was added.
Streptomyces cultures were grown at 30 °C. Liquid
cultures were shaken vigorously on an orbital shaker, in a
baffled flask when available.
Streptomyces spore suspension.
Spores were harvested from plate cultures of
Streptomyces according to a modified version of the
procedure of Hopwood et al. (1985). All solutions and
apparati were sterilized by autoclaving prior to use.
Water, typically 9 ml, was added to a well-sporulating plate
(2-3 ml for slant cultures). The agar surface was scraped
with a sterile loop to suspend the spores. The liquid was
then removed with a pipette and transferred to a test tube.
The suspension was mixed vigorously on a vortex mixer for 1-
2 minutes (min), then filtered by vacuum through fiberglass
wool to remove mycelial fragments (Fig. 8). The filtrate
was centrifuged for 10 min at 1800 x g in a Sorvall H1000B
rotor. The supernatant was immediately poured off. After
resuspending the pellet in the remaining drop (approximately
0.5ml), lml of water was added, mixed well, and the
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47
Figure 9. Apparatus used for preparation of spore
suspensions and removal of mycelial fragments. Modified
from the procedure of Hopwood et al. (1985) .
-
48
Crude spore
suspension
Fiberglass wool plug
Vacuum^
Filtered spore
suspension
125 ml Filter flask
-
49
mixture transferred to a screw-top vial containing 0.5ml
80% (w/v) glycerol to achieve a final concentration of 20%
glycerol. The suspension was mixed and stored at -20°C.
Selection of 5-fluorouracil resistant strain.
The 5-fluoropyrimidine analogues are toxic only after
being converted by the corresponding pyrimidine salvage
enzymes to the nucleotide level (0'Donovan & Neuhard, 1970).
Positive selection of spontaneous mutants utilizing these
analogues, in a manner similar to that of Martinussen &
Hammer (1995) , was used to obtain strains lacking specific
salvage enzymes. Selection of a 5FU resistant strain of S.
griseus was done by spreading a spore suspension of the
wild-type strain onto a minimal plate. To the center of the
plate, a crystal of 5FU was placed. Growth of the plate was
monitored. A zone of clearing indicated that the compound
was toxic. Colonies which arose within the zone were
allowed to sporulate and then collected using a sterile
cotton swab. These spores were transferred to a fresh plate
of minimal medium and streaked for isolation. Several
isolated colonies were then selected and transferred to
separate plates. Spores from colonies on these plates were
collected for spore suspensions. The spores were again
exposed to the analogue. Strains which showed no zone of
clearing were resistant to 5FU.
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50
Harvesting of bacterial cultures.
Cells for assays were typically prepared by inoculating
a flask containing 100 ml of Streptomyces minimal liquid
medium with 50-100 jul of a dense spore suspension of the
required Streptomyces strain. After 3 days incubation, 5 or
10 ml of this starter culture were transferred to one or two
liter volumes of the same medium. After 40 to 48 hours (h),
the cells were harvested by centrifugation in a Sorvall RC5C
centrifuge in a GS-3 rotor at 10,000 xg for 20 min at 4 °C
with the centrifuge brake turned off. Most of the
supernatant was carefully poured off to avoid disturbing the
loose pellet. The pellet was resuspended in the remaining
amount (10-15 ml) . The cells from 500 ml of the culture
were transferred to 50ml, disposable conical tubes. The
centrifuge tubes were washed with 10 ml of distilled water
to remove residual cells and the wash added to the cells in
the conical tubes. The conical tubes were then centrifuged
in a Sorvall H1000B rotor for 15 min at 1800 x g at 4 °C. The
supernatant was poured off. The cell pellet was either
frozen at -20 °C for storage or used immediately.
Cells for total DNA isolation were harvested by vacuum
filtration using a modified version of the method of Hopwood
et al. (1985). Cells were first grown in nutrient broth
with 34% (w/v) sucrose. The mycelium was then collected by
filtration in a Buchner funnel containing two sheets of
Whatman No. 50 filter paper. The mycelium was washed with
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51
100 ml of 10% (v/v) glycerol per 500 ml of culture, and the
paste then transferred to a sterile conical tube using a
sterile spatula. The cell paste was stored at -20 °C or
used immediately.
Preparation of cell extracts.
