DEOXYRIBONUCLEIC ACiD - DEPENDENT RIBONUCLEIC
ACE) ?’!33LYMERASE OF PSEUDOMONAS PUTIDA:
PURIFICATION AND CHARACTERIZATION

Thesis for the Degree of Ph. D.
MICHIGAN STATE UNIVERSITY
JAMES CARL JOHNSON
1971

    
  

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Michigan. State
University

 

 

This is to certify that the

thesis entitled
DEOXYRIBONUCLEIC ACID-DEPENDENT RIBONUCLEIC

ACID POLYMERASE 0F PSEUDOMONAS PUTIDA:
PURIFICATION AND CHARACTERIZATION

presented by

JAMES CARL JOHNSON

has been accepted towards fulfillment
of the requirements for

9‘ D degree 1n WO‘Z/

WW 460$)”;

Major professor

Date 'QM / lf7/
' II /

0-7639

 

ABSTRACT

DEOXYRIBONUCLEIC ACID-DEPENDENT RIBONUCLEIC
ACID POLYMERASE OF PSEUDOMONAS PUTIDA:
PURIFICATION AND CHARACTERIZATION

 

BY

James Carl Johnson

The objectives of this study were to obtain homo-

geneous DNA-dependent RNA polymerase from Pseudomonas

 

putida, to determine the size and structure of the enzyme,
and to study properties of the enzyme-mediated synthesis
of RNA.

Two forms of RNA polymerase (nucleoside triphos-
phate RNA nucleotidyltransferase, EC 2.7.7.6) from g.
putida were resolved by chromatography on phosphocellulose
and subsequently purified to greater than 98 per cent of
homogeneity. As determined by sodium dodecyl sulfate—
polyacrylamide gel electrophoresis, the polypeptide sub-
unit structures of the two forms of the enzyme were dZBB'o
and azBB'. The molecular weights of polypeptides a, 8, B',
and 0 were 44,000, 155,000, 165,000, and 98,000, respec-
tively. The sedimentation coefficients of 0288'0 and
0288' as determined by sucrose gradient centrifugation in

0.40 M potassium acetate were 13 S and 12 8, respectively,

 

James Carl Johnson

but in 0.05 M potassium acetate, the sedimentation coef-
ficients of the aggregate forms of aZBB'o and oZBB' were

19 S and 25 S. Similar results were obtained by analytical
ultracentrifugation. Polyacrylamide gel electrophoresis

of aZBB'o or of 0288' resulted in several bands of protein.
Each protein band was enzymatically active in the unprimed
synthesis of poly A-poly U, thereby suggesting that the
multiple bands represented aggregate forms of the enzyme.
35S—labeled B. putida RNA polymerase a288'o was

purified from labeled cells which had been grown on a I

 

minimal growth medium containing 35S-labeled sulfate. As

shown by sodium dodecyl sulfate-polyacrylamide gel electro-

phoresis the 35S-labeled-enzyme was at least 98 per cent

homogeneous. The specific radioactivity of the enzyme was

7 Cpm per mg of protein. Analysis of 358 content

of each polypeptide subunit showed that the amount of 35S

2.3 X 10

0f 8', B, and 0 relative to a was 3.6 to 3.6 to 2.2 to 1.0,
respectively. The amount of 8' plus 8 in the Initial
thract was 1.2 per cent of the total protein as determined
by sodium dodecyl sulfate polyacrylamide gel electrophor-
esis. From the amount of 8' plus 8 relative to the total
Protein and the formula weight of oZBB'o, it was calculated
that there were approximately 5,000 molecules of dzBB'o
per E. putida cell.

The reaction catalyzed by RNA polymerase was fol-
lowed by measuring the incorporation of 3H labeled-

ribonucleoside monOphosphates into RNA or by measuring the

James Carl Johnson

formation of inorganic pyrophosphate. An enzymic method
for the determination of inorganic pyrophosphate was
developed. Inorganic pyrophosphate was quantitatively
determined from the amount of NADPH formed via the action
of UDP—glucose pyrophosphorylase, phosphoglucomutase, and
glucose-6-phosphate dehydrogenase. The method for the
determination of inorganic pyrophosphate was used for the
assay of RNA polymerase by coupling the generation of
inorganic pyrophosphate by RNA polymerase with NADPH
formation. A mole of inorganic pyrophosphate released by
RNA polymerase resulted in the reduction of a mole of
NADP+. By means of the radioactive assay of RNA polymer-
ase or the assay for inorganic pyrophosphate, it was

shown that dZBB'o was 4- to 5-fold more active in tran-
scribing native bacteriophage gh-l DNA than was 0288'.
With denatured DNA or poly d(A-T) as template, aZBB' was
twice as active as aZBB'o. The rate of the DNA-directed
POlymerization reactions catalyzed by azBB'o in the
presence of pancreatic ribonuclease was increased 10 per

cent, whereas that rate catalyzed by ozBB' was increased

80 to 110 per cent.

 

DEOXYRIBONUCLEIC ACID-DEPENDENT RIBONUCLEIC

ACID POLYMERASE OF PSEUDOMONAS PUTIDA:

 

PURIFICATION AND CHARACTERIZATION

BY

James Carl Johnson

 

A THESIS

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY

Department of Microbiology and Public Health

1971

DEDICATION

To my parents who provided the
opportunity and encouragement
and to my wife who was blessed
with patience and understanding.

ii

 

ACKNOWLEDGEMENTS ,

My sincere thanks and appreciation for the help
and guidance in all phases of my graduate program are
expressed to Dr. John A. Boezi. His insistance on clarity a
and simplicity in the design of experiments and in writing
has been a great help to me. My thanks also go to

Dr. Harold L. Sadoff whose efforts through the Department

 

 

of Microbiology and Public Health enabled me to continue
the research program with Dr. Boezi.

Sincere appreciation is also expressed to the
other members of my Ph.D. guidance committee, Dr. Philipp
Gerhardt, Dr. Ralph N. Costilow, and Dr. Robert R. Brubaker.
I thank the other members of the Biochemistry and Micro-

biology and Public Health Departments with whom I have

been associated for advice and discussions. Thanks are

given to Gary Gerard, Bob Blakesley, Howard Towle, Kathy

Rose, and Drs. Lucy Lee, Robert Armstrong, Ken Payne, and

Seizen Toyama for discussions. I am indebted to

Mrs. Monique DeBacker for technical assistance.

I thank the Departments of Biochemistry and Micro-
biology and Public Health and the National Science

Foundation and the National Institutes of Health for

financial support.

iii

I especially thank the Department of Microbiology
and Public Health for presenting a Graduate Office

Scholarship to me in the spring of 1969.

iv

‘I-hjl" 'v

 

CURRICULUM VITAE OF JAMES CARL JOHNSON

May 1, 1971

Biographical Sketch

 

Name: James Carl Johnson

Born: October 2, 1942 in Madison, Wisconsin
Citizenship: United States of America
Marital Status: Married

Education

 

(1947-1959) Elementary and Secondary Education--pub1ic
schools in Madison, WisconSIh and Waterloo,

Iowa.

Participant in Northeast Iowa Science
Fairs (1957, 1958, 1959).

(1960-1964) Undergraduate Education--Iowa State
University, Ames, Iowa.

Ma'or: Biochemistry
National Science Foundation Undergraduate

Program participant (1961, 1962, 1963).
Title of Project: Purification and
Physical Characterization of Feather

Keratin.
Academic and Thesis Advisor: Dr. Malcolm

Rougvie.

Graduation Date: May, 1964.

(1964-1967) Graduate Educatigf—Department of Biochemistry,
East Lansing,

Michigan State University,
Michigan.

Master of Science Thesis Title: Purifica-
tion an C aracterization of DNA-
dependent RNA Polymerase from
Pseudomonas putida A. 3.12. .

Thesis Advisor: Dr. John A. Boezi.

Stipend: National Institutes of Health
Training Grant.

 

(1967-1971) Graduate Education--Department of Micro-
bioIbgy and Public Health, Michigan State
University, East Lansing, Michigan.

Doctoral Thesis Title: DNA-dependent
RNA Polymerase of Pseudomonas putida:
Purification and Characterization.

Thesis and Academic Advisors: Dr. John A.,
Boezi and Dr. Harold Sadoff.

Stipend: N.I.H. Teaching Grant

 

 

 

(1969) Received a Graduate Office Scholarship from
the Department of Microbiology and Public
Health.

Publications and Abstracts

1. Boezi, J. A., L. F. Lee, and J. C. Johnson,
"Characterization of Bacteriophage gh-l of
Pseudomonas fluorescens A 312," Bacteriological
Proceedings, 66, No. V27, 113 (1966).

 

2. Johnson, J. C., and J. A. Boezi, "RNA Polymerase
of Pseudomonas putida," Bacteriological Pro-
ceedings, 68, No. P115, 131 (1968).

3.1 Johnson, J. C., M. Shanoff, S. T. Bass, J. A.
Boezi and R. G. Hansen, "An Enzymic Method for .

Determination of Inorganic Pyrophosphate and Its
Use as an Assay for RNA Polymerase, Analytical

Biochemistry, 26, 137 (1968).

’ " Polymerase
4. Johnson J. C. and J. A. Beri, .RNA
from Pseudomonas putida," Federation Abstracts,
891 (1970).

. . M. DeBacker, and J. A. Boezi, .
J C ' Acid-dependent Ribonucleic ACid

domonas putida," Journal of
246 1222 (197II.

5. Johnson,
"Deoxyribonucleic
Polymerase of Pseu
Biological Chemistry;

5. Gerard, G. F., J. C. Johnson, J. A. Boezi, The

Release of o from Pseudomonas putida RN: Poly;
merase during the DNA-directed RNA Syn e512 9

' ' ss 1971.
Process," Federation Abstracts (in pre )

Egaching Experience

””196” —2r———-———————-£’*De W “first:grasses $3.225.
804 an .
Uggéigraduate Laboratory 059 superVised by

Dr. E. Benne. . .
400 Honors Laboratory superVised by Dr. F

Rottman.
vi

 

 

(1969) Department of Microbiology and Public Health.
Microbiology Undergraduate and Graduate
Laboratory 401 supervised by Dr. Ralph
Costilow.

 

vii

 

TABLE OF CONTENTS

GENERAL INTRODUCTION

LITERATURE SURVEY . . . . . . . .
Introduction . . . . . . . . . .
The Essential Role of RNA Polymerase in_vivo
Bacterial RNA Polymerase Structure . . . .
Protein Factors Affecting RNA Synthesis

Sigma I I I I I I I I I
Psi . .

 

Rho I I I I I I I I I I I I I
M-factor . . . . . . . . . . . .
References . . . . . . . . . . . .

ARTICLE 1

An Enzymic Method for Determination of Inorganic
Pyrophosphate and Its Use as an Assay for RNA
Polymerase, J. C. Johnson, M. Shanoff, S. T.
Bass, J. A. Boezi, and R. G. Hansen, Analytical
Biochemistry, ggJ 137 (1968). .

ARTICLE 2

Deoxyribonucleic Acid-Dependent Ribonucleic Acid
Polymerase of Pseudomonas putida, J. C. Johnson,
M. DeBacker, and J. A. Boezi, Journal of
Biological Chemistry, 246, 1222 (1971)

 

ARTICLE 3

35
Purification and Properties of S-labeled RNA
Polymerase of Pseudomonas putida, J. C. Johnson
and J. A. Boezi (manuscript in preparation).

viii

Page

24

30

42

 

GENERAL INTRODUCTION

The model for gene expression, developed nearly
fifteen years ago, states that genetic information encoded
in DNA in the form of deoxyribonucleotide sequences is
transcribed into ribonucleotide sequences of RNA. Trans-
scription is the process involving base pairing, whereby
the genetic information contained in DNA is used to order
a complementary sequence of bases in an RNA chain. Several
species of RNA molecules which are transcribed from a DNA
template have been described. These Species include
messenger RNA, ribosomal RNA, and transfer RNA. Each of
these species of RNA is specifically involved in the
synthesis of polypeptides. Ribosomal RNA is the structural
nucleic acid component of ribosomes. Transfer RNA is the
Species of RNA that is able to accept and covalently com-
bine with an amino acid and to hydrogen bond with a
messenger RNA nucleotide triplet. Only messenger RNA,
however, carries the information which specifies the
Primary structure of polypeptides. Translation of the
information encoded in messenger RNA to form the primary
structure of polypeptides is the second step in gene

exPression.

 

 

Transcription of DNA is the essential first step
in gene function. Considering the importance of gene
function in biology, one would like to know the details of
control and mechanism of processes required for transcrip-
tion. One approach to the study of transcription has been
to search for the activities which are required for
in yigrg synthesis of RNA from DNA templates in cell-free
extracts. This approach was adopted by several laborator-
ies nearly ten years ago and led to the discovery of an
activity which catalyzed the synthesis of RNA from a DNA
template.

The enzyme that catalyzes transcription of DNA is
RNA polymerase. It was first isolated from bacterial
extracts but has recently been isolated from extracts of
a variety of prokaryotic and eukaryotic cells. Genetic
experiments have been used to demonstrate that the RNA
polymerase prepared from bacteria is the major, if not
the only, enzymatic activity responsible for the synthesis
of messenger, ribosomal, and transfer RNA. Recently, an
effort has been directed toward the determination of the
subunit polypeptide structure of the purified enzyme.
This approach has led to the discovery of protein factors
which have the property of influencing the enzyme-mediated
reaction in vitro. Until now these studies have been
generally restricted to the RNA polymerase obtained from
the extensively studied bacterium, Escherchia_coli. Two

varieties of RNA polymerase have been prepared from E. coli

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These varieties differed in structure by a single polypep-
tide chain, which was called sigma. Sigma was found to

cause the RNA polymerase molecule to initiate synthesis at

correct sites on the DNA templates used for i§_yit£g
synthesis of RNA.

When work was begun for this thesis, only a partial
purification of the enzyme from E. ggli_had been reported.
It was decided that a second example of RNA polymerase
from bacterial sources should be provided for study.
Pseudomonas putida was utilized as the bacteria from which
RNA polymerase was purified beCause the bacteria was being
used in related studies of RNA structure and methylation.
In addition, a bacteriophage which was specific for
P, putida was isolated. It was of interest to determine
whether the host RNA polymerase would transcribe the bac-
teriophage DNA. The procedure described herein for the
purification of RNA polymerase resulted in two forms of
the enzyme which were nearly homogeneous. One of the
forms contained 100 per cent of the amount of sigma capable
of being associated with the enzyme. The other form of
RNA polymerase was completely devoid of sigma.

This thesis is organized into four major sections.
The first is a literature review in which much of the
information on bacterial RNA polymerase structure and
factors has been described. The second and third sections

are composed of articles describing the assay, purification

 

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and structure of the enzyme. These articles have been
included in the form of reprints from the journals in which
they were published. The fourth section consists of a
manuscript which is to be published. It concerns the puri-
fication and properties of 3SS-1abe1ed RNA polymerase from
P, putida and the use of the radioaCtive enzyme to inves-
tigate the release of sigma factor following the initia-

tion of RNA synthesis.

 

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LITERATURE SURVEY

Introduction

 

The first reports of the detection, isolation, and
purification of a DNA-dependent RNA polymerase from bac-
terial sources occurred in the early 1960's (1,2,3,4,5).
As of now, a prodigious literature concerning bacterial
RNA polymerase has accumulated. The extent and variety of
topics treated in the literature are beyond the scope of
this survey. The following reviews and symposia should be
consulted for a comprehensive treatment of the literature
(6,7,8,9,10,11).

RNA polymerase has been purified from the following
bacterial sources: Escherchia coli (several strains)

(1,2,5,12,13,14); Azotobacter vinelandii (3,15); Micrococcus

 

luteus formerly Micrococcus lysodeiktus (4); Bacillus
subtilis (16); and Pseudomonas indpgophera (17). The most
extensively purified enzymes, those obtained from the
various strains of E. ggli and the RNA polymerase from

A. vinelandii, are homogeneous as determined from sedimen-
tation and electrophoretic studies. E. 391$ RNA polymerase
has been the subject of most of the kinetic and structural

studies.

 

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6

The Essential Role of RNA Polymerase in Vivo

Evidence for the role of DNA-dependent RNA polymer-
ase was provided through a study of E. coli mutants having
alterations in a structural gene(s) for RNA polymerase which
rendered the cell temperature sensitive or resistant to the
rifamycin antibiotics. A mutant of E, 92;; which rendered
the cell temperature sensitive (growth at 30° but not at 42°)
was unable to incorporate labeled uridine into RNA at the
non-permissive temperature (18). RNA polymerase isolated
from the mutant grown at 30° was active when tested Ea IiEEE
at 30°, but was considerably less active relative to the
wild type RNA polymerase when tested EE_yEE£g_at 42°. Sub—
sequent analysis indicated that the E, ggEE_mutant had a
structurally altered RNA polymerase molecule. The nature
of the alteration has not been determined.

