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

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

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.

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

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

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

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.

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

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.

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

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.

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

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

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

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

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

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

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.

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

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

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.

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

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

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-

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