Cell extract was prepared by breaking the cells using
sonication. Cell pellets from freshly harvested cells or
frozen samples were resuspended in the appropriate breaking
buffer for the assays to be performed. Typically, 1ml of
ATCase breaking buffer (2 mM /3-mercaptoethanol, 20 ptM ZnS04,
and 50 mM Tris-HCl, pH8.0) was added per 1 g of wet weight
of pellet. The cell suspension was sonicated using a
Branson Cell Disruptor 200 for 5 min intervals while the
tube was in an ethanol-ice water slurry to control heating
of the sample. The sample was mixed by inversion, and the
sonication repeated for a total of 15 min sonication per
sample. The sonicated suspension was transferred to an
SA600 centrifuge tube and centrifuged at 4 °C for 1 h at
33,000 xg. The resulting supernatant was transferred to a
sterile 15 ml, disposable conical tube and stored at room
temperature until the cell extract was used in an assay.
Assays on crude extract were performed within 48 h of cell
extract preparation.
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52
Aspartate transcarbamoylase assay.
Cell extract was prepared as described earlier using
ATCase breaking buffer.
ATCase activity was assayed by measuring the amount of
carbamoylaspartate (CAA) produced in 20 min at 30 °C
according to the method of Gerhart & Pardee (1962), with
modifications using the color development procedure of
Prescott & Jones (1969). The optimal pH for ATCase in S.
griseus was first determined by assaying ATCase activity
with pH varied from 6.5 to 10.0. All the subsequent assays
were done at the optimal pH.
It was also necessary to determine the best storage
conditions for the S. griseus extracts. During initial
experiments, it was found that the extracts rapidly lost
activity during storage at 4°C. This was similar to the
cold-inactivation of the wheat-germ ATCase (Grayson, 1979).
A cell extract was prepared and then divided into 0.5ml
aliquots. An initial assay was done on the sample to
establish a zero-point for comparison. An aliquot was
stored under each of the following conditions: 1) room
temperature; 2) filtered through 0.2 fjM membrane and stored
at room temperature; 3) heated at 65°C for 5 min.,
centrifuged 2 min. at 10, 000 x g, and stored at room
temperature; 4) stored at 4 °C; 5) stored at 37 °C; 6) with 1
mM UTP and stored at room temperature; 7) in 20% glycerol
and stored at 4 °C; and 8) in 20% glycerol and stored at
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53
-20 °C. ATCase assays were performed on these samples after
storage for 1 day, 2 days, 7 days and 14 days.
Assays were performed to determine the Km for
aspartate
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54
ATCase assay tubes were prepared in advance, without
addition of carbamoylphosphate, and preincubated at 30 C
for 4 min. The reaction was initiated by the addition of
100 pi of carbamoylphosphate (Smgml"1) for the aspartate
curve and, for the carbamoylphosphate curve, by 100 /il of
carbamoylphosphate of varying concentrations (0.081 mM - 5.2
mM) . At 20 min, 1 ml of the color mix was pipetted into the
reaction tubes. The color mix contained 2 parts of 5 mg ml"1
antipyrine in 50% (v/v) sulfuric acid and 1 part of 8 mg ml"1
2,3-butanedione monoxime in 5% (v/v) acetic acid. Color was
developed by incubating the tubes in a 65 °C water bath with
the tubes exposed to the light of the room and capped by
marbles to limit evaporation. After 2 h of incubation, the
absorbance was read in a Beckman DU-40 spectrophotometer at
a wavelength of 466 nm. Controls were utilized to determine
background readings in tubes containing all reactants except
cell extract and in tubes with cell extract but lacking the
substrate carbamoylphosphate. All readings were blanked to
these controls.
The nmol CAA was determined using a CAA standard curve.
The standard curve was prepared using known concentrations
of CAA ranging from 50 to 500 nmol in the standard assay
reaction mix and under the same color development
conditions.
The kinetic curves were generated by plotting specific
activity of the enzyme (nmol CAA min"1 (mg protein)"1) versus
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55
the concentration of aspartate or carbamoylphosphate. The
Km, and Kmcp were determined from the intercepts of the
x-axis on Lineweaver-Burk plots (l/velocity versus
1/substrate).
When checking fractionation samples for ATCase
activity, the volume of sample used was typically 100 /xl,
with the water adjusted accordingly to achieve a final
reaction volume of 1ml. Aspartate concentration was 10 mM
for all such samples and the reaction time was 20 min. Stop
tubes were typically incubated for 2 h at 65 °C.
Repression of ATCase in cells grown in the presence of
exogenous pyrimidine compounds (100 pig ml"1) was also studied.
ATCase activity was measured as previously described using
varied aspartate and the specific activity of ATCase in
cells grown with pyrimidine compounds was compared to the
specific activity of cells grown in minimal medium without
any additions.