Rifamycin or its derivative rifampicin when added to
a growing culture of E, ggli_inhibited growth (19,20,21).
Wild type E, ggl£_cells were not able to incorporate labeled
uridine into RNA following addition of rifampicin to the
culture. It was shown that rifampicin inhibited the DNA-
directed synthesis of RNA by RNA polymerase E3 yi££9_(22,23).
Other studies determined that rifamPiCin binds to RNA poly-
merase and blocks initiation of RNA synthesis (24,25). Rifam-
picin resistant mutants of E, ggii_have been isolated which
synthesize an altered RNA polymerase molecule (21). Rifam_
picin did not bind to the altered enzyme. Consequently, the

antibiotic did not inhibit RNA synthesis i2.2££2,°r i2 vitro.

 

The studies on temperature sensitive and rifamycin
resistant mutants together with the extensive biochemical
studies on the RNA polymerase lead to the conclusion that
the RNA polymerase as isolated by the various purification
procedures is the "genetic transcriptase" (26), that is,
the essential enzyme responsible for most if not all RNA

synthesis from a DNA template in the bacterial cell.

Bacterial RNA Polymerase Structure

 

The subunit structure of purified E, 99EE_RNA
polymerase has been investigated. SDS or urea polyacryla-
mide gel electrophoresis of RNA polymerase which was
denatured by means of SDS or urea in the presence of
reducing agents resolved either three or four polypeptide
subunits (27,28). The RNA polymerase which had been
chromatographed on phosphocellulose contained three poly-
peptide subunits designated 8', B, and a, with molecular
weights 165,000, 155,000, and 39,000 respectively. The
RNA polymerase which was not purified by phosphocellulose
chromatography contained 8', B, a, and o polypeptide sub-
units. Sigma (0) has a molecular weight of 95,000 (28).
The molar ratio of the polypeptides obtained from the
phosphocellulose purified enzyme was 18' : 18 : 2a. The
minimal polypeptide subunit formula for this enzyme was
“288'. The RNA polymerase which was purified by gel
filtration and sucrose or glycerol gradient centrifugation,

but not by phosphocellulose chromatography contained

 

8', 8, a, and o in a molar ratio of 18' : 18 : 2a : lo.
The.minimal subunit formula for this enzyme was azBB'o.
The minimal formula weights calculated for a288' and 0288'0

5 and 4.93 X 105 respectively. These minimal

were 3.98 X 10
fOrmula weights correspond to the monomeric forms of the
enzyme.

In addition to the 8', 8, o, and a polypeptide
subunits of o288' and a288'o, a polypeptide (omega, w) of
molecular weight 9,000 was variably associated with the
purified enzyme (13,29). It is not known whether w is a
structural component of the enzyme or whether it has any
function in RNA synthesis.

Sephadex G-200 chromatography of denatured a288'
in the presence of SDS or urea resulted in the separation
of 8 plus 8' from a (27). The 8 plus 8' polypeptide sub-
units were separated on DEAE-cellulose in urea (27). The
molecular weights of the 8 plus 8' and the a subunits
determined by SDS-polyacrylamide gel electrophoresis were
found to be the same as those determined by sedimentation
equilibrium studies with the isolated subunits.

Amino acid compositions of a and of 8 plus 8' as
well as the a288' enzyme have been performed (27). Methio-
nine is the only N-terminal amino acid of either a, 8, or
8'.

Prior to the establishment of the polypeptide sub-
unit structure of RNA polymerase, studies of the sedimen-

tation properties of the enzyme were confusing. Two,

 

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three, or more species of molecules were observed in sedi-
mentation velocity experiments with highly purified RNA
polymerase (30,31). A partial understanding of the effects
of ionic strength, subunit composition, preferential
hydration, and irreversible dissociation has helped to
clarify much of the confusion. Berg and Chamberlin ana-
lyzed the two species of RNA polymerase, o288' and a288'o,
by sedimentation velocity and equilibrium experiments (29).

Apoenzyme (o288') in buffers of ionic strength above 0.26

0
20,w

dependence of sedimentation coefficient on the protein

behaved as a single sedimenting species (S =12.6). The

concentration was slight. When apoenzyme was centrifuged
in buffers with ionic strength below 0.26, a variety of
aggregated species was observed with average sedimentation
coefficients of 44 to 48 S. Holoenzyme (o288'o) in buffers

with ionic strength of 0.12 or higher sedimented as a

0
20,w

in low ionic strength buffers, the holoenzyme aggregated

0
20,w

a full complement of o sedimented as a mixture of apo- and

single species with S =15.0. When o288'o was sedimented

to a dimer with S =23.0. RNA polymerase with less than
holoenzyme at both low and high ionic strengths. The
molecular weights determined by sedimentation velocity
analysis were in agreement with those calculated from SDS-
Polyacrylwmide gel electrOphoresis studies. Molecular
weight determinations of 0288' and 0238'0 by sedimentation

equilibrium.have not been definitive.

 

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Sedimentation of E, SQli.RNA polymerase in increas-
ing concentrations of urea indicated that the 3288' form
could undergo large conformational changes or could dis-
sociate into smaller species (32). RNA polymerase dis-
sociates into its polypeptide subunits in high urea
concentrations (27,33). The 9 S species derived from RNA
polymerase in low urea concentrations may represent a
mixture of a8 and a8' (32). Renaturation of urea or
lithium chloride dissociated enzyme to obtain a partially
active enzyme which has the characteristics of the non-
dissociated enzyme has been described (34,35).

The polypeptide subunit structure of the E,
vinelandii and the E, subtilis RNA polymerase was found

to be similar to the E, coli polymerase. The E, vinelandii

 

RNA polymerase consists of an apoenzyme with polypeptide
subunit formula o288' and a holoenzyme a288'o (36). The
molecular weights of a, B, 8', and 0 were virtually iden-
tical to those from E, EEli.RNA polymerase. E, subtilis
RNA polymerase, although not totally purified, contained
Pelypeptide subunits of molecular weight 155,000, 120,000
57,000 and 45,000 (37). Two polypeptides at molecular
weight 155,000 were resolved. The molar ratio of the
155,000 dalton material to the 45,000 dalton material was
1:1. Chromatography of the enzyme on phOSphocellulose
removed the 57,000 dalton polypeptide from the enzyme and

resulted in an enzyme activity with altered template

 

11

specificities. The 57,000 dalton polypeptide is therefore
similar to E, ggii_RNA polymerase a factor in function but
not in size.

Infection of E, EQli.bY T4 bacteriophage results
in modification of the host RNA polymerase subunit struc-
ture. The a polypeptide chains are modified apparently by
the addition of 5' AMP in a T4 phage-specific reaction
(38,39). This modification occurs early in infection.
Adenylation of RNA polymerase from E, ggl£_by ATP with an
activity from uninfected cells has also been reported (40).
The adenylated enzyme is less active in RNA synthesis than
is the non-adenylated form. Both the 8 and 8' polypeptides

are modified late in T infection (39,41). The host 0

4

subunit is replaced by a T -specific 0 factor early in

4

infection (42,43). The T -specific 0 has the same molec-

4
ular weight as the host 0, and it confers different tem-
plate specificities to the apoenzyme. Late in T4 infection
this T4-specific 0 may be replaced by yet a different T4-
specific 0 (41).

Infection of E. ggli_with bacteriophage T7 results
in the Q2 ERIE synthesis of an entirely new type of RNA
polymerase molecule (44). The T7-specific RNA polymerase
is a product of gene 1 and is distinct from the host poly-
merase in that its activity is not inhibited by rifamycin,

streptolydigin, or by antibody specific for the host

enzyme. The enzyme consists of a single polypeptide

I_-—-_—v-‘r.4 ,V"... .. ' .'

 

 

12

subunit of molecular weight 110,000 as determined by SDS-
polyacrylamide gel electrophoresis.

Structural alteration of E. subtilis RNA polymerase
during sporulation has been reported (37). The sporula-
tion RNA polymerase has a polypeptide of 110,000 daltons
which is not found in the enzyme from vegetative cells. The
sporulation enzyme was found to contain only one of the
155,000 dalton polypeptides found in the vegetative RNA

polymerase.

Protein Factors Affecting RNA Synthesis

A series of sequential reactions serve to describe
the steps in RNA synthesis by RNA polymerase from a DNA
template. These steps in RNA synthesis are association,
initiation, elongation, termination, and dissociation.
During association, the RNA polymerase molecule interacts
with and binds to sites on the DNA template. Initiation
occurs with the formation of the first phosphodiester bond
between the 5'-termina1 and the second ribonucleotide.
Elongation follows with the sequential addition of ribo-
nucleoside monophosphates and the release of pyrophosphate.
At termination the synthesis of RNA ceases. Dissociation
follows termination with the release of the RNA product

and the RNA polymerase from the DNA template.

Sigga
Several proteins have been discovered which have

the property of affecting one or more of the steps in RNA

 

 

13

synthesis by bacterial RNA polymerase Eg'yiggg. One such
protein is the sigma (a) polypeptide subunit of E. 32;;
RNA polymerase (28). The E, ggli_sigma subunit has been
isolated from holoenzyme (a288'o) by chromatography on
phosphocellulose (28).

The efficiency by which a DNA template may be
transcribed by RNA polymerase is affected by the presence
or absence of o. The efficiency by which RNA polymerase
transcribes a particular DNA template £2.XiE£2 is measured
from the overall rate of RNA synthesis which reflects the
rate limiting step in either association, initiation,
elongation, termination or dissociation. The initiation
reaction may be followed by means of the exchange reaction
which occurs between inorganic pyrophosphate and the ribo-
nucleoside triphosphates involved in initiation (45).
Measurements of RNA synthesis and the pyrophOSphate
exchange reaction showed that transcription from E, ggli

bacteriophage T was dependent upon the presence of sigma

4
(46). It was concluded that c was probably necessary for
the formation of the first phosphodiester bond catalyzed
by RNA polymerase.

The asymmetry of RNA synthesis from native DNA is
affected by the presence of o in the reaction. RNA syn-
thesis $2.!i12 is asymmetric and must initiate and

terminate at specific sites on the DNA template. RNA

synthesis in vitro using holoenzyme is asymmetric on

14

E, 32;; bacteriophage T7 or A DNA and only the early phage
genes are transcribed (6,47). Also, E, ggli_holoenzyme
catalyzes asymmetric transcription of DNA from E. subtilis
bacteriophage SP01 (48), and animal viruses SV40 and adeno-
virus (49,50). In contrast, E, gg£i_apoenzyme catalyzes
symmetric transcription from a variety of templates (T4,

F1, and T bacteriophage DNA) (47,51,52). RNA synthesis

7
by apoenzyme is initiated at non-specific sites on the DNA
molecules. Therefore, it may be concluded that o affects
a specific interaction between the enzyme and functional
sites on the DNA. The nature of the functional sites
which specify holoenzyme recognition may be the same or
similar on a variety of DNA molecules from different
sources. The functional sites for holoenzyme initiation
may be equivalent to the promotor regions (53). Thus,
sigma may direct the initiation of specific RNA chains by
identifying specific DNA promotor regions to which the
enzyme may bind.

The effect of c on the binding of RNA polymerase
to DNA has been studied by a nitrocellulose filter binding
assay (54,55). It was found that 0 was not required for
formation of an RNA-polymerase-DNA complex. The stability
0f the complex was greatly enhanced, however, by the
presence of o. The formation of the DNA-RNA.polymerase
holoenzyme complex was temperature sensitive which suggests

that a cooperative reaction was involved (39). No more

 

15

than 7 or 8 molecules of holoenzyme could tightly bind to
T7 DNA. These data suggest that o is involved in the
formation of a highly stable enzyme-promotor complex which
may entail opening of the DNA helix in localized specific
regions. 1

From the above studies it may be hypothesized that

o is required for recognition of promotor regions on DNA,
or o is required for tight binding of the apoenzyme to the
DNA, or both. By the first hypothesis, the specific infor-
mation for site recognition is located in the 0 subunit.
By the second, the site recognition information is located
in the apoenzyme. Then, sigma would act as an allosteric
effector for tight, site-specific binding. Sigma, in the
absence of apoenzyme, will not bind to DNA (55).

At some stage of RNA synthesis by E. EQli and E,
yinelandii holoenzyme i2 y£E£2_o is apparently released
from the enzyme (46,52,56,57). This has led to the postu~
lation of the sigma cycle by Traverse and Burgess (56).
The essential feature of the sigma cycle is that once RNA
synthesis starts, sigma is released to bind to other apo—
enzyme molecules which may subsequently bind to DNA at
Specific promotor sites and initiate RNA synthesis.
Release of c has not been demonstrated EE_yEyg_and there
has been no direct demonstration of the release of o from
holoenzyme ig_vitro until now (58 and unpublished obser-

vations). The rationale for the release of sigma from

holoenzyme during RNA synthesis remains obscure.

16

The incubation of holoenzyme with a variety of
ribopolymers in the absence of nucleoside triphosphates
results in the apparent release of o (36). This suggests
that if c release occurs Eg.yiyg_it may be mediated by
RNA. However, much of the RNA synthesized E2_yiyg is
bound to either ribosomes in the case of messenger RNA
(57) or protein in the case of ribosomal RNA (60) prior to
release from the DNA-enzyme-RNA complex and, thus, the

RNA may be unavailable for mediation of c release E3 vivo.

iii

A class of proteins which act as positive control
elements in transcription have been isolated. These pro-
teins which act as secondary specificity determinants have
been named psi (W) (61). One such psi factor which is
Specific for the ribosomal RNA genes (Tr) has been parti—

ally purified from E, coli (61). In the presence 0f Tr,

but not in its absence, E. coli RNA polymerase holoenzyme

catalyzes the synthesis of ribosomal RNA 33 vitro using

E. coli DNA as the template. The specificity of transcrip-

tion by apoenzyme is not affected by Wr-

W activity has been found to be associated with
r

purified QB replicase (61). The RNA replicating enzyme

purified from bacteriOphage QB infected E. coli contains

a Phagespecific polypeptide chain of 69,000, and three

hOSt specific polypeptides of molecular weights 33,000,

 

.-. .l

.w

Avv‘.
u

1"

‘4

~A

0"

 

 

)-

 

17

47,000, and 74,000 (62,63). The Tr activity is associated
with the two small host specific polypeptides.

Another psi-like factor which has positive control
over the lac operon is CAP (64). CAP is a protein which
requires bound cyclic AMP for its activity. The cyclic
AMP-CAP complex is necessary for the transcription of the

8-ga1actosidase genes by holoenzyme £2 vitro (65).

Rho

RNA chain termination and release may, in part, be
mediated by a protein isolated and extensively purified
from E. EQEE_ribosome-free extracts. This protein has
been named rho (p) (66). The protein is probably composed
of four subunits each with a molecular weight of 50,000 as
determined by SDS-polyacrylamide gel electrophoresis and
sedimentation velocity experiments. Analysis of the size
of RNA transcription products formed by E, gel; holoenzyme
using A DNA as the template in the absence of added p
determined that most of the RNA products had sedimentation
coefficients which were greater than 22 S. In the presence
of added p, two classes of RNA having sedimentation coef—
ficients of 7 or 12 S were formed. The 7 and 12 S RNA
products made Eg.y£E£9 are the same size as the early A-
specific messenger RNA made iglyiyg. The 12 8 RNA molecule
is probably the messenger RNA of the N gene of A DNA. The
Protein product of the N gene is required for transcription

of the late 1 genes Ea vitro by E. coli holoenzyme (67).

 

18

Thus, it has been suggested that the N gene protein is an
anti-p factor, a protein which interferes with the activity
of p and permits DNA transcription into the late A genes

(67,68).

M-factor

Psi and rho proteins were obtained from the super—
natant fraction of E, 22E; cells. Another protein, M-
factor, has been purified from the ribosomal fraction of
the cell (69). Like sigma, M-factor is a protein which
has a sedimentation coefficient of 4 S and increases the
rate of RNA synthesis by E, 22l£.RNA polymerase holoenzyme
ig_yi££9, Mrfactor binds to purified E. Egli RNA poly-
merase holoenzyme. The available data suggest that M-
factor participates in RNA synthesis i2.!i££2.b¥ combining
with holoenzyme and affecting an event which occurs during

or close to initiation (70).

0

Al).

 

f.‘

10.

ll.

12.

13.
14.

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Rabussay, D., and Zillig, W., Fed. Exper. Biol. Soc.
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Hayashi, M., Hayashi, M. N., and Spiegelman, 8., Proc.
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Burgess, R. R., J. Biol. Chem., 244, 6168 (1969).

Burgess, R. R., Travers, A. A., Dunn, J. J., and
Bautz, E. K. P., Nature, 221, 43 (1969).

 

Berg, D. and Chamberlin, M., Biochemistry, E, 5055
(1970).

Richardson, J. P., Proc. Natl. Acad. Sci. U.S.A., E5,
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Smith, D. A., Martinez, A. M., Ratliff, R. L.,
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21

Ishihama, A., and Hurwitz, J., Federation Proc., EE,
659 (1969).

 

Walter, G., Seifert, W., and Zillig, W., Biochem.
Biophys. Resy_Commun., EE, 240 (1968).

 

Lill, U. I., and Hartmann, G. R., Biochem. Biophyg.
Res. Comm., 32, 930 (1970).

Sumper, M., Riepertinger, C., and Lynen, F., Fed.
Europe. Biochim. Soc. Letters, E, 45 (1969).

Krakow, J. S., and Von der Helm, K., Cold Spring
EarborlSymposia on Quantitative Biology, E2,
73 (1970).

 

Losick, R., Shorenstein, R. G., and Sonenshein, A. L.,
Nature, 227, 910 (1970).