Dihydroorotase assay.
With an equilibrium constant of 1.0, the DHOase
reaction can be assayed in either direction. Here, DHOase
was assayed in the degradative direction. The DHOase assay
reaction tubes contained in a 1 ml total volume: 1 mM EDTA;
20 jtil fraction sample; 100 mM Tris, pH8.6; and ddH20 to
which 100 pi of 20 mM dihydroorotate (DHO) in 0.1 M phosphate
buffer, pH 7.5, were added to start the reaction. A
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56
background control was prepared for each cell extract used
which received cell extract, but no DHO. A blank was
prepared for each assay which contained all reactants, but
no enzyme. All readings were zeroed using these controls
before calculations were performed. After a 20 min
incubation at 30 °C, 1 ml of color mix (same as used for
ATCase assay) was added directly to the assay tubes. These
tubes were incubated at 65 °C for 2 h and then the absorbance
read at 466 nm. Conditions for storage of the enzyme were
determined in the same manner as was done for Streptomyces
ATCase. Kinetic studies were performed using varying
amounts of the substrate. A blank was prepared for each
concentration of DHO used, and all readings were zeroed to
the blanks prior to calculations.
Growth Curve.
A growth curve was prepared for S. griseus 10137. A
flask containing 2 liters of Streptomyces minimal liquid
medium was inoculated with 20 ml of a starter culture. At
the time of inoculation, a zero point sample was taken. Two
50 ml aliguots were removed and placed in pre-weighed, 50 ml
conical tubes. One was used for dry weight determination
and the other for ATCase and DHOase assays. Samples were
taken at 8 h intervals throughout the growth of the culture
up to 72 h. All samples were stored at 4 °C until collected
by centrifugation at 1800 x g. Pellets were stored at
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57
-20 °C. Samples for dry weight determination were dried
together in a vacuum desiccator for 120 h. ATCase and
DHOase assays were performed on cell extracts from each time
point.
Protein concentration determination.
Protein was quantified by the method of Lowry et al.
(1951) . Bovine serum albumin (BSA) at 0-100 fig was used to
produce a standard curve. The BSA was divided into 11 tubes
at 10 fig increments as follows: the 0 fig tube contained
only 200 fil ddH20, while the 10 fig tube contained 190 fil ddH20
+ 10 fil 0.1% BSA (100 fil of 1% desiccated BSA in water
diluted in 900 fi 1 ddH20 yields a solution of BSA at 1 fig
ixl'1) . The rest of the tubes used in the standard curve
were set up accordingly. Three different volumes (5 fil, 10
fil, and 20 fj.1) of the unknown sample cell extracts,
typically 1:10 dilutions in ddH20 of each sample, were added
and brought up to 200 fil total volume with ddH20. To the
standard and unknown tubes, 800 fil alkaline copper reagent
(0.5ml 2% sodium-potassium tartrate and 0.5 ml 1% copper
sulfate mixed together before adding 49 ml of 2% sodium
carbonate in 0.1 N sodium hydroxide) were added. The tubes
were allowed to stand at room temperature for 10 min. Folin
reagent (100 fil, 2 N commercial preparation diluted 1:1 with
ddH20) was added to all tubes while mixing on a vortex
mixer. After incubating at room temperature for 30 min, the
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58
absorbance was read at 660 nm. The BSA concentration was
plotted against the A660 to establish a standard curve.
Purification strategies.
Several purification strategies were utilized with
Streptomyces aspartate transcarbamoylase (ATCase) and
dihydroorotase (DHO). These strategies are outlined below,
with the specific procedures discussed in following
sections. All steps of each strategy were performed at room
temperature.
The first strategy was begun by streptomycin sulfate
precipitation of cell extract. The supernatant was then
fractionated using ammonium sulfate. The resuspended pellet
of the fraction containing maximal ATCase activity was
passed over a Sephacryl S-400 size-exclusion column. The
column fractions which contained maximal ATCase activity
were pooled and concentrated. The concentrate was then
fractionated using high performance liquid chromatography
(HPLC) ion-exchange.
In an alternate strategy, the extract was first passed
over a Sephadex G-200 size-exclusion column. The fractions
containing peak activity were pooled and concentrated. The
concentrate was separated through a Sephacryl S-400 size-
exclusion column. The tubes containing maximal activity
were pooled and then fractionated with ammonium sulfate.
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59
Streptomycin sulfate precipitation.