 

Goff, C. G., and Weber, K., Cold Spring Harbor
Symposia on Quantitative Biology, 3;, 101 (1970).

Zillig, W., Zechel, K., Rabussay, D., Schachner, M.,
Sethi, V. 8., Palm, P., Heil, A., and Seifert, W.,
Cold Spring Harbor Symposia on Quantitative
Biology, 2;, 47 (1970).

Chelala, C. A., Hirschbein, L., and Torres, H. N.,
Proc. Natl. Acad. Sci. U.S.A., fig, 152 (1971).

Travers, A., Cold Epring Harbor Symposia on Quanti—
tative Biology, EE, 241 (1970).

Travers, A., Nature, 223, 1107 (1969).
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Chamberlin, M., McGrath, J., Waskell, L., Nature, 228,
227 (1970).

Krakow, J. S., J. Biol. Chem., 241, 1830 (1966).

Dunn, J. J., and Bautz, E. K. F., Biochem. Biophys.
Res. Commun., 3;, 925 (1969).

Summers, W. C., and Siegel, R. B., Natugg, 223, 1111
(1969).

Grau, 0., Ohlsson-Wilhelm, B. M., and Geiduschek, E. P

gold Sprin Harbor 8 mposia on Quantitative
Biology, 32, 221 (1970).

'I

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22

Westphal, H., and Kiehn, E. D., Ibid., p. 819.

Green, M., Parsons, J. T., Pina, M., Fujinaga, K.,
Caffier, H., and Londgraf—Leurs, I., Ibid., p. 803.

Sugiura, M., Okamato, T., and Takanami, M., Nature,
225, 598 (1970).

Bautz, E. K. F., Bautz, F. A., and Dunn, J. J.,
'Nature, 223, 1022 (1969).

Arditti, R. R., Scaife, J. G., Beckwith, J. R., J. Mol.
Biol., _E, 421 (1969).

Jones, 0. W., and Berg, P., J. Mol. Biol., EE, 199
(1966).

 

Hinkle, D. C., and Chamberlin, M., Cold Spring Harbor
Symposia on Quantitative Biology, E2, 65 (1970).

 

Travers, A. A., and Burgess, R. R., Nature, 22 , 537
(1969).

Darlix, J. L., Sentenac, A., Ruet, A., and Fromageot,
P., Europe. J. Biochem., ii, 43 (1969).

 

Gerard, G. F., Johnson, J. C., and Boezi, J. A., Fed.
Proceedings, 1971 (in press).

 

Miller, 0. L., Beatty, B., Hamkalo, B., and Thomas,
C. A., Cold Spring Harbor Symposia on Quantitative
Biology, EE, 505 (1970).

Mangiarotti, G., Apririon, D., Schlessinger, D., and
Silengo, L., Biochemistry, Z, 456 (1968).

 

Travers, A. A., Kamen, R. I., Schleif, R. F., Nature,
228, 748 (1970).

Kamen, R., Nature, 228, 527 (1970).

 

Kondo, M., Galleroni, R., and Weissmann, C., Nature,
228, 525 (1970).

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Acad. Sci. U.S.A., EE, 104 (1970).

Eron, L., Arditti, R., Zubay, G., Connaway, S., and
Beckwith, J. R., Proc. Natl. Acad. Sci. U.S.A.,
_6_8_, 215 (1971) .

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67.

68.

69.

70.

 

23

Roberts, J., Cold Sprin Harbor S m osia on Quantita-
tive Biology, EE, lgI (I970).

Schwartz, M., Virology, 32, 23 (1970).

 

 

 

Davidson, J., Pilarski, L. M., and Echols, H., Proc.
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Davidson, J., Brookman, R., Pilarski, L., and Echols,
H., Cold Spring Harbor Symposia on Quantitative

Biology, EE, 95 (1970):

ARTICLE 1

AN ENZYMIC METHOD FOR DETERMINATION OF INORGANIC

PYROPHOSPHATE AND ITS USE AS AN ASSAY FOR RNA

POLYMERASE

BY

J. C. Johnson, M. Shanoff, S. T. Bass,

J. A. Boezi, and R. G. Hansen

Reprinted from Egalytical BiochemistEy, 26, 137 (1968)

24

  

.A '1 fl!" . .
431146": '

    

 

osphafe and Its Use as an Assay for
RNA Polymerase . ‘

  
  
  
  
  
  
 
  
  
  
 
  
   
   
 

:l " IN, MIKE SHANOFF, S. T. BASS, J. A. BOEZI,
‘3 AND R. G. HANSEN

-. 0! Biochemistry and Microbiology, Michigan State University,
East Lansing, Michigan 48823

Received February 13, 1968

v "'0 pyrophosphate (PP,) is a product of many biosynthetic
;; In biopolymer formation PP, arises (a) in protein synthesis
; lingo of amino acid activation, (b) in nucleic acid synthesis at

aversion in the formation of the glycosyl donor, and (d) in lipid
,: at the stage of fatty acid activation.

usual method of PP, analysm involves its hydrolysis to inorganic
hosphate (P,) followed by P, determination by the Fiske and
:i-w method (1). Both inorganic and organic phosphate com-
3v“ n n: interfere in this analysis.

‘iklbrecht, Bass, Seifert, and Hansen (2) reported the purification and
._ * ; ’ation of UDP-glucose pyrophosphorylase from calf liver. The
thility of this crystalline enzyme has made possible the develop-
M of a specific enzymic method for the determination of PP,. Using
‘_‘.}hsphoglucomutase and glucose-G-phosphate dehydrogenase—both of
W are commercially available, and UDP-glucose pyrophosphorylase,
m can be quantitatively determined from the amount of NADPH
. bread in the following series of reactions:

o,

 
 
    
   
 
 

    
   
 

Exfozgmrylase
PP: + UDP-glucose ——p————' UTP + glucose-LP
phosphoglucomutue
‘ , glucose-LP ———————~ gluoose-fi-P
’7 amt...
(km? + NADP ————-——-—-+ gluconolactone-G—P + N ADPH
. .ble assay conditions, one equivalent of NADPH will be
" fi- 'ofPPp
-vi.e., coupling the determination of PP, with the forms.

     
  
 

  
   
 

            

    

 

 

138 JOHNSON, SHANOFF, BASS, BOEZI, AND HANSEN

tion of NADPH via the three enzyme-catalyzed reactions, can also he
used as an assay for enzymes which generate PP, as a reaction product.
A description of the enzymic method for the determination of PP, and
the application of the method for the assay of DNA-dependent RNA
polymerase is the subject of this report.

MATERIALS AND METHODS

NADP, UDP-glucose, glucose-6—phosphate dehydrogenase, and the 5’-
phosphate derivatives of adenosine, guanosine, uridine, and cytidine were
obtained from P-L Biochemicals, Inc. Calf thymus DNA, pancreatic
RNase (type I-A) , and phosphoglucomutase were purchased from Sigma
Chemical Co. DNase I (electrophoretically purified) and inorganic
pyrophosphatasc were obtained from Worthington Biochemical Corp.
[‘H]-ATP and [sHl-CTP were purchased from Schwarz BioResearch,
Inc. Actinomycin and nogalamycin (3) were gifts from Merck, Sharp,
& Dohme, and The Upjohn Company, respectively.

UDP-glucose pyrophosphorylase was isolated from calf liver by the
procedure of Albrecht, Bass, Seifert, and Hansen (2) and recrystallized
twice. RNA polymerase was purified from Pseudomonas putido A312
by a procedure that will be the subject of another communication (4).
For the experiments reported here, ammonium sulfate fraction II (A.S.II)
of RNA polymerase was used. No inorganic pyrophosphatase activity
has been detected in this fraction. Using 100 [Lg/ml P. putida bacte-
riophage gh-l DNA as template, A.S.II had a specific activity of 2500—
3000 mnmoles CTP converted into a trichloroacetic acid insoluble form per
hour per milligram protein (radioactive assay). In the spectrophotometric
assay, A.S.II had a specific activity of 5000—5500 mnmoles NADPH
formed per hour per milligram protein.

P. putida bacteriophage gh-l DNA (5, 6) was purified by the method
described by Thomas and Abelson (7). The concentrations of both dl-I
DNA and calf thymus DNA were determined from their ultraviolet ab-
sorptions using the extinction coefficient Emil" = 200. Calf thymus DNA
was denatured by heating at 100° for 10 min, followed by quick 000131188

For the determination of PP, by the UDP-glucose pyrophosphorylaie
assay system, the reaction mixture contained, in 0.5 ml: 50 mole We:
acetate buffer (pH 8.0), 1 nmole magnesium acetate, 0.2 mole
0.2 ”mole UDP-glucose, excess phosphoglucomutase and g1 u -..-. 7
phate dehydrogenase, from 5 to 20 mnmoles PP,, and errands
glucose pyrophosphorylase so that reaction was complete in "
NADPH formation was measured at 340 111,, in a : «hp.

3 equipped With a Gilford BW‘
.3; g; _ .'

éaES—s Eu} gar!-

 

...
_ a

   
 
 
 
 
 
 
 
 
 
 
 
 
 
 

     
 
   
   
 
    
 
 
    
   

  

 

     
  
 
  
   
  
   
 
   
   
  
   
 
 
  
 
 
 
 
 
 
 
 
  
  
  
 

‘ 3 mo moo r011 PP, DETERMINATION 139

   

" '. = using the molar extinction coefficient of 6.22 X 10’ (9) . "' ";
mixture employed for the formation of RNA com-
“NA by RNA polymerase contained, in 0. 5 ml. 10 moles
7 ; .'.' uer (pH 8.0), 1 mole magnesium acetate, 0.25 mole
V' “site, 0.2 mole each of ATP, GTP, CTP, and UTP, 0.1
', 0.1 mole UDP-glucose, excess phosphoglucomutase,
hate dehydrogenase, and UDP- glucose pyrophosphoryl-
a u 1 DNA, and 4 to 17 ,ug RNA polymerase. Before initiating
: .. with RNA polymerase, a 20 min incubation was required
’, reaction of PP, contaminating the nucleoside triphosphates.
formation was measured at 340 my in the spectrophotometric
‘ the radioactive assay, [3H] -CTP (5 X 108 cpm/nmole)o
(3 X 10‘ cpm/nmole) was used. Material, insoluble in 10%
*I v I cetic acid, was collected on nitrocellulose membrane filters
' ed for radioactivity in a liquid scintillation spectrometer.
l'eaction mixture for polyriboadenylate (poly A) synthesis by
7., . 1 nerase contained, in 0. 5 ml: 10 nmoles Tris acetate (pH 8 0),
magnesium acetate, 0.25 nmole manganese acetate, 0.2 mole
,1le mole NADP, 0.1 nmole UDP-glucose, excess phosphoglu-
j. , glucose-6-phosphate dehydrogenase, and UDP- glucose pyro-
' lacs, 50 pg heat-denatured calf thymus DNA, and 70 pg RNA
= w For the radioactive assay, [3H]-ATP (1 X 101 cpm/gmole)

RESULTS
.1. {nation of Inorganic Pyrophosphate by the Pyrophosphorylase
System. The results presented in Table 1 demonstrate the equiv- - 1 .
between PP, added to the pyrophosphorylase assay system and L 'r‘ 7,;
H formed. For the three experiments reported, using from 5 to i ’ -~‘_r
«I‘Iles of PP, per assay mixture, an equivalent amount of NADPH
armed. An equivalence between .PP, and NADPH was also found
TP,," 5'-mono-, di-, or triphosphate derivatives of adenosine or
31 ., a mixture of the four nucleoside triphosphates, RNA polymerase,

 
 
      
   
 
 

  

TABLE 1
.... tion of Inorganic Pyrophosphate by Pyrophosphorylase Assay System

PP: added. mole- NADPH formed, moles
Expt. 1! Expt. [[1 Exm. I Expt. II Expt. 1n

 
 

 

 

 

 
     
   

 

    

         
       

5.2 5.6 4.9 5.0 5_3
10.0 10.0 10.1 10.0 10.4
.0 15.1 14.8 15.2 16.0
.0 20.2 19.5 21.1 20_o

      

 

 

    

JOHNSON, sumo”, BASS, BOEZI, AND HANSEN

 

 

TABLE 2
Efiect of Various Agents on Determination of Inorganic Pymphouphatl

Additions to pyrophosphorylase NADPH.
assay system mole-

15 mumoles PP, 15.0—15.5
+150 mnmoles P, 15.5
+15,000 mpmoles P, 14.8
+200 mpmoles AMP 15.0
+200 mumoles ADP 16.2
+200 mnmoles ATP 15.9
+200 mpmoles UMP 14.5
+200 mpmoles UDP 15.3
+200 mumoles UTP 14.5
+200 mpmoles each of ATP, GTP, UTP, and CTP 14.6
+35 ug RNA polymerase 10.1
+250 ug denatured calf thymus DNA 16,1

 

or denatured calf thymus DNA were added to the assay mixture (Table
2). In other experiments, it was shown that actinomycin or nogalamycin
do not interfere in PP, determination.

Spectrophotometric Assay for RNA Polymerase. The requirements for
the reduction of NADP by the pyrophosphorylase assay system can-
comitant with the generation of PP, by RNA polymerase were detarv
mined. The results are given in Tables 3 and 4. If RNA polymerase, gh-l
DNA, the four nucleoside triphosphates, the purine nucleoside triphos-
phates, or the pyrimidine nucleoside triphosphates were omitted from the
reaction mixture, no NADPH was formed (Table 3). As shown in
Table 4, when DNase, inorganic pyrophosphatase, actinomycin or
nogalamycin was added to the reaction mixture, little or no NADPH ml
formed. However, when RNase was added to the reaction mixture, the

formation of NADPH was stimulated. The stimulatory efiect of RNase .

on PP, formation by RNA polymerase has been reported by Krakoi
(10) and Maitra and Hurwitz (11).

In Figure 1 the relationship between the rate of NADPH format!”
TABLE 3 "

Requirements for NADPH Formation Concomitant with Synthesis of REA:
’ ‘ Compomuol "’
motion mixture

 

NADPH.
mole/min/ml

 

.00

 

 

  

‘ 1 men 103 PP, DWINATION

 
 
 
    
 
  
   
  
 
   
  
  
   
  
   
   
  

 
 
 
 

TABLE 4
of Various Agents on NADPH Formation .‘
Concomitant with Synthesis of RNA . . . .

 

N ADPH.
mumolee/min/Inl

 

O
3;
i
5'
E
cook-o.—
asses
cl

 

van t of added RNA polymerase is given. Under the assay

employed, the relationship was linear to the formation of 2

NADPH/min/ml.

1 amount of RNA synthesized is proportional to the amount

polymerase used (12, 13). Likewise, the total amount of

formed was proportional to the amount of RNA polymerase

the assay system. For example, when 8.5, 17, 25.5, and 34 ug/ml

;~ .1; ::e were used, 19, 34, 50, and 64 mpmoles/ml NADPH ‘7
..,~. - , respectively.

kinetics of NADPH formation as measured in the spectrophoto-

.leeay and RNA synthesis as measured by the conversion of [3H]

Md ['H]-ATP into a trichloroacetic acid insoluble form are given

- 2. Following a short lag, the rate of NADPH formation was ,
l‘es/min/ml. The rates of incorporation of UMP and AMP were I ' ' _‘ 5
0.0 mpmoles/min/ml, respectively. At the plateau, approximately . }
lee/ml of NADPH was present, and 16 and 9 mamoles/ml of .
and AMP had been incorporated into RNA. - aft“

 

‘6'
I

!°
0
1

b

 

 

   

NADPH (mpmoles/min/ml)

 

. 1

IO 20 30
RNA POLYMERASE (Mg/ml)

~ '1' between rate of NADPH formation and mount of 11m

    
   

 

    

   
   
    
 
   
 
 
     
 
   

142 JOHNSON, SHANOFF, BASS, BOEZI, AND HANSEN

For comparison of the results of the two assay procedures, it should- 1
pointed out that the spectrophotometric assay measures the formation
of NADPH as a consequence of the production of PP, by RNA polymers-
ase. PP, is generated by RNA polymerase concomitant with the 111-»
corporation of each of the four nucleotides into the intrachain phoepho:
diester link of RNA. No distinction is made between the synthesis of
trichloroacetic acid soluble or insoluble polyribonucleotides. The radio
active assay measures the incorporation of one of the four nucleotidm
into polynucleotides which are insoluble in trichloroacetic acid. syn-
thesis of oligo- or polynucleotides which are soluble in trichloroacetic
acid is not detected.

sszaaiefii

 

 

 

 

O.

2 NADPH

< so - f

3:: .

m E

2 8

o '5 4O -

E

L

o :1.

I .5,

o_ _ CMP

9, 2° Men»: ..... -

2 017 ..-_._.§C—.————O-
1 1 1 1
20 4O 60 80

TIME, in minutes

FIG. 2. Time course of NADPH formation and CMP and AMP incorporation
into RNA.

The base composition of the RNA product has not been determined
and is not necessarily identical to that of gh-l DNA (mole % G + C =
57.0) which served as template (5). However, if it is assumed that the
base composition of RNA is the same as the gh- 1 DNA template, the
total nucleotides incorporated into the TCA insoluble RNA product may
be calculated from the amount of CMP and AMP incorporated. ‘
amounts to 56 mpmoles/ml from the CMP data and 41 m ... ‘
from the AMP data. The former estimate is in good agreement; the
is somewhat lower than the amount of NADPH found in the :‘a . ,
tometrie assay. However, the fact that the estimate of "
nucleotides incorporated into RNA 1s difierent when cal . ‘

 

  

     
    
 
     
  
  
   
  
 
   

if“!!! moo ma PP, DETERMINATION 143
tion concomitant with the generation of PP. during
u 'b'oadenylate (poly A). The kinetics of NADPH
l-AMP incorporation into poly A are presented in
. : rates of NADPH formation and AMP incorporation
- 'r, somewhat more NADPH was present near the end
indicating the synthesis of some trichloroacetic acid
i 01' the incorporation of UTP, produced in the UDP-

... .horylase reaction, into polymer. No NADPH was
RNA p01ymerase was omitted from the reaction mixture or
, .-.. calf thymus DNA was used in place of heat-denatured
51 DNA.

  

 
 
  
 

   
   
  
 
  
 
   
 
   
   
   

 

 

 

 

2 I00 - NADPH
E
\
n
2
2 so ' AMP "_o
e ,A/
v ’1’ O
0. __ I
2 60 /
<
5
I 40 -
(L
O
‘2‘ o
20 - I NADPH (minus RNA polymerase
I, or with unheated
I calf thymus DNA)
I _4—.-

 

 

so 120 130
TIME, in minutes

. . DISCUSSION

”amation of PP, via the action of UDP-glucose pyrophos-
"" PhOSPhOglucomutase, and glucose-6-phosphate dehydrogenase
' "' to be accurate, sensitive, and specific. The method is limited
the sensitivity of the quantitative determination of NADPn
" Ipecificity of the enzymes. Neither P. nor any of the organic
Compounds tested interfered with PP, determination. 3
Hinalietho'd for measuring PP; has been adapted for use as
;wm polymerase. Using this method, the formation of

r“

4.. ‘ "1'
k A. .‘I
' ' k
1 \ .L.‘ 7
O
4
0 ,V‘

 

    
  
  
 
  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
   
  
 
 
 
 
 

 

144 JOHNSON, SHANOFF‘, 111133, 301121, AND HANan

NADPH—as a consequence of the generation of PP, by RNA polymefi
ase—can be easily measured at a rate of 0.1—0.2 mnmole/Inin/ml. M
thermore, the enzymic assay procedure permits a continuous monitoring- In
of the RNA polymerase reaction. Consequently, a study of the ‘- -'«1, '
kinetics under a variety of conditions can be conveniently performedli‘
addition to these attributes, the spectrophotometric assay has certain
advantages over the radioactive assay. The amount of NADPH formed
is a direct measure of the total nucleotides incorporated into the intra-
chain phosphodiester link of the polyribonucleotide product. In the I“,
radioactive assay, a knowledge of the base composition of the productis
required in order to calculate the amount of polymer formed from the
incorporation of one nucleotide. Furthermore, the solubility of the
polymer product in trichloroacetic or perchloric acid is not a factor inthe
spectrophotometric assay as it is in the radioactive assay. .
The enzymic method for the determination of PP, should find applica- I,“
tion for the assay of many enzymes which generate PP, as a reaction 1
product. Some examples are DNA polymerase (14), RNA-dependent
RNA polymerase (15) polyriboadenylate polymerase (16), the amino-
acyl-sRNA synthetases (17), and the fatty acid activating enzymes (18).
Although PP, is a product of many biosynthetic processes, the intra- on
cellular concentration of PP, is low due, supposedly, to the action of I“
ubiquitous inorganic pyrophosphatases (19). The hydrolysis of PP, may -
be considered to be an energetically wasteful process since the energy
of the anhydride bond is lost as heat. However, in the phosphorylation
of glucose in liver microsomes (20) and in the formation of phase
phoenolpyruvate in the propionic acid bacteria (21) PP, serves as on
energy source. Futhermore, PP, has been implicated in photophos-
phorylation in Rhodospirillum rume (22) and in oxidative phos-
phorylation in Acetobacter suboxydans (23) and in Escherichia
(24). Knowledge of the intracellular levels of PP, in a variety of arm-
nisms might provide a clue to the metabolic fate of PP,. Methods have
not been available that are sufficiently sensitive or specific for the
termination of PP, in biological material. The use of this present : _
cedure, which is limited only by the sensitivity of the quanti "
determination of NADPH and by the specificity of the enzymes,- 1 ll
help clarify the function of PP, in metabolism.

 
   
 

IIIIFFsEEEcF!

   

   

 
 
 

   

Fra-

    
   
  
 

  

SUMMARY
1. An ensymic method for the determination of PP, has _
god. PP, can be quantitatively estimated from the ‘
the action of EDP-glucose 1

‘4 -,-' an.

      

  

' ‘30 men FOR PP, DETERMINATION

  
 

$11 phosphate compounds tested interfered in the PP,

    
   
 

{has been used for the assay of DNA-dependent RNA
coupling the generation of PP, by RNA polymerase .1
Ormation. The RNA polymerase catalyzed syntheses of g A
. 'boadenylate have been assayed by this procedure. . “ ‘

 
   

  
    
 
 
 
 
  
  
 
  
 
  
 
  
 
  
 
 
  
  
 
  
  
  
 
  
  
 
  
 
  

ACKNOWLEDGMENTS . ' . 1 1‘ ‘ ‘.'

. ‘ 7 e- ,.
,1 ." l” technical assistance of Madame M. DeBacker is acknowledged. ff," V

.M by National Institutes of Health Grant AM 11156. Michigan . _ T.) ‘
‘ 1.; rinlent Station Journal No. 4289. . _. 5'1.
W _ REFERENCES 5- ‘1
- ‘, H., AND Swallow, Y., J. Biol. Chem. 66, 375 (1925). ‘

.. , G. J., Base, S. T., Smear, L. L., AND HANSEN, R. G., J. Biol. Chem. ‘ 1:

3 (1966). , r

, B. K., am: sum, 0. G., Proc. Natl. Acad. Sci. Us. 54, 566 (1965). - 1 -‘ »’.

~' :31 C-o AND Bow, J. A., manuscript in preparation.

1t. m Bow, .1. A., J. Bacterial. 92, 1821 (1966). 1 _ ..
1’1. m Bow, J. A., J. Virology 1, 1274 (1967). . ,
1» C. A., AND ABELSON, J ., in “Procedures in Nucleic Acid Research" (8. 1.

   
   
  
 
  

1'1 and David R. Davies, eds.), p. 553. Harper & Row, New York, 19605 . f s"

. W. A” AND GILF‘ORD, S. R., Anal. Biochem. 2, 601 (1961). ‘ , .' i
' . A., 111m Hakeem, B. L., Biochem. Prepn. 3, 24 (1953). . - . .;

1 .1. 3, J. Biol. Chem. 241, 1830 (1966). . T‘s?
U., AND Huawrrz, J ., J. Biol. Chem. 242, 4897 (1967 ). . 1.1

- H. m Komn, M. W., Proc. Natl. Acad. Sci. U. s. 51, 807 (1954).

1' 2 N. J. P., J. Mol. Biol. 21, 115 (1966).

n. A., Science 131, 1503 (1960).

I. Am: ammu, 3., Proc. Natl. Acad. Sci. U. s. 54, 579 (1965).

1 J- T., 03112, P. J., m1 Hvawrrz, J., J. Biol. Chem. 237, 3786 (1962).

, - E. B., m) zAuscmx, P. C., J. Biol. Chem. 221, 45 (1956).
1 1111,1115 WAKIL, s. J., J. Biol. Chem. 204, 453 (1953).

1 R. C. m» Lam, H. A., Biochim. Biophys. A;‘?195:£)189 (1961). ‘

W. J ., m) Nashua, R. C., J. Biol. Chem. 239, 275 .

1’ H. G., Duns, J. J., AND LocHMiiLLsa, H., J. Biol. Chem. 241, 5692 (1966).

"“1“, M., Nature 216, 241 (1967).

‘ ' mn. L., KING, T. E., AND CHELDELIN,
_ 11.957).

11.1? 1 ' ~ nan. L., Biochim. Biophys. Am 34, 586 (1959).

V. H., J. Biol. Chem. 227, 135

 

 

ARTICLE 2

DEOXYRIBONUCLEIC ACID-DEPENDENT RIBONUCLEIC ACID

POLYMERASE 0F PSEUDOMONAS PUTIDA

 

BY

J. C. Johnson, M. DeBacker, and J. A. Boezi

Reprinted from Journal of Biological Chemistry,

216, 1222 (1971)

30

it‘lllt

\

.

 

 

 
 

 

 

 

T1111: Jornmn OF BIOLOGICAL CHEMISTRY
1111.246, No. 5, Issue of March 10, pp. 1222—1232, 1971
Printed in U.S.A.

Deoxyribonucleic Acid-dependent Ribonucleic
Acid Polymerase of Pseudomonas putida*

(Received for publication, June 5, 1970)

J. C. JOHNSON, M. DEBACKER, AND J. A. BOEZI

From the Departments of Biochemistry and of Microbiology and Public Health, Michigan State University,

East Lansing, M ichigan 48823

SUMMARY

Two forms of RNA polymerase (nucleoside triphosphate-
RNA nucleotidyltransferase, EC 2.7.7.6) from Pseudomonas
putida were resolved by chromatography in 50% glycerol on
phosphocellulose and purified. As determined by sodium
dodecyl sulfate-polyacrylamide disc gel electrophoresis, the
subunit structures of the two forms of the enzyme were
aside and agfifi’. The molecular weights of a, B, B’, and a
were 44.000, 155,000, 165,000, and 98,000, respectively. The
sedimentation coeflicients of azfifi’a and 0121313" as determined
by sucrose gradient centrifugation in 0.40 M potassium acetate
were 13 S and 12 S, respectively, but in 0.05 M potassium ace-
tate, the sedimentation coefficients were 19 S and 25 S.
Similar results were Obtained by analytical ultracentrifuga-
tion. Multiple protein bands resulted from polyacrylamide
disc gel electrophoresis of 0263'0' and azBB’. Each band was
enzymatically active in the unprimed synthesis of poly A 1p01y
U. RNA polymerase azfifi'a was 4- to 5-fold more active in
transcribing P. putida bacteriophage gh-l DNA and coliphage
T1 DNA than was (1.1352 With calf thymus DNA as a tem-
Plate. the two forms were equally active in RNA synthesis.
With denatured DNA or poly d(A-T), a236’ was twice as
active as was azBB’a. The rates of the DNA-directed polym-
erization reactions catalyzed by agfifi’a and agfifi' were 81'
fected difierently by added RNase. The initial rate of the
reaction catalyzed by azfifi’a was increased 10%. Whereas
that catalyzed by azfifi’ was increased 80 to 110%.

DNA-dependent RNA 1.)olymcrasc (ribonucleoside triphos—
lete-RNA nucleotidyltransferase, EC 2.7.7-6) has be?“ P11”—
ficd and characterized from a number of bacterial sources (1-6).
The enzyme from Escherichia coli has been the most extensively
studied. It has been purified by a variety of procel'lurcs .(l,
7‘16). and its catalytic and physical moperties have been studied
bl‘fl number of laboratories (9, 10, 12, 13, 17—31)-

A method utilizing phosphocellulose chromatography for the

c This work was supported in part by Grant (iB-‘i’lllkl from the
butional Science Foundation. This is blichil-Can Agriculture L)“
Periment Station Article 5112.

purification of E. coli RNA polymerase has been described by
Burgess (22). The subunit structure of enzyme prepared by this
method, designated core enzyme, was azfifi' (23). Burgess,
Travers, Dunn, and Bautz (24) showed that a, a subunit which is
involved in the initiation of RNA synthesis, was separated from
the core enzyme by phosphocellulose chromatography. The
subunit structure of the holoenzyme or complete enzyme was
agfifi’a. At the present time, only the subunit structure of the
E. coli enzyme has been documented in detail. Knowledge of
the structural as well as the catalytic properties of RNA polym-
erase from other sources must be acquired before a generalized
view of structure and function in the transcriptional process can
be set forth.

This report describes the purification of RNA polymerase from
Pseudomonas putida. Two forms of the enzyme have been re-
solved by phosphocellulose chromatography. Each form has
been characterized with respect to its subunit composition and
certain of its physical and catalytic properties.

EXPERIMENTAL PROCEDURE

.l-Iaterials—«Whatman DEAE-cellulose (DE-1) and phospho-
cellulose (P-l) were purchased from Reeve Angel, New York,
New York. UDP~glucose, NADP, glucose 6—phosphate dehy-
drogcnase, dithiothreitol, and the unlabeled 5’-phosphate de-
rivatives of the ribonucleosides were obtained from P-L Bio-
chemicals. 3H-Labeled ribonucleotides were from Schwarz
BiOResearch. Calf thymus DNA, herring sperm DNA, pan-
creatic RNase (type LA), and phosphoglucomutase were from
Sigma. Pancreatic DNase I (electrophoretically purified) and
catalase were obtained from Worthington. UDP-glucose pyro-
phosphorylase was isolated from calf liver (25) and recrystal-
lized twice. RNA polymerase from E. coli K—12 was a gift from
C. Scharrenberger and E. K. F. Bautz, Rutgers University.
Poly d(A-T) was purchased from Miles Laboratories, Inc., Elk-
hart, Indiana. Actinomycin D and nogalamycin were gifts
from Merck Sharpe and Dohme and The Upjohn Company,
respectively. Rifampicin was purchased from Mann. Acryla-
mide and bis-acrylamide were from (‘analco, Rockville, Mary-
land, and recrystallized according to the procedure of Loening
(26). Ethidium bromide and (Toomassie brilliant blue were
obtained from Calbiochem and (“olab Laboratories, Inc., Glen-
,mod, Illinois, reSpectively. Diethyloxydiformate was pur-
chased from Naftonc, Inc., New York, New York. Nitrocellu-

1222

Issue of March 10, 1971

lose. membrane filters, type B-6, were from Schleicher and
Schuell, Inc., Keene, New Hampshire.

Growth of P. putida—P. putida (the same or similar to ATCC
12633) was grown in lOO-liter volumes in a New Brunswick
Fermacell, model F-130. The growth medium contained the
following in grams per liter: yeast extract, 5; glucose, 8; NaCl, 8;
(NHJgHPOi, 6; KH2P04, 3; MgSOi-7H20, 1; and FeCl;, 0.005.
The cells were grown at 33° with aeration at 8 to 10 cu ft per min
into the early stationary phase and then harvested in a Sharples
centrifuge. The 1800 to 1900 g (wet weight) of packed cells
were stored at -20°.

Preparation of DNA and RNA-—P. putida bacteriophage gh-l
DNA (27, 28) and E. coli bacteriOphage T4 DNA were purified
by the method of Thomas and Abelson (29). (‘alf thymus DNA
was further purified by SDsl-phenol extraction. P. putida DNA
and E. coli ’H-DNA were prepared by the procedure of Thomas,
Berns, and Kelly (30). After RN ase treatment and phenol
extraction, the DNA preparation was mixed with diethyloxydi-
formats and incubated at 22° for 30 min. Following phenol
extraction, ethanol precipitation, and isopropyl alcohol f ractiona-
tion, the DNA was dissolved in a butler containing 0.01 M Tris
acetate (pH 8.0), and 0.1 M sodium acetate and dialyzed for 10
hours. Denatured DNA was prepared by heating dilute solu-
tions of DNA at 100° for 20 min, followed by quick cooling in
ice water. E. coli-soluble and ribosomal 3H -R N A were prepared
as described by Payne and Boezi (31), Native DNA and RNA
concentrations were determined spectrophotometrically based
on the extinction coefficient Eti’t = 200.

Radioactiw Assay of RNA Polymerase—The radioactive assay
of FNA polymerase measured the incorporation of CMP into a
form insoluble in trichloracetic acid. The reaction mixture
contained 20 mM Tris acetate (pH 8.0), 4 mM magnesium ace—
tate, 1 mM manganese acetate, 60 mM ammonium acetate, 0.4
out each of ATP, GTP, (TP, and FTP, and 110 pg per ml of
DNA and RNA polymerase. CT P was labeled with 3H at 5 X
103 Chm per nmole. Incubation was at 30° for 10 min unless
otherwise indicated. After incubation, 50- to 250p] samples
of the reaction mixtures were mixed with 100 pl of 0.1% SDS.
(.‘old 10‘}, trichloracetic acid (5 ml), which had been filtered
through Celite, and 250 pg of herring sperm DNA were added.
After 15 min at 0—4°, insoluble material was collected on a nitro—
cellulose membrane filter. The filter was washed with four 5-ml
Minions of cold 10% trichloracetic acid, dried, and then moni-
tored for radioactivity in a liquid scintillation spectrometer.
The scintillation fluid (5 ml) contained 4 g of 2,5 bisl2-(5-tert-
l)utyl-henzoxazolyl)]thiophene per liter of toluene. One unit of
RNA DOIymerase activity was defined as that amount of enzyme
Which catalyzed the incorporation of l nmole of (.‘MP per hour
3t 30°. The specific activity was the number of units per mg of
ltmtxein. Protein concentrations were determined by the method
of Lowry et al. (32) with bovine serum albumin as a standard.