A streptomycin sulfate solution was made by mixing 10%
(w/v) streptomycin sulfate in a 10 mM phosphate buffer, pH
7.5 in the manner of Adair & Jones (1972). A one-half
volume of this 10% streptomycin sulfate solution was slowly
added to a known volume of cell extract (typically 30 ml)
while stirring slowly. After stirring for 1 h, precipitates
were removed by centrifugation for 30 min at 33,000 x g at
4 °C. A 0.5 ml sample of the supernatant was removed for
later assays and protein analysis. The remainder of the
supernatant was transferred to sterile conical tubes where
it was measured before proceeding to the next purification
step. Although Adair & Jones (1972) dialyzed their samples
before proceeding, excessive loss of activity was seen for
dialyzed Streptomyces extracts. Upon noting this in pilot
purifications of the samples, the dialysis step was omitted
in all subsequent experiments.
Ammonium sulfate fractionation.
Ammonium sulfate fractionation first involved
determination of the correct percent saturation for
precipitation of the desired protein. Depending on the
sample source, either streptomycin sulfate supernatant or
column fractions, the saturation for precipitation varied
due to differences in initial salt contents of these
samples. To determine the correct percent saturation for
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60
precipitation, a sample was fractionated in 5% (w/v)
ammonium sulfate saturation increments beginning at 15%
saturation at 25 °C. The pellet at each step was
resuspended in ATCase buffer, typically 400 fil, while 500 (il
of the supernatant was saved in a sterile 1.5 ml
microcentrifuge tube. Activity for ATCase and DHOase was
monitored in the pellet and supernatant. In this manner, it
was determined that maximal ATCase activity for Streptomyces
from streptomycin sulfate supernatants precipitated between
40% and 55% ammonium sulfate saturation at 25 °C while
ATCase activity from column fractions precipitated at 45%
ammonium sulfate saturation at 25 °C.
In the purification experiments using streptomycin
sulfate precipitation, the initial ammonium sulfate cut was
made by slowly adding ammonium sulfate to the streptomycin
sulfate supernatant until reaching the desired saturation.
The sample was stirred for 1 h. Precipitates were removed
by centrifugation for 30 min at 33,000 xg at 4 °C. The
supernatant was transferred to a sterile conical tube and
the volume noted. A 0.5ml supernatant sample was removed
and the pellet was resuspended in 0.5-1 ml ATCase buffer.
Both were stored for ATCase and protein analysis.
Additional ammonium sulfate was slowly added to the
supernatant to bring to the second saturation percentage.
The sample was stirred for 1 h before centrifugation for 30
min at 33,000 xg to remove precipitates. A 0.5ml
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61
supernatant sample was removed and stored for ATCase and
protein analysis. The pellet was resuspended in 0.5-1 ml
ATCase buffer and stored at room temperature.
When using ammonium sulfate precipitation of S-400
column fractions, only one cut was done. Ammonium sulfate
was slowly added to the combined column fractions until
reaching the desired saturation. The sample was stirred for
1 h. Precipitates were removed by centrifugation for 30 min
at 33,000 xg at 4 °C. The supernatant was transferred to a
sterile conical tube. The pellet was resuspended in 0.25ml
ATCase buffer and stored at room temperature.
Sephadex 6-200 column chromatography.
A 1 cm x 30 cm column was filled to a bed height of
23.5 cm with Sephadex G-200 size-exclusion matrix. Size-
exclusion buffer (1 mM EDTA, 20 jiM zinc acetate, 0.5 mM
/3-mercaptoethanol, 150 mM potassium acetate and 10 mM Tris-
HC1, pH 8.2, in ddH20) was degassed by vacuum and used as
the elution buffer. The column was loaded with 1 ml of
extract which was mixed with 50 fil of Blue Dextran (0.01 g
ml"1) and 2 /xl of Phenol Red (0.01 g ml"1) . After the entire
sample was in the gel matrix, the buffer reservoir was
connected, and collection of 1 ml fractions began. For all
determinations, the Blue Dextran peak tube was designated
Tube #1 and all other tubes were labelled relative to this
tube. Samples were stored at 25 °C.
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62
Each fraction was assayed for both ATCase and DHOase
activity. The assays were performed as described later in
this paper.
Sephacryl S-400 column chromatography.