SPeClTOletometric Assay of RNA Polymerase—The spectro-
photometric assay of RNA polymerase as described by Johnson
ii ‘11- (33) Coupled the formation of inorganic pyrophosphate t0
NADP‘r reduction. In addition to the components used in the
r{Idioactive assay, the reaction mixture contained 0.2 mM
NADP+, 0.2 mm UDP-glucose, and excess phosphoglucomutase,
glucose 6‘DhOSphate dehydrogenase, and UDP~glucose pyrophos—
l’hOTYlase. Incubation was at 30°. Calculation of the number

1The abbreviation used is: SDS, sodium dodecyl sulfate.

J. C'. Johnson, M. DeBacker, and J. A. Boezi 12233

of nanomoles of NADPH formed was made with the molar extinc-
tion coefficient of 6.2 X 10’ (34).

Assay of Other Enzyme Activities—The assay for exo- and endo-
DNase and RNase measured the solubilization of 3H-l)NA
and 3H-soluble RNA and 3H-ribosomal RNA. An additional
assay for endonuclease measured the change in sedimentation
patterns of ’H-DNA in alkaline sucrose gradients. Inorganic
pyrophosphatase was determined by measuring the change in
inorganic pyrophosphate by using the coupled UDP-glucose
pyrophosphorylase assay system (33). ATP-AMP phospho-
transferase was measured by the formation of 3H-ADP from 3H-
ATP and AMP. Enzymes which hydrolyze the ribonucleoside
triphosphates were measured by the formation of ribonucleoside
mono- and diphosphates which were identified by paper chroma-
tography in isobutyric acid-NHiOH-HQO (66: l :33). ATP-R NA
adenylyltransferase was measured as described by Payne and
Boezi (31). (.‘atalase was determined spectrophotometrically
by measuring the disappearance of H202 at 240 nm. lil)l-’—
glucose pyrophosphorylase was measured as described by Al-
brecht et al. (25).

Polyacrylamide Disc Gel Electrophoresis—Polyacrylamide gels
were prepared according to the general procedures of Davis (35)
and Ornstein (36). Clear gels were prepared from a solution
which was 3.75% in acrylamide by mixing together 1 part water,

 

  
  
  

 

 

 

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50 ‘ IOO
FRACDON NUMBER

FIG. 1. Phosphocellulose chromatography 0f “RAE-cellulose
fraction. Dialyzed DEAR-cellulose fraction (960 ml) was applied
at 3 ml per min to a phosphocellulose column (5 x 40 cm) which
had been equilibrated with phosphate—glycerol bufi'er (002 M
potassium phosphate, pH 7.5, 0.005 M 2-mercaptocthanol, 0.000] M
EDTA, and 50% glycerol (v/v)). Following application of the
enzyme fraction, the column was washed with 4 liters of the Phos-
phate-glycerol buffer at. a rate of 5 ml per min. A linear 4-liter
gradient from 0.0 to 0.4 M KCI in phosphate-glycerol bufi‘er was
used to elute RNA polymerase. The column was developed at 6
ml per min and 170 fractions of ‘20 ml each were CUllected.
O“—*O, absorbence at 280 nm; A—— bA, CMP incorporation with
gh-l DNA as the template; O—~——O, CMP incorporation with calf
thymus DNA as the template; AwA, KCI concentration as
determined from conductivity measurements; F. (in the
inset), the ratio of CMP incorporation with gh—l DNA as the
template to that with calf thymus DNA as the template. For the
assay of RNA polymerase, 10-pl samples of the fractions were as—
sayed in (SO-pl reaction mixtures according to the method described
under “Radioactive Assay of RNA Polymerase.” Fractions 93
through 115 (Phosphocellulose Fraction I) and 140 through 153
(PhOSphocellulose Fraction II) were pooled,

I50

1224

1 part Solution A (30.3 g of 'l‘t'is, 0.23 ml of N,N.i\7'..\"~tetra-
methylethylenediamine, 48 ml of l x ”Cl, and water to a total
of 100 ml), 2 parts Solution ll (15.0 g of acrylamide, 0.55 g of
bisacrylamide, and water to 100 ml), and 4 parts Solution (‘
(0.14 g of ammonium persulfate and water to 100 ml). (llass
tubes with an internal diameter of 5 mm were filled with the gel
mixture to a height of 7 cm. Water was layered on top of the gel
mixture to form a level interface. bellowing polymerization,
the gels were brought to the temperature of electroi)horesis and
then sttl'ijected to electrot)l'ioresis for 30 min at 5 ma per gel prior

 

 

 

 

 

 

 

 

 

 

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50 100

FRACHON NUMBER

FIG. 2. “BAH-cellulose chromatography of I’hosphocellulogo
Fractions I (upper) and [I (lower). l)ialyzed Pl‘iosphoeellulose
Fraction I (510 ml) was diluted to 1020 ml with Buffer ASH-glyc-
erol (0.01 M Tris-”Cl, pH 8.0, 0.005 M 2-mercaptoethanol, 0.01 M
MgC-lg, 0.000] M EDTA, and 50?}. glycerol (v/v)), The diluted
enzyme fraction was applied at 3.3 ml per min to a DEAR-cellulose
column (2.5 X 19 cm) which had been equilibrated with Buffer
ASH-glycerol. The column was then washed with 00 ml of Buf’fer
ASH-glycerol. A linear 400-ml gradient from 0.0 to 0.40 M NaCl in
Buffer ASH-glycerol was used to elute RNA polymerase. The
column was developed at 3.3 ml per min and 125 fractions of 3 ml
each were Collected (upper). O~—~O, absorbanee at 280 um;
..—., CMP incorporation with gh-l DNA as the template;
As A, NaCl concentration as determined from conductivity
measurements. For the assay of RNA polymerase, 5-,ul samples
of the fractions were assayed in (SO-pl reaction mixtures according
to the method described under “Radioactive Assay of RNA Polym-
erase." Fractitms ti? through 85 were pooled. Dialyzed I’ht;g-
phoeellulose Fraction II (200 ml) was diluted to 400 ml with Butler
ASH—glycerol and applied at 2.5 ml per min to a IH'JAIC-cellulose
column (2.0 X 17 cm). The column was then washed with 80 ml of
Buffer ASH-glycerol. A linear 250-1111 gradient from 0.0 to 0.40 M
NaCl in Buffer ASH-glycerol was used to elute RNA polymerase.
The column was developed at 2.7 ml per min and 130 fractions of 2
ml each were collected Newt-7f). O-‘A—VO, absorbance at 280 nm;
. -fio, (‘31P incorporation with gh-l DNA as the template;
‘—-—«‘, Na(‘l concentration as determined by conductivity meas—
urements. For the assay of RNA polymerase, lO-nl samples of
the fractions were assayed in (30441 reaction mixtures as described
under “Radioactive Assay of RNA Polymerase.” Fractions 74
through 00 were pooled.

DNAstlcpcndenl REX-l Polymerase of Pseudomonas punt/a.

to sample application.
was stained for 4 to 12 hours with 0.3“} (‘UOI’IlttSs‘lt‘ blue in 10'";
trichloracetic acid containing 5"; methanol.
was removed by rinsing the gels with 10'] trichloracetic acid,

Vol. 240, No. 5
1901],,“ng electrophoresis, the protein
l'nbound stain

SDS-polyacrylamitle disc gel electrophoresis was perforim-tl

accort‘ling to a mmlification of the prm-et‘lure of Shapiro, Vit‘iticla.

and Maizel (37).
acrylamide were prepared by the procedure (‘lescrihed above with

Acrylamide-h‘l)S mixtures containing 3.750.

the exception that Solution .\ contained 0.8 M sodium phosphate
(pll 7.1), 0.8“; SDS, and 0.23 ml of N.N.N'..\"—tetrametliyl
etbylenediamine per l00 ml of solution. For acrylanude-S08
mixtures with 5‘} acrylamide, Solution B contained 20 g of
acrylan'iide, 0.735 g of bisacrylamide, and water to 100 ml. For
acrylamide-QIH mixtures with 3‘; tu-rylmnitle, Solution ll con-
tained 12 g of acrylamidc, 0.44 g of bisacrylamide, and water
to 100 ml. S1’N-polyacrylamide gels 7 or H cm in height were
prepared. Following electrophoresis, the SDS-t)olyacrylaniide
gels which contained 10 pg of protein or more were stained hr
the procedure described above. Sl)S-t)olyacrylantide gels which
contained less protein were stained with 0.4"}; (‘oomassic blue in
10"; trichloracetic acid and 20";- methz-inol for 12 hours. De
staining was performed by first washing the gels 6 hours in 10';
trichloraceti' acid and 33"; methanol and then rinsing the gels
in 10‘,‘ trichloraceti' acid until areas of the gel without protein

were colorless.
mastitis

Part/imitate of RNA Polymerase

The entire purification procedure was performed at 0- 4".

. lnitia/ I:'.1'tr(1ct~#.>\ block of frozen 1’. putida cells, 9510 g (wet
weight), was cut into small pieces. Buffer .-\Sll (0.01 M 'l‘ris-llt'l
(pll 8.0), 0.01 M .\Ig(.'l—_i, 00001 M EDTA, and 0.005 M 2-inercap-
toethanol) was added (900 ml) and the mixture was stirred until
a smooth cell suspension was obtained. The cell suspmision me
twice passed through the pressure chamber of a .\Ianton-(iziulilt
lab homogenizer (38‘) which was maintained at 7,000 psi. The
process, which provided nearly complete rupture of the cells.
resulted in a viscous cell extract. Pancreatic DNasc 1 (ll-5 #L’
per tnl) was added to the cell extract and the mixture was stirred
for 15 min. Following centrifugation at 12,000 X g fer 30 min.
the supernatant fraction was decanted and saved. The Dell“
fraction was suspended in 200 ml of Butler ASH and reccntri»
fuged. The resulting supermitant fraction was combined with
the supernatant fraction from the first centrifugation 01111131
extract, 2,150 ml).

[fig/i. Speed Supernatant l’rttcttbrt—Jl‘lie initial extract \th
centrifuged at 78,000 X g for 90 min. The clear, straw-mlnrwl
supernatant fraction was collected by decantation. 'l‘ltt’ t't‘llt‘l
fraction was suspemled in 200 ml of Buffer ASH and recenti'i-
fused. Supernatant f1‘tt('tlt)ttS from the first and set-mid centrif-
ugations were combined (high speed supernatant fraction, 1,73”
ml).

:ltmno-nium Sulfate (60";) Fractions—'l‘he high speed S111191113”
tant fraction was diluted with buffer ASH to a protein («inventin-
tion of 2.5 mg per ml. Solid ammonium sulfate (410311 “'3‘
added to the diluted high speed supernatant fraction (2500 ”ll;

to give 30‘"; saturation. After stirring for 30 min, the stispett-

4

sion was centrifuged at 12,000 X g for 30 min. The supernatant
. . , . ."l
fraction was brought to 60f“; saturation with the addition of 4J-
9: 01 ammonium sulfate. After stirring for 30 min, insolnltlt‘

r’: .‘ '

Issue of March 10, 1971 J. C. Johnson, M. DeBacker, and J. A. Boezi 1225

Tutu: I
Summary of purification

The purification of RNA polymerase through the DEAR-cellu-
lose fraction was conducted on material from 990 g (wet weight)
of cells. After this, only one-half of the DEAE—cellulose frac-
tion was processed. RNA polymerase activity was measured
by the method described under “Radioactive Assay of RNA
Polymerase.” gh-l DNA was used as the template.

 

 

 

Enzyme mation l as; \ tear first;
mg rmi]: unfit/mg
Initialextract.........,...._....13.0 X 10‘ 99X10‘ 8
High speed supernatant fraction 6.1 X 10‘ 70 X 10‘ 11
60‘} ammonium sulfate fraction. 51 X 10‘ 270 X 10‘ 53
DEAR-cellulose fraction. . . . . , . . . 1.0 X 10‘ 330 X 10‘ 210
Phosphoccllulose Fraction I ..... 73 16 X 10‘ 2200
l’hosphocellulose Fraction II. . . . 62 2.6 X 10‘ 420
DEAEcellulose Fraction PCI. .. 30 9.6 X 10‘ 3200
DEAE-cellulose Fraction PCII . . 22 1.4 X 10‘ 640

material, which included RNA polymerase, was collected by
(entrifugation. Bufier ASH (800 ml) was added and the mix-
ture was stirred. The resulting solution was dialyzed for 9 hours
in 12 liters of Buffer ASH (60C; ammonium sulfate fraction,
940 ml).

MAE-cellulose Fraction—Acid and base~washed DEAE-
cellubse suspended in Bufier ASH was poured into a column and
packed under air pressure of 1 to 2 p.s.i. The packed column
(14.2 X 60.0 cm) was washed with Buffer ASH until the pH of
the outflow was 7.0 to 7.2. The 60‘}; ammonium sulfate frac—
tion was diluted with Bufi'er ASH to a protein concentration of
8.5 mg per ml (6 liters) and applied to the column at a rate of 160
ml per min. Following application of the diluted enzyme frac-
tion, the column was washed with 16 liters of Buffer ASH con-
taining 0.1 M NaCl. RNA polymerase was eluted with Bufier
ASH containing 0.2 M NaCl. Fractions containing RNA p0-
lb'memse activity were pooled (DEAE-cellulose fraction, 5 liters).

Phosphocellulose Fractions [ and [l—«Acid- and base-washed

phosphocellulose was suspended in phosphate-glycerol buffer
(0.02 M potassium phosphate (pH 7.5), 0.0001 M E1)TA,0.005 M
2‘mt’rcaptoethanol, and 50% glycerol (v/v)), poured into a col—
umn, and packed by gravity. The packed column (5 X 40 cm)
“to first washed with 750 m1 of phosphate-glycerol buffer con—
mlninfi 0.5 M KCI and then with phosphateglyccrol buffer until
chloride ion was no longer detected in the outflow. Th9 PH 0f
the outflow was 7.5.
'The DEAE-cellulose fraction was dialyzed for 12 hours in 50
liters of a buffer containing 0.01 M Tris-HCI (pH 8.0), 00001 M
EDTA, and 0.001 M 2-mercaptoethanol. Dialysis was con-
tinued for 20 hours with one bufl'er change in 20 liters of phos-
l’hate‘glyccrol buffer. During dialysis against phosphate-
glycerol buffer, the DEAE-cellulose fraction was reduced in
Volume to 1620 m1.

Gli'cerol (300 ml) was added to the dialyzed DEAE—cellulose
metion. The fraction was divided into two equal parts. 0116
Rm (960 ml) was stored at —20° for processing at a later date.
“18 other part was applied to the phosphocellulose column at a
me 013 ml per min. Following application of this enzyme
fmetion, the column was washed with 4 liters of PhOSI’hi‘te'

 

  

FIG. 3. Schlieren sedimentation pattern of Fractions PCI
(upper) and PCII (lower). Fraction PCI at a concentration of3.4
mg per ml in a buffer containing 0.05 M Tris acetate (pH 8.0), 0.01
M magnesium acetate, 0.001 M 2-mercaptoethanol, and 0.40 M po-
tassium acetate was centrifuged at 4.6" in a Kel F centerpiece in
the An-l) rotor of the Spinco model E analytical ultracentrifuge.
The picture was taken at 29 min after reaching a speed of 56,000
rpm. The schlicren optical system was used and the phase plate
angle was 65°. Fraction PCII at a concentration of 3.3 mg per ml
in a buffer containing 0.05 M Tris-HCI (pH 8.0),0.01M magnesium
chloride, 0.001 M 2—mercaptoethanol, and 0.40 M potassium chlo~
ride was centrifuged at 20.1" in the Kel F centerpiece in the An-D
rotor of the Spinco model E analytical ultracentrifuge. The pic-
ture was taken 38 min after reaching a speed of 56,000 rpm.

glycerol buffer at a rate of 5 ml per min. A linear gradient from
0.0 to 0.4 M KCI in phosphate-glycerol buffer was used to elute
RNA pelymerase (Fig, 1). Two peaks of RNA polymerase
activity were resolved. The first peak eluted at 0.18 M KCI
and the second eluted at 0.30 M KCI. Fractions of the first peak,
which had a ratio of enzyme activity of 3.0 with ghsl DNA as
template to that with calf thymus DNA as template, were pooled
(Fraction PCI, 480 ml). Fractions of the second peak, with an
enzyme activity ratio of 0.5, were pooled (Fraction PCII, 200 ml),

DEA E-rclluvlose Fractions PC] and PC] I —~DEAE-cellulose

1226

suspended in Buffer ASlI-glycm‘ol (Buffer ASH with 50‘";-
glycerol (v /v)) was used to prepare two columns. The. columns,
2.5 X 19 em and 2.0 X 17 cm, were first washed with 200 ml of
Buffer .—\Sll~glyccrol containing 0.5 M Na(‘l, followed by 500 ml
of Buffer ASH-glycerol.

In preparation for chromatogra])hy, l’hoslihocellulose Frac-
tions 1 and II were ‘ach dialyzed for 10 hours in 10 liters of
Buffer ASH-glycerol, and then were diluted 2—fold. l’hospho-
cellulose Fraction I was applied to the larger DEAR-cellulose
column and Fraction II to the smaller column. After each
column was washed with a column volume of Buffcr ASH-
glycerol, RNA polymerase was eluted with a linear gradient of
0.0 to 0.4 M Na(‘l. The elution profiles are presented in Fig. ‘2.
For both colnn‘nis, the peak of RNA polymerase activity was
eluted at 0.17 M Na(‘l. l’iak fractions from each column were
pooled and dialyzed for 10 hours in ‘2 liters of a solution at 75‘}

saturation with ammonium sulfate. The white flocculent pre-

 

 

 

 

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F RACTION NUMBE R

FIG. 4. Sticrose gradient centrifugation of Fractions PCI
(upper) and l’tr‘ll (lower) in 0.40 M potassium acetate. Fraction
PCI (50 pg) or Fraction PCII (107 pg), catalase (79 fig), and UDP-
glucose pyroplnisphorylase (10 rug) were dissolved in 0.10 ml ”f a
buffer containing 0.05 M Tris acetate (p118.2),0.005 M 2-mercupto-
ethanol, 0.005 M magnesium acetate, and (HO M potassium acmmfl
The enzyme mixtures were layered on 5 to 20‘} linear sucrose
gradients (v1.8 ml) prepared in the above mentioned buffer and
centrifuged in a Spinco model L-2 centrifugé at 34,000 rpm in the
SW 39 rotor for 9 hours at 3°. After centrifugation, a hole ,1“;
punched in the bot tom of each centrifuge tube and 31 fractions per
gradient were collected. (Tatalase and [EDP-glucose pyrophos-
phorylase were assayed on 30— and 5-pl samples, respectively, as
described under “Assay of Other Enzyme Activities.” RNA
polymerase was assayed on 15—pl samples in 125—141 reaction mix—
tures with gh-l DNA as described under “Radioactive Assay of
RNA Polymerase" except that reaction mixtures were incubated
for ‘20 min. Recovery of Fraction PCI activity was 06‘3“}, and that
for Fraction PCII was 100%. Am‘, catalase as measured by
the change in absorbance at 240 nm; .——., UHF-glucose pyro-
phosphorylase as measured by NADPH forn‘iation; Q——-..Q,
Fraction PCI (upper) or Fraction PCII (lower) as measured by
CMP incorporation.

DNA-(lcpcmlcul RNA Polymerase of Pseudomonas punt/(L

Vol. 246, No. 5

eipitate which formed was collected by centrifugatitm at 20,000
x g for 20 min and was dissolved in a buffer containing 0.0.1") M
Tris acetate (plf 8.0), 0.01 M triagnesium acetate, 0.000] n
lilYl‘A, 0.001 M dithiothreitol, and 50?} glycerol at a protein
com'cntration in excess of 10 mg per ml. The enzyme fraction
from each column was dialyzed for 12 hours in 200 ml of the same
buffer (l)1£.-\l‘}-cellulose Fraction l’f‘l, 1.95 ml, and l)li.—\E-ccllu-
lose Fraction l’('l1, 1.35 ml).

Summary and Comments on l’urfticrilionHA summary of the
burification is presented in Table I. The determination of RNA
polymerase activity in the enzyme fractions through the l)li.\li-
cellulose. fraction is complicated by the presence of several
cc'mt‘aminating enzyme activities, such as DNase and RNase,
and by fragments of I’. putido DNA.

P. putida cell rupture by means of the Manton-(laulin lah
homogenizer was easy and efficient. One kilogram of frozen
cells could be broken in a few minutes and cell rupture was es-
scntially complete as judged from the direct microscopic obser-
vation of the broken cell suspension.

l’hosphocellulose chrormitogra})hy was performed in 50’,
glycerol to stabilize the enzyme and to prevent the conversion of
Fraction PCI to Fraction l’(‘11. When purified Fraction l’t‘l
was chromatograiihed on phosi)hocellulose in 50“; glycerol, it
remained intact and eluted as a single peak. When Frau-tion l’t'l
was chroniatographed in 5"; glycerol, some conversion to Frac-
tion l’t‘ll was observed.

('ritcria 0f l’urity————'l‘he purity of RNA polymerase Fractions
l’(‘l and l’(‘ll was evaluated by the following criteria. At the
final step of the purification, chromati)graphy on l)l‘lAF.-ccllu-
lose, specific enzymatic activity was constant throughout the
peak fractions (Fig. 2). In buffers containing 0.4 M salt, a single
symmetri'al schlieren pattern was observed in the analytical
ultraccntrifugc (Fig. 3). 1n i)i)lyacrylamide disc gel electro-
phoresis, comigration of protein and enzymati' activity were
found. In Sl)S—polyacrylamiile disc gel electro}whoresis, ol‘ll.V
subunits which were derived from R NA polymerase \Vt’l‘t’ olt-
served. Data for the. last two criteria are given below.

Fractions PCI and l’(‘ll were examined for the presence of
contaminating enzymes which might interfere in the assay of
RNA polymerase. No DNase and RNase activities \\'t'I‘t‘
detected. When 5 to 25 times the amount of protein iudinaril.‘
used in the assay for RNA polymerase was incubated for 4 hours
With 311-11NA or with 3H-soluble or 3ll-ribosmnal RNA, no
radioactivity was solubilized. Also, the alkaline sucrose til?“
dient profile of 311-11N‘A was unchanged after incubation with
Fraction l’(‘l or with Fraction l’(‘11. Other activities “lath
were not detected were inorganic pyrophos]ihatasc, ;\'1‘1)._\,\ll’
l’l“l-‘lthutl'ansferase, ATP—BNA adenylyltransferase, and "'l'
zymes which hydrolyze the rilmnucleosidc triphosphates to their
mono- or diphosphatc derivatives.

A few of the preparations of Fractions PCI and PC” \Wl't‘
contaminated with materials that sedimented at 5 S and at 25 5.
These contaminants amounted to 5 to 15“} 0f ”“9 Will ”ml
were removed by rechromatograIthy on phosi)hocellulosc HI” 1‘."
sucrose or glycerol gradient ccntrifugatioi‘i.

Churactcrimtion of Fractions PCI and PCI]

Storage—It‘ractions PCI and PCII were. stored in liqnit.l 1111“"
gen at. protein concentrations greater than 10 mg l)”. ml 1.“ :1
buffer containing 0.05 M Tris acetate (pH 8.0), 0.01 3“ "“‘li'wimin
acetate, 0.0001 M EDTA, 0.001 M dithiothrcitol, and 0” '

V

"4+. ‘5

Issue of March 10, 1971

   
    

 

 

 

 

 

 

 

 

 

 

 

“l2
2?
A ~ 8
«318
9 E
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5 o_4.t_=
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59 ~e
4 -14

5 [0 IS 25

FRACHON NUMBER
F105 Sucrose gradient centrifugation of Fractions FCI
Fraction

(upper) and PCII (lower) in 0.05 M potassium acetate.
PCI (77 pg) or Fraction PCII (80 pg), catalase (59 pg), and UDP-
glucose pyrophosphorylase (8 pg) were dissolved in 0.10 ml of a
buffer containing 0.05 M Tris acetate (pH 8.2), 0.005 M 2-mercapto-
ethanol, 0.005 M magnesium acetate, and 0.05 M potassium acetate.
The enzyme mixtures were layered on 5 to 20% linear sucrose
gradients (43 ml) prepared in the above mentioned buffer and
cent-|'lf‘1t§ed in a Spinco model L—2 centrifuge at. 34,000 rpm in the
SW 39 rotor for 9 hours at 3°. After centrifugation, a hole was
punched in the bottom of each centrifuge tube and 30 fractions
P0P gradient were collected. Enzyme assays were performed as
d(”scribed in the legend to Fig. 4. Recovery of Fraction PCI
“emit." “'88 83% and that for Fraction PCII was 59%. A—w—A,
fatalase as'measured by the change in absorbance at 240 nm;
9““.’ UDP'glucose pyrophosphorylase as measured by
MDPH formation; O—O, Fraction PCI (upper) or Fraction
PC” (lower) as measured by CMP incorporation.

glycerol. No loss of activity over a period of several months
“as observed, even after repeated freezing and thawing. Frac-
tions PCI and PCII were also stored at —20° for several weeks
Without loss of activity.

('llram'olet Absorbances—In several different preparations of
Fractions PCI and PCII, the 1123031250 ratios were 1.5 to 1.6.
The hum for each was 278 nm. For either Fractions PCI or
I’Cll, a protein concentration of 1 mg per ml as determined by
L0Wry protein analysis gave an absorbance of 0.67 at 280 nm.
‘Sedimentation Coefiicienta—T he sedimentation coefficients of
lmctions PCI and PCII were greatly influenced by the ionic
s”With of the solvent. The results demonstrate that both
Fmctions PCI and PCII aggregate in solvents of low ionic
:‘Im‘gth. When measured with the analytical ultracentrifuge
"1a buffer containing 0.05 M Tris acetate, 0.01 M magnesium
acetate, 0.001 M 2-mercaptoethanol, and 0.40 M potassium ace-
tate (DH 8.2), the sedimentation coefficient of Fraction PC‘I was
1230362") and that of Fraction PCII was 11.8 (836???“ - When
mgasured in the above buffer containing 0.05 M potassium ace-
tate rather than 0.40 M potassium acetate, the sedimentation
(’Wflicient of Fraction PCI was 19_9 (335?? and that. of Frac-
thll PCII was 26.5 (8535:”),

.w

J. C. Johnson, M. DeBacker,

1227
and J . A. Boezi

M‘Tfl

20a

in

 

 

 

 

 

O
f

ABSORBANCE AT 550 nm

05~ "I

J

 

 

 

 

 

 

 

 

 

 

 

 

l l i I 1 g 1
2 4 6 2 4 6
DISTANCE FROM ORIGIN (Ch/II

FIG. 6. Polyacrylamide disc gel electrophoresis of Fractions
PCI (left) and PCII (right). Polyacrylamide gels containing
3.75% acrylamide were prepared as described under “Polyacryl-
amide Disc Gel Electrophoresis.” Fraction PCI (20 #8) 01' Frac-
tion PCII (20 pg) in 10 pl of a buffer containing 0.01 M Tris acetate
(pH 8.0), 0.005 M magnesium acetate, 0.005 M 2-mercaptoethanol,
and 5% glycerol was layered on a gel. Electrophoresis was per-
formed at 5 ma per gel for 50 min at 0 to 4° in a buffer containing
0025 M Tris-0.2 M glycine, pH 8.9. Following electrophoresis,
the protein contained within the gels was stained with Coomassie
blue. Each gel was then scanned at 550 nm with a Gilford linear
transport.

Fractions PCI and PCII were also analyzed by sucrose gra-
dient centrifugation in buffers containing 0.40 M and 0.05 M
potassium acetate. The results are presented in Figs. 4 and 5.
In the buffer with 0.40 M potassium acetate, Fraction PC‘I co-
sedimented with the UDP-glucose pyrophosphorylase marker
(Fig. 4). Under the same conditions, Fraction PCII sedimented
somewhat more slowly than the UDP-glucose pyrophosphoryl-
ase marker, but slightly faster than the catalase marker. With
catalase as the reference (330,... = 11.3), the sedimentation co-
efficient of Fraction PCI was 13 S and that of Fraction PCII
was 12 S. In the buffer containing 0.05 M potassium acetate
the faster sedimenting forms, i.e. the aggregates of Fractions PCT
and PCII, were observed (Fig. 5). Again, with catalase as
the reference, the sedimentation coefficient of Fraction PCI was
19 S and that of Fraction PCII was 25 S.

Polyacrylamide Disc. Gel EIBCITOPhOTeWR'Iulti|)le protein
bands (probably due to aggregation) were observed for Fractions
pcl and PCII in polyacrylamide gels (Fig. 6). The proteins
contained within duplicate gels which had not been stained with
Coomassie blue were tested for enzymatic activity by an assay
in sit-u (39). Following electrophoresis, the gels were incubated
in reaction mixtures containing ATP, UTP, and Mn2+ to pro-
mote the unprimed synthesis of poly A ~poly U. After incuba-
tion, the polynucleotides were stained with ethidium bromide.
Polynucleotides were formed in all regions of the gels which
contained protein, including the protein at the origin and that

l 228

’7'," 1.?

:7" we

.vvir.

2
l

I
.0.

 

FIG. 7. SDS-polyacrylamide disc gel electrophoresis of RNA
polymerase. Escherichia coli (lefl), Pseudomonas putida Fraction
PCI (center), and combined E. coli and P. pulida Fraction PCI
(right). Reaction mixtures containing 75 pg of E. coli (holoen-
zyme) or P. putida Fraction PCI or 75 pg of each were incubated
in 150p] of a mixture containing 0.1M sodium phosphate (pH 7.1),
1.0% SDS, 1.0% 2~1nercaptoethanol, and 5% glycerol for 3 hours at
37°. The mixtures (3 pl) were layered upon SDS-polyacrylamide
gels (11 cm) prepared from an acrylamide-SI)S mixture containing
3.75% acrylamide. Electrophoresis was performed at 25° for
4% hours at 8 ma per gel in 0.1 M sodium phosphate, pH 7.1, and
0.1% SDS. The protein was stained with Coomassie blue.

small amount which had migrated about 3.5 cm from the origin.
Gels incubated in reaction mixtures from which ATP, UTP, or
MnH were omitted did not stain with ethidium bromide.
Subunit Structures of Fractions PCI and PCII—The subunit
structures of F ractions PCI and PCII were examined by the
sDS-polyacrylamidc disc gel electrophoresis technique (37, 40).
By means of this technique, the constituent subunits were

DNA-dependent RNA Polymerase of Pseudomonas putida

Vol. 246, No.5

 

 

 

 

l l l T l
30L- _.
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20- _.
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[I] -——o bovlnoummalbumln
3 6"" \o--—\o oyruvotl klnou "
5— _.
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DISTANCE FROM ORIGIN (cm)

FIG. 8. Molecular weight determination by SDS—polyacryl-
amide disc gel electrophoresis. Curve .4, each protein was sep.
arately incubated in a mixture containing 0.1 M sodium phosphate.
(pH 7.1), 19} SDS, and 19}, 2-mercaptoethanol for 3 hours 3137”.
In the case of myosin, the incubation mixture also contained 6 M
urea. After incubation, each protein was layered on a SDS-poly-
acrylamide gel prepared from acrylamide-SI)S mixtures contain-
ing 5% acrylamide. Electrophoresis was performed at 7.5 ma per
gel for 3 hours at 22° in 0.1 M sodium phosphate, pH 7.1, and 0.1‘}
SDS. Curve B, proteins were incubated together in a mixture
containing 0.1 M sodium phosphate (pH 7.1), 1% SDS, 1?}. 2»mer~
captoethanol, and 5% glycerol for 3 hours at 37°. After incuba-
tion, the mixture was layered on a SDS~polyacrylamide gel pre-
pared from an acrylamide—SI)S rnixt ure containing 5‘} acrylamide.
Electrophoresis was performed at 1'2 ma per gel for 3 hours at “25°
in 0.1 M sodium phosphate, pH 7.1, and 0.1% SDS. The proteins
were stained with Coomassie blue and the distances of migration
from the origin were measured. The molecular weights of the
polypeptides used as standards were 220,000 for myosin (47),
165,000 for E. coli B’ (‘23), 155,000 for E. coli IS (23),95,000 for E. coli
a (24), 68,000 for bovine serum albumin (48), 57,000 for pyruvale
kinase (49), 40,000 for aldolase (50), 39,000 for E. coli a (2‘3), 37.011)
for pepsin (51), 31,000 for pancreatic DNase I (52), and 13,7000"
pancreatic RNase, type l-A (53). A, subunits of P. pnlida RNA
polymerase; 0 (Curve A) and 0 (Curve 8), polypeptides used as
standards.

identified, their molecular weights were estimated, and the sub»
unit. structural formulas were determined.

Fraction PCI was incubated at 37° for 3 hours in 0.1 a sodium
phosphate (pH 7.1), 1% SDS, and 1% 2-mercapt0cthanol. Th9
resulting subunits were separated by eleCtI‘OPhOWSI“ 0“ SDS—
polyacrylamide gels. Four subunits were resolved (Fifl- 7’.
center gel). For comparison, the subunits derived from E- 0’“
RNA polymerase (holoenzyme) were analyzed in a similqr
manner (Fig. 7, left gel). The four E. coli subunits, which hare
been designated 6’, B, a, and a in order of increasing “‘Ob‘I't-‘I
(23, 24), and the four P. putida subunits were similar. In k"91"
ing with the nomenclature used for the E. coli subunits, the R
Putida subunits derived from Fraction PCI were designated B” d’
a, and a. By means of the same procedure, the B’, )3, and“
subunits but not the 0’ subunit, were derived from Fraction Pi.”-
The 5', e, and (1 subunits of Fractions PCI and PCII had MW“
mobilities.