A 3 cm x 100 cm BioRad column was gravity-packed with a
Sephacryl S-400 size-exclusion matrix. The same size-
exclusion buffer was used as with the G-200 column. The
column was first loaded with 0.5ml of blue dextran (0.01 g
ml"1}. The loading apparatus was washed with 0.5ml of
column buffer prior to loading 1-3 ml of sample. A gravity
flow system was used, and flow rate was typically 0.8ml
min"1. The void volume of typically 150 ml was removed and
discarded, after which 80 tubes of 5 ml fractions were
collected. In a few experiments, 2.5ml fractions were
collected in the tubes where ATCase was expected to elute.
Samples were stored at 25 °C.
Each fraction was assayed for both ATCase and DHOase
activity. These assays were performed as described later in
this paper.
Concentration of size-exclusion fractions.
The fractions from the size-exclusion chromatography
containing maximal ATCase activity were pooled and
concentrated using a Centriprep-30 concentrator (Amicon,
Inc., Beverly, MA). The concentrators were centrifuged at
1500 x g for 15 min at 25°C. The filtrate was decanted and
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63
centrifugation repeated according to the instructions until
the retentate was concentrated to approximately 3 ml in
preparation for use on an S-400 column and 0.7 ml in
preparation for HPLC fractionation.
High performance liquid chromatography ion-exchange.
The concentrated sample was loaded on a Waters Protein-
Pak Q 8HR AP Mini (5 mm x 50 mm) ion-exchange column. The
initial buffer (Buffer A) was 20 mM Tris-HCl, pH8.2, which
had been filtered and degassed. The flow rate was 1 ml
min"1. A linear gradient was applied over 40 min to go from
100% Buffer A to a 25%:75% mixture with Buffer B, which was
20 mM Tris-HCl, pH8.2 and 1 M KC1. During the gradient
change, 0.5ml fractions were collected. Samples were
stored at 25 °C. Each fraction was assayed for both ATCase
and DHOase activity. The ATCase and DHOase assays for
fractionation samples were used as described later in this
paper.
Nondenaturing polyacrylamide activity gels.
Cell extract was prepared as described previously for
the ATCase activity assay. Samples of typically 24 [j.1 of
cell extract were mixed with 6 jul of 5X loading buffer
(312.5 mM Tris, pH 6.8, 50% v/v glycerol, and 0.05% w/v
bromophenol blue in ddH20) . The total volume of 30 (xl was
loaded onto a nondenaturing polyacrylamide gel with a 5%
stacking gel and an 8% separating gel. The gel was
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64
typically run in a Bio-Rad Mini-Protean II cell at 100V for
2.5 h at room temperature. The gel was prepared by first
pouring the separating gel, which contained 2.67ml of the
stock solution of acrylamide (29% w/v acrylamide and 1% w/v
bis-acrylamide in ddH20) , 2.5 ml of Buffer B (1.5 M
Tris-HCl, pH 8.8), and 4.83 ml of ddH20. Ammonium persulfate
(0.02 g) was added to the mixture to remove dissolved
oxygen. Just prior to pouring the gel, 5 /z 1 of N,N,N',N'-
tetramethylethylenediamine (TEMED) was added and mixed in by
gentle inversion. The solution was poured into the gel
plates leaving a 2 cm gap at the top. The separating gel
was covered with a layer of N-butanol to prevent drying and
allowed to stand at room temperature for 1 h to polymerize.
The butanol layer was poured off, and the stacking gel
poured. The stacking gel contained 0.67ml of the stock
solution of 30% acrylamide, 1ml of Buffer C (0.5 M Tris, pH
6.8), and 2.3 ml of ddH20. Ammonium persulfate (0.01 g) and
TEMED (5 /xl) were added. Once the stacking gel was poured,
a comb was inserted between the plates to form the wells.
The stacking gel was allowed to polymerize for 30 min. The
running buffer contained 25 mM Tris and 192 mM glycine in
ddH20 at a pH of 8.8.
For molecular mass determinations, samples were loaded
on a 4-20% Bio-Rad polyacrylamide ready-gel. These gels
were run at 150V for 3 h at room temperature using the same
non-denaturing running buffer described above. Partially-
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65
purified ATCase from Escherichia coli K12 (holoenzyme and
catalytic trimer) and Pseudomonas aeruginosa PA01 with known
molecular mass were used as standards, as were Sigma protein
molecular weight markers urease and BSA.
The gels were stained specifically for ATCase activity
by a procedure developed by Bothwell (Bothwell, 1975) and
further modified by K. Kedzie (1987), who used histidine
instead of imidazole. In this study, two slight
modifications to the procedure were made as indicated. When
the electrophoresis was complete, the plates were separated,
and the gel was placed in 250 ml 50 mM histidine, pH7.0, at
room temperature (modified from ice-cold at this st