‘. 1.. g _

 

 

 

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lssuc of March 10, 1971
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IN ARBITRARY UNITS

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PROTEIN APPLIED TO GEL (pg)

F100. Relationship between amount of each subunit tllld
amount of protein applied to Si)S-polyacrylamide gels. Frac—
lions PC] or PCII was incubated in a bufl'er containing 0.1 M
sodium phosphate (pH 7.1), 1’}; SDS. and 1‘} ‘2-mercaptocthmlol
foils hours at 37°. After incubation, 20 to 240 pg of Fraction PCI
IMO to '2“ng of Fraction PCII were layered on Sl)SspIIl.\-'il~(‘l'."l*
amide gels prepared from a 5“}; acrylamide-SIH mixture. Elec-
‘Wplloresis was performed as described in Fig. 8 (Curve .1). Fol-
lowing electrophoresis, the gels were either scanned at 280 nm
iniinctliatcly or at 550 nm after the proteins were stained with
(‘uomassie blue. The areas under the subunit peaks were meas-
ured, O—~-O, 8’ plus i3 subunit; A——~A, a subunit; .——»., 0'
Sf‘l'llllll'. Upper left, Fraction PCI, 550 nm scans; upper right,
lrm‘llm'l PCII, 550 nm scans; lower lcfl, Fraction PCI, 280 nm
scans; lower right, Fraction PCII, 280 nm scans.

The I‘nolcculur weights of the P. puli'do. subunits were deter-
illlllt'tl from a standard curve which related the log: of the I'nolccu-
l3" “'(‘iL’ht to the distance of migration from the origin (Flt-{- 3)-
Tllf' molcculzir weights of B’, B, 0', and a were estimated to be
US$000, 155,000, 98,000, and 44,000, respectively. The molccu-
ltll‘ weights of the B’ and B subunits of the P. pllt‘itfa enzyme wci'c
Jilcutiml with those of the, E. coli enzyme. The molecular
Nights of the a and a P. putida subunits were somewhat fil'i‘iltt‘l'
than those of the E. coli enzyme. (sec Fig, 7, rig/If .0”).

. ln attempts to dissociate the subunits into smullcr stI'III-turcs,
inn-tion l’(‘I was heated at 50° for 3 hours or 1000 for 10 min
mill .\1 sodium phosphate (pH 7.1), 1"; SDS, and l "I; Buttercup-
IIIIct'lmnol prior to application on a SDS-i)(ilyzit‘l‘l'l‘lml‘l“ 31“]-
l3J3, 0, and a were the only structures detected. Fraction l’i'll
“ill." (““ll'lt‘d for 15 hours in 0.1 M Tris-0.03 M citric acid 0’” H"),
1', SDS, 3 M urea, l9}, 2-mercuptoethanr)l, and 0.01"}- EDT-\-
.\ltcr dialysis, one sample was applied directly to :I SIN-1W)"
{ll‘l‘}'-ltlll'll(l0 gel containing 8 M urea, a second sample was llt‘:ll(‘(l
arm“ for 1 hour, and a third sample. was heated at 75'” for 1 hour
prior to application on the gel. [3" (3, and a were tilt" 0110' WW”
imlfk‘l detom‘d. When Fraction PCI or l’(‘ll was Ilcnuturt‘tl
“l ‘ M guunidine hydrochloride and 0.1 M Ilitliiotlircitol and 01011
. , . 01 at. 40 for 1:) hours pIIoI to (“til“l’lmlm-‘i

./ . C. Johnson, M. DeBacker, and

J . A. Boezi

TABLE II

' l‘t
’0, [0 at 81101171
Gift)" 0! [3’ 'i" [3 ("Hf 0’ 8'! _

Ibnnits rcluln
filo/(1r t‘ont'cnh‘ - [——

 

 

 

 

 

 

 

 

__._______fi~. ' l R I t' l Rclaltiw’
' Relative C 3 “"9 mo or
RNA l . m lecu :Ir . tra-
Hcthod Used in pon- Subunits j ttmgfunt waiilihi.“ “)illifnnq
quantitative determination {merase1 subunits , subuults subunits
‘ ”it
{P — “FEW-Ii ' 1 1
1 ’ I 42 13.6 '
Gel scans at 500 um 1 PCI £3 + i3 l 1 1 2.2 0.5
(after CoomaSSIe a i 1 0 1.0 I 1‘0
blue staining) , a , . ». 1.1
PCII a + ii 4.1 3-b
a 1.0 1.0 1.0
‘ ” -’ ’ .(i 1.0
»‘ ‘ . a 1 2 ‘ [‘1 ,. I)(I I d + d . 3.8 l l P
(10] scans at P5011 a l 1.1 . 2.2 ' 0.0
a 1.0 l 1 0 1.0
PCII in' + e 3.9 ‘-. :.u 1.0
l a . 1.0 l 1.0 10

|
__. ___._—_

 

ABSORBANCE AT 550 nm

 

 

 

L J I L

1
3-2 3.4
DISTANCE FROM ORIGIN

3.0 3' 6

(Cm)

F“ 1” “““r‘i'nml'ic tracing of 5’ and (3 region of SilS-bolv-
:Icrylumidc gel of Fraction PCI. Fraction p01 was dellutured
according to the procedure described in Fig 7. A gradient SDS-
P00110131amide gcl was prepared by the Procedure of )‘Izirgolis and
Kcnrick (5“ ”5“”; 3‘7 ““d 5“? atfrylztiiiitlc-SI)S mixtures. The
5"} acrylzunidc-RIH mixture was made 10"; in glycerol lustabilizc
t he acrylumide gradient during polymerization. Denatured
F'W'tim PCI (1'5 “in “"13 layered on the gradient. Sl)S~pol\-_
acrylmnide gel. l£19“1""’I’l“”“*‘l5 was performed at 8.0 nm per gel
for 4% hours at “25° in 0.1 M sodium phosphate. DH 7.1, and 0.1%
SDS. The protein was stained with Coonmssic blue. The densi-
met I'ii' “'30ng {It 5:30 nm was made with a (lilford linear transport.

only thc 0', I3, 0', and a hands of Fraction l’(‘I and the b”, B, and
or bonds of Fraction PCII were oliscrvctl.

The amount of pl’ plus t3 and of cr relative to a in Fraction PCI
and thc amount of (3’ plus 5 relative to a in Fraction PCII was
dctcrmincd from measurements of the area under each subunit
peak in dcnsimctric tracings of Sl)S-imlvucryluInidc m‘ls con-

tuininu‘ dill‘crcnt amounts of tlic “WWI-in], The arm qulcr Pilt'lt

Tm”: III
of DNA «lependcnl synthesis of RNA by
Fractions PCI and. PCI]
mtained the components de-
Assay of RNA Polymerase.”

in the reaction mixture was 9
gh-l

(.‘h (r ractc r is! tea

The complete reaction mixture Ct
scribed under “Spect rophotomet ric
The concentration of Fraction PCI
pg per ml, whereas that of Fraction PCII was ‘25 pg per ml.
DNA was used as the template.

 

 

NADPH formation in fraction

l
Components of I
reaction mixture I

 

 

 

PCI I Pcn
—# A *F— iii ——’—F — i — I mimics/mt):1m!
Complete... .. I 1.7 I 1.0
Minus RNA polymerase. . . . . _ . , . . . - (l 0
Minus gh-l DNA. . .. , ..._ I o I 0
Minus ATP, UTP, UTP, CTP. . . . , . I 0 0
Minus ATP, GTP. . . . . . . . 0 0
Minus UTP, CTP. . . . . .. . . , .I 0 0
Plus4pg/ml I)Nase....... . ‘. 0 0
Plus 4 pg/ml actimmiycin . . . . . I 0 I 0
Plus 4 ugg’nil nogalamycin. . . . . . . I 0 I 0

 

Tuna; IV
Effects of DNA templates on rates of RNA synthesis by
Fractions PCI and PCI]

All reaction mixtures, with the exception of those containing
poly d(A~T), contained the components of the reaction mixture
described under “R adioactive Assay of RNA Polymerase." CM 1’
incorporation into RNA was measured. The reaction mixture
with poly d(A—T) contained 20 find Tris acetate (pH 8.0), 1 mM
manganese acetate, 0.4 mm each of UTP and 3hI-ATF' (0 X 10“
cpm per nmole), and poly th-T) at 0.4 optical density unit at
200 not per ml of reaction mixture. AMP incorporation into poly
r(A-U) was measured. Incubation was at 30° for 10 min for all
reaction mixtures. For each template, ribonucleotide incorpora-
tion into polymer was measured at several different Concentra-
tions of Fractions PCI and PCII to make certain that the amount
of incorporation was proportional to the amount of added enzyme.

 

 

 

 

 

i
"(XWP orANlP incorporationI

I into fraction ' Activity ratio of
I

DNA template l’iaction l’t‘l to
I

‘ Fraction Pen

PCI PCII

 

 

I I
I "motes/Irvmg protein I

gh-l........... .. ,. . ... 3200 050 4.9
T... ............... .. . 580 I 130 4.5
Calf thymus ............. .l 1100 I 1200 I 0.0
Pseudomonas putt’da. . I 370 ' 200 I 1.3
Denatured gh-l ........... I 280 I 620 I 0.5
Denatured calf thymus, . . I 00 I 210 I 0.4

. I 830 I 1400 I 0 . ti

Poly (HA-T). ...

subunit peak was i‘i‘ieasured on gels which had been scanned at
550 nm after they were stained with (‘ooniassic blue or at 280 nm
for those which were not stained. ’In these analyses, [3" and t3
were not resolved because of the large amount of protein applied
to the SDS-polyacrylamidc gels. The results are. presented in
Fig. 9. By either measurement, a lin‘ar relationship for the
area under each subunit p ink with respect to the amount of pro-
tein applied to the gel was established. From the slopes of the
lines in Fig.2. 0, the amount of 8’ plus 6 and a relative to a for
Fraction l’( ‘l and the amount of [3’ plus 3 relative- in a for Frac-

1)NA-(Iependcnt RNA I’olym

Vol. 246, X0. 5)

.s-curlomonas panda

erase of 1)

I
60i— 60‘—

(nmoles/ml)

4O 40r-

20 20

INCORPORATlON

 

        

 

 

CMP

    

4.5

(min)

I5 30
TIME

45 6O 75

(mm)

Flo. 11. Kinetics of RNA synthesis by Fractions PCI and PC”.
The reaction mixtures contained the components described under
“Radioactive Assay of RNA Polymerase.” The concentration of
PCI was. 37 pg per ml and that of Fraction PCII was 38
CMP incorporation into RNA with gh-l DNA as the
template (left): O——~O, Fraction PCI; A—- ’A. Fraction PCII.
CMP incorporation into RNA with calf thymus DNA as the tem-
plate (right): I ——l, Fraction PCI; O~—-——O, Fraction l’(.‘ll.

Fraction
pg per ml.

tion l’(‘ll was determined. The results are tabulated in Table
11. There were two 5" + 1} subunits and one a subunit for every
two 0: subunits in Fraction l’(‘I and one ,B’ + ,8 subunit for each
a subunit in Fraction PCII.

The amount of 8' relative to B in Fraction PH and in Frac-
tion l’(‘lI was determined by means of (‘oomassie blue-stained
Si)S-polyacrylamide gels. The amount of protein applied on
these gels was small and ,8’ and B were resolved. For both Frac-
tions l’(‘I and PCII, the amount of B’ was equal to (3. .\ densi-
metric tracing of the B’ and 6 region of a Si)S-ptilyaci'_\'lninide
gel of Fraction l’(‘I is presented in Fig. 10.

The subunit. structural formula for Fraction PM was therefore
assigned as 02/313'0 with a molecular weight of 506,000 or an in-
tegral multiple of it. The. sedimentation coefficient of 135. indi-
cated that the molecular weight of Fraction P(‘l was about
500,000. Thus, the subunit structural formula for Fraction l’t'l
“'“S “255,0 With a molecular weight of 506,000. Based upon the
size indicated by a sedimentation coetiicient of 12 S and the
subunit content, the structural formula for Fraction P(‘ll was
(1230' and its molecular weight was 408,000.

DAV‘ILdirWICd Synthesis of RNA by Fractions PCI and I’CH

Herrera! Characteristics of RNA Synthesis~Smue of the charac-
tt‘l'ih‘tit's 0f the DNA-dependent RNA synthesis by Fractions 1’”
and l’(‘II as determined with the siiectropliotometrit‘ aw." 31“
presented in Table. III. Rifampicin (data not shown) lttlllltltt‘ll
the synthesis of RNA by Fractions PCI and PCII, with a mu-
ct-intration of 0.03 pg per ml producing a 50‘}. inhibition. Tilt"
co)icentration of the antibiotic is similar to that required to $1in
50"} inhibition of RNA synthesis by E. coli RNA it‘ih’"‘“"““"
(4i , 42).

The rate of R NA synthesis as measured by the ratcot'ilit‘t"1‘i‘*"
ration of ('Ml’ into a form insoluble in trichloracetic acid was
proportional to the amount of enzyme used. For example, wht‘“
1.5, 4.5, and 7.5 pg of Fraction I’C‘I were added to 12541] “WW”
mixtures with gh-l DNA as the template, 0.81, 2.5, and 3.‘
"m0!“ ”f (“11” l‘Osiiectively, were incorporated in 10 min.
When 1.5, 4.5, and 7.5 pg of Fraction PCII were used, 0.19, 0.5;).
and 0.72 nmole of ("‘M P, respectively, were. incorporated in l”

\\

Issue of March 10, 1971

TABLE V
Effect of RNase on DNA dependent reaction

The reaction mixtures contained the components listed under
"Spectrophotometric Assay of RNA Polymerase.” The concen-
tration of Fraction PCI was 9 pg per ml in the reaction mixtures
whicheontained gh-l DNA and 49 pg per ml in the reaction mix-
tures which contained calf thymus DNA. The concentration of
Fraction PCII was 25 pg per ml in the reaction mixtures which
contained gh-l DNA and 51 pg per ml in the reaction mixtures
which contained calf thymus DNA. RNase, which had been
heated to 100° for 10 min, was at a concentration of 4 pg per ml.

 

 

 

 

 

 

'NAnPn formation
_ Percentage of
R.\A polymerase DNA template increase _in
l Plus Minus "118 by RNase
; . RNase RNase
I __ _ . l _ __
"moles/"r in/ml
rat-non PCI gh-l 1.4 1.3 - 10
. Calf thymus 2.5 2.3 10
l
, I
Fraction PCII gh-l 2.7 1.3 110
Calf thymus 4.4 2.4 80

 

 

min. The concentrations of DNA, the four nucleoside triphos-
phates, divalent metal ions (Mg2+ and Mn“), and monovalent
cations (NH;+ or K+) which gave the fastest rate of RNA syn-
thesis by Fraction PCI were the same. as those which gave the
fastest rate of synthesis by Fraction PCII. The cmtcentratimlS
ofthc reagents were those used in the standard l‘flt‘llUttCthG assay
mixture.

fillets of DNA Templates on Rates of RNA Synthesis—JPN
rates of Fraction P(‘l- and Fraction P(.‘II-catalyzcd RNA syn-
thcsis with a number of DNA templates were determined. These
results are presented in Table IV. With P. putida bacteriophage
gh-l l).\' A or coliphage T4 DNA as template, the specific activity
of Fraction PCI (enzyme with a) was 4 to 5 times greater than
that of Fraction PCII (enzyme without a). With denatured

l).\'.\ or poly d(A-T) as template, the specific activity of Frac-
tltm PCI was half that of Fraction PCII. The kinetics of RNA
"l'ltlllt‘s‘ls by Fractions PC‘I and PCII, with gh-l DNA and calf
”Nous DNA as the template, is presented in Fig. 11. Although
(llflelring in Slope and in extent, the shapes of the curves were
-‘lllll itl'.

,Ellé'l‘f of RNase~—The rate of the DN.-‘\-directed polymeriza—
tion of the nucleoside triphosphates is increased by the addition
of RNase to the reaction mixture. This increased rate of reac-
tion, monitored by the increased rate of inorganic pyrophosphate
l‘ll‘llltllltflt, has been reported by Maitra and H urwitz (13), John—
son cl (:1. (33), and Krakow (43). The RNA polymerase used in
these published studies was not defined with respect to its com-
plement of a.

l‘lte efi'ect of RNase on the DNA-directed reaction catalyzed
lti' Fraction PCI (enzyme with a) and by Fraction PCII (enzyme
“,‘lh‘mt 0) Was investigated. The results are presented in Table
l. The initial rates of the reactions catalyzed by Fraction l’('ll
were increased 80 to 1109.}, Whereas {1,059 catalyzed by Fraction
l’('[ were increased only 10?}.

DISCUSSION

1 1W0 forms of the enzyme RNA polymerase, agBfi'O‘ ”Ml 0265',
lAllie been purified fmm P. Ptltida. (.‘lirontatogl‘fiIth 01" phos-
lttocellulose in 50% glycerol separated the enzyme into the two

J. C. Johnson, M . DeBacker, and J. A. Boezi

1231

forms, with azfifi'a (Fraction PCI) eluting at. 0.18 M KC1 and
(1236’ (Fraction PCII) eluting at 0.30 M KCl. Each form of the
enzyme was purified and characterized with respect to its subunit
composition and to certain of its physical and catalytic proper-
ties. Some preparations of 0236,0' were contaminated with a
small amount of a yellow-colored material which was removed
by sucrose or glycerol gradient centrifugation. This material
which sedimented at 25 S has been identified as an a-keto acid-
dehydrogenation enzyme complex.

The B’, 6, a, and 0: subunits of the P. putida enzyme probably
are polypeptide chains rather than aggregates of polypeptide
chains. Only these polypeptide subunits were detected by SDS-
polyacrylamide disc gel electmphoresis for enzytne that had been
denatured by a number of procedures. These procedures in-
cluded denaturation of aZBB’a and (1266' a t. elevated temperatures
in SDS, urea, or guanidine hydrochloride in the presence of the
sulfhydryl reagents 2—mercapt0ethanol or dithiothreitol. It is
not known whether the two a polypeptide chains of the P. putida
enzyme are identical. Furthermore, it is not known whether
the a of (1238,0' is a unique polypeptide chain or a mixture of
polypeptide chains, each chain having a molecular weight of
98,000. The role of E. coli 0 as an initiating factor of RNA syn-
thesis suggests that there might be more than one type of a
polypeptide chain per cell (~14).

The subunit structure of P. putida RNA polymerase is similar
to but not identical with that of the E. coli enzyme. The subunit
structure of the E. coli core enzyme is 0286’ and that of the holo-
enzyme, or complete enzyme, is a255'0 (23, 24). The molecular
weights of the a, B, B’, and 0 subunits of E. coli RNA polymerase
are 39,000, 155,000, 165,000, and 95,000 respectively. These
subunits probably are polypeptide chains rather than aggregates.
Only these subunits were observed by SDS-polyacrylamide disc
gel electrophoresis for enzyme that had been denatured by a
number of procedures (23). The molecular weights of a mixture
of E. coli [3’ and B and of a were determined by equilibrium cen-
trifugation in urea and in guanidine hydrochloride and shown to
be essentially the same as those determined by SDS~polyacrylarm
ide disc gel electrophoresis (23).

The molecular weights of the 6’ and 6 subunits of the P. putida
enzyme were identical with those of the E. coli enzyme.
measured by SDS}tolyacrylamide disc gel electrophoresis. The
molecular weights of the a and a P. putida subunits were some-
what greater than those of the E. coli enzyme. In all prepara—
tions of the E. coli enzyme, except those fractionated on hydroxv-
apatite and possibly those fractionated on phosphocellulose by
means of a shallow KC‘l gradient, a small protein, as (molecular
weight 9000), was present (23). No on has been detected in
preparations of the P. putida enzyme.

Both agBB’a and 0123,8’ of P. putida transcribed a number of
DNA templates, but with different efficiencies. With P. panda
bacteriophage gh-l DNA as the template, (XQBBIO' was 4 to 5 times
more active in RNA synthesis than was (1266’. Although differ-
ing in rate and in extent, the kinetics of gh-l DNA~directed R NA
synthesis by agBB’a and by agflfi’ is similar.

The initial rates of DNA-directed polymerization reactions
catalyzed by agflB'a and (1268’ were affected differently by RNasg
The rate of the reaction catalyzed by agfifi’a was increased 10‘},
whereas that catalyzed by agfiB' was increased 80 to 110%. The
increased rate of polymerizatitm in the presence of RNase, was
probably due to a relief of RNA product inhibition (45, 46).
Ammrently, the RNA synthetic process originated by 0383’ was

1:232

.\ pI‘oIltH't. inhibition than that. originated

l)\' aofifi’a. It is likelv that, this difference in SUS(‘(‘])t1l)lllt._\' to
product inhibition for agfiij’a and (1253’ mi» due to dItTeienu s i

the structure of the enzymes of the l)NA~eiizyine-nziseent RNA

complexes involved in RNA ehuin elongation rather than in the
vme eoiiiiilex involved

structures of the enzymes of the. DNA-enz .
in the initiation iiroeess. For E. coli l’NA ])()l_\'111£‘1’t1S0, 0 1S
released following: initiation, leaving the core enzyme (0255') to
catalyze eliuiii elongation (44). An zinnli‘igous process iirol‘mlily
occurs with the P. pulirla enzyme. Thus, for the P. pulirlu
enzyme, the emiforniation of agfifi' wliieli eatulyzes (‘lltllll elonga-
tion must depend on wl'ietlier or not a' was involved in tlie initia—

liioi‘e suseeptilile to RX

tioii [11‘0('GSS.
ltEl’lillliNC‘lCS

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45. 11mm iii: “...XN!)1\'I)NR.\I.),.\I.W'., Proc..\'(I.l..-lruil..Sit.l'.S..l..
51, #40] (19(34).

4t}. Fl.‘(‘HS, .19., 3111.1.1‘1'1‘1‘5, It. I... ZILLIG, W., .\.\'D Worn-1i. (1..

4!).

50.
:31 o

. 'lANi'oIin, (3., KAwAiiAiiA. K., AND LAI’ANJE. S..

fur. J. li’ioi'lirm.. 3, 153 (1967).

. (ii-iltsllMAN, 1.. C., .\.\'l) l)iii-;ioizN. 1)., Biophyy .l_, 9, A233)

(11min.

,]_ .tim'r.
(Vie/n. Son, 89, 72‘.) (1967).

S'i‘izixxiiz'i-z, M. A., ANI.) I)I;AI.. W. (T., JIL, Bim‘lmuix/I‘y-5v 13””
(lsiuo).

KA\\'AIHRA. K.. AND TANFoIm. C., Biorhixmixli‘y. 5, 157K (1913“?

WILLIAMS, ll. (3.. J R., .\.\'l) HAJAooIiAI.Ax, '1‘. ( 1...]. Biol. ('hrni.
241, 4951 (1906).

. LINiiiiIcIm. U., Bioi-hemixtrg, 6, 335 (1967).
51. Hms, C. H. W., Mooiiiz, S.., ANI) S'ricix, W. H., J. Biol. ('lu-W

219, (3‘23 (11156).

. MAnooms, J., .\.\'l) KicNIiiI‘K, K. (1., Anal. Bloi'llt‘lll.. 25. 347

(19138).

ARTICLE 3

PURIFICATION AND PROPERTIES OF 355-LABELED RNA

POLYMERASE OF PSEUDOMONAS PUTIDA

 

BY

J. C. Johnson and J. A. Boezi

(Manuscript to be published)

42

SUMMARY

35S-labeled Pseudomonas putida RNA polymerase
(aZBB'o) was purified from labeled cells which had been
grown on a minimal growth medium containing 35S-labeled
sulfate. As shown by SDS-polyacrylamide gel electro-
phoresis, the 35S-labeled enzyme was at least 98 per cent
pure. The specific enzymatic activity of the enzyme was
7,800 nmoles of CMP incorporated per hour per mg of pro-
tein at 30° using gh-l DNA as the template. The specific
radioactivity of the enzyme was 2.3 X 107 cpm per mg of
protein. As shown by analysis of the isolated polypeptide
subunits, the amount of 358 of 8'. 8, and a relative to d
was 3.6 to 3.6 to 2.2 to 1.0, respectively. The amount
of 8' plus B in the Initial Extract was 1.2 per cent of
the total protein as determined by SDS-polyacrylamide gel

electrophoresis.

43

INTRODUCTION

Radioactive RNA polymerase of high specific radio—
activity would be a useful reagent in certain studies of

enzyme structure and function. A radioactive assay of

3H-, 14C-, or 35S-labeled protein could be used as a

method of protein quantitation in place of the usual color-
imetric assays. The colorimetric assay of Lowry which is
the most commonly used method of protein quantitation and
one of the most sensitive can not be used in many instances
because of interference caused by such common biochemical

reagents as Tris and mercaptoethanol (l).

3H-, 14C-, and 35S-labeled Escherichia coli RNA

polymerase apoenzymes (aZBB') have been prepared (2,3,4).
Labeled E. coli holoenzyme (a288'0> or labeled RNA poly-

merase from other sources has not been prepared. The

3 14
Specific radioactivities of the H- and C-labeled E.

coli enzymes were low. Consequently, the usefulness of

the preparations was limited. This study describes the

. 35
Preparation and characterization of S-labeled Pseudomonas

' ' ' dio-
putida RNA polymerase (aZBB'o) of high SpelelC ra

activity and high specific enzymatic activity.

44

EXPERIMENTAL PROCEDURE

Unless otherwise stated, the EXPERIMENTAL PRO-
CEDURES used in these studies were the same as those

described in the preceding chapter of the thesis.

35

Materials. S-labeled sodium sulfate with

 

specific activity 845 mCi per mmole and Omnifluor were
purchased from New England Nuclear. Triton x-lOO was
from Sigma. Agarose (Bio-Gel A-l.5 m) was obtained from

Bio—Rad Laboratories.

Growth of Pseudomonas putida. The growth medium

contained the following in grams per liter: glucose, 20;

NH Cl, 2; Na HPO 6; KH PO 3; NaCl, 8; MgC12.6H20,

4 2 4’ 2 4’

0.08; NaZSO4-10H20, 0.03; Mn(C2H302)2-4H20, 0.005; CaClzy

O, 0.005; and NaZMoO4-2H20, 0.005. For
35

0.005; FeCl -6H

3 2

the production of 35S-labeled cells, 15 mCi of S-labeled

sodium sulfate were added per liter of growth medium.

The cells were grown at 33° on a gyrorotatory shaker in

2.8-1 Fernbach flasks containing 500 ml of growth medium.

The doubling time for the culture was 100 minutes. When

the cells reached the late logarithmic phase of growth,

they were harvested and stored at -20°. From a three

liter culture of 35S-labeled cells which had been harvested

at an optical density (660 nm) of 1.3 units per ml, the
Yield of cells (wet weight) was 9.7 g.

45

46

35S-labeled Proteins in polyacrylamide

Assay of
gels, Polyacrylamide gels were cut into 2 mm transverse
fractions using a stainless steel support and cutting
guide. The fractions were placed in scintillation vials.
After the addition of 0.2 ml of 30% H202 to each vial,
they were incubated at 65 to 70° for 9 hours or at 100 to
110° for 2 hours. Following incubation, 5 ml of a mix-
ture containing 6 parts of Omnifluor solution (18.1 g of
Omnifluor per gallon of toluene) and 7 parts of Triton
X-100 were added to each vial (5). The vials were capped,

shaken, and assayed for their 35S-content in a scintilla-

tion spectrometer.

RESULTS

Purification of RNA Polymerase
The purification of 35S-labeled RNA polymerase
was performed at 0 to 40°.

Initial Extract. Frozen 35S-labeled Pseudomonas

 

 

putida (9.7 g) and frozen unlabeled P. putida (11.7 g)
were mixed with washed glass beads (40 g) and ground in a
mortar with a pestle. After cell rupture had occurred,
30 ml of buffer (10 mM Tris-HCl, pH 8.0, 10 mM MgC12,
0.1 mM EDTA, 1 mM dithiothreitol, 200 mM KCl, and 15%
glycerol (v/v)) were added. The mixture was stirred
until a smooth suspension was obtained. Pancreatic DNase
I was added to a final concentration of 2.5 ug/ml. The
mixture was stirred for 15 minutes, then centrifuged at
12,000 X g for 30 minutes. The supernatant fraction was
decanted and saved. The pellet fraction was suspended in
30 ml of the above buffer and recentrifuged. The result—
ing supernatant fraction was combined with the supernatant
fraction from the first centrifugation. (Initial Extract,
63 ml.)

High Speed Supernatant Fraction. The Initial
Extract was diluted to 80 ml with the above buffer, then
centrifuged at 78,000 x g for 90 minutes. The supernatant
fraction was collected by decantation. (High Speed Super-

natant Fraction, 64 ml.)

47

48

60% Ammonium Sulfate Fraction. Solid ammonium
sulfate (23.2 g) was added to the High Speed Supernatant
Fraction to give 60% of saturation. After the mixture was
stirred for 15 minutes, the suspension was centrifuged at
12,000 X g for 15 minutes. The pellet fraction was dis—
solved in 15 m1 of the above buffer and dialyzed against
2 liters of the buffer for 15 hours. (60% Ammonium
Sulfate Fraction, 26 ml.)

DEAE-Cellulose Fraction. The 60% Ammonium Sulfate

 

Fraction was diluted 4-fold into buffer (10 mM Tris-HCl,

pH 8.0, 10 mM MgCl 0.1 mM EDTA, 0.1 mM dithiothreitol,

2:
and 15% glycerol (v/v)) and applied to a DEAR-cellulose
column (5 by 15 cm) at a flow rate of 8 ml/min. The
column was then washed with 500 ml of the above buffer
containing 0.1 M NaCl. RNA polymerase was eluted with the
above buffer containing 0.2 M NaCl. The fractions giving
the bulk of the absorbance readings at 280 nm were pooled
(230 ml). The pooled fractions were brought to 60% of
saturation with solid ammonium sulfate (83.2 9). After
the mixture was stirred for 15 minutes, the susPension was
centrifuged at 12,000 X g for 15 minutes. The pellet
fraction was dissolved in 11 m1 of phosphate buffer (20 mM
potassium phosphate, pH 7.5, 1 mM dithiothreitol, 0.1 mM
EDTA, and 15% glycerol (v/v)) containing 0.2 M KCl and
dialyzed against 2 liters of the phosphate buffer with

0.2 M ROI for 15 hours (DEAE-cellulose Fraction, 11 m1).

49

Phosphocellulose Fraction I. The DEAF-cellulose

 

Fraction was diluted to 31 ml with phosphate buffer and
applied to a phosphocellulose column (2 by 7 cm) at a flow
rate of 1 ml/min. The column was washed with 150 m1 of
phosphate buffer at a flow rate of 2 ml/min. A linear
gradient from 0.1 to 0.5 M KCl in phosphate buffer was
used to elute RNA polymerase. Fractions (2 ml) were col-
lected at a flow rate of 2 ml/min. Fractions which con-
tained RNA polymerase activity that had eluted at 0.18 M
KCl were pooled. The pooled fractions were dialyzed
against 2 liters of 20 mM potassium phosphate, pH 7.5,

1 mM dithiothreitol, 0.1 mM EDTA, 200 mM KCl and 50%
glycerol (v/v) for 5 hours and then against 2 liters of
ammonium sulfate solution (75% of saturation, pH 7.5) for
12 hours. Insoluble material was collected by centrifuga-
tion at 12,000 X g for 30 minutes, then dissolved in 10 mM
Tris acetate, pH 8.0, 0.1 mM EDTA, 0.5 mM dithiothreitol,
500 mM KCl and 7.5% glycerol (v/v). (Phosphocellulose
Fraction I, 0.83 ml.)

Agarose Fraction I. Agarose beads were suspended
in water, settled to remove fine material, and equili-
brated in 10 mM Tris acetate, pH 8.0, 0.1 mM EDTA, 0.5 mM
dithiothreitol, 500 mM KCl and 5% glycerol (v/v). A
column (1.5 by 85 cm) was packed by gravity and washed with
the above buffer at a flow rate of 0.2 ml/min. Phospho-

cellulose Fraction I was layered on the top of the column

50

and washed through at a flow rate of 0.2 ml/min. Fractions
(1.5 ml) were collected. The elution profile is presented
in Figure l. Fractions 38 through 46 were pooled, dialyzed
against 2 liters of 20 mM potassium phosphate, pH 7.5,
1 mM dithiothreitol, 0.1 mM EDTA, 200 mM KCl, and 50%
glycerol (v/v) for 5 hours, then against 2 liters of ammo-
nium sulfate solution (75% of saturation, pH 7.5) for 12
hours. Insoluble material was collected by centrifugation,
dissolved in 20 mM potassium phosphate, pH 7.5, 1 mM
dithiothreitol, 200 mM KCl, and 2% glycerol (v/v), and
dialyzed against 200 m1 of this buffer for 12 hours.
(Agarose Fraction I, 0.2 ml.)

Glycerol Gradient Fraction I. Agarose Fraction I
was layered on a 10 to 30% linear glycerol gradient (4.8
ml) prepared in 50 mM potassium phosphate, pH 7.5, 1 mM
dithiothreitol and 200 mM KCl and centrifuged in the SW 39
rotor of the Spinco model L-2 centrifuge at 35,000 rpm for
19.1 hours at 4°. After centrifugation, a hole was
punched in the bottom of the centrifuge tube and twenty

35S-content of each fraction

fractions were collected. The
was measured (Figure 2). Fractions 5 through 9 were pooled
and dialyzed against 500 ml of 50 mM Tris acetate, pH 8.0,
10 mM magnesium acetate, 1 mM dithiothreitol, 0.1 mM EDTA,
200 mM KCl, and 50% glycerol (v/v) for 8 hours. (Glycerol

Gradient Fraction I, 0.58 ml.) Glycerol Gradient Fraction

I was stored at -20° at a protein concentration of 1 mg/ml.

51

.cwaoom mHm3 we cmsounu mm mCOfiuomum .H mHoHpH< mo
=Ammmumewaom 42m mo wmmmm m>HuomoHUmmv mhspmooum Hmucmfi
Iflummxmz ca pmbflnowmp posses one ou mcflpsooom mmHsuxHE
coeuommu a: mma :H mmHQEMm Halm co pmcwfiumump mm3

A4 4V >9H>fluom wmmumE»Hom <zm .va mnemEonuommm
coHumHHHucHom cflsvfla ha mmHmEmm Hanm co pmcflfiumump
mmz AOllllov cwmpoum Umambmalmmm .pmuomaaoo mHm3
comm HE m.H mo msofluomum om can cflE\HE N.o um Ummo
|Hm>mp mmz CEDHOU was .Houmowam wm paw .Hox 2E oom
.Houamunuoaspas :5 m.o .«eom 2s H.o .o.m mm .wumumom
mane 2s as ea emumunaeesam ammn can guess season
mmoumod EU mm an m.H 6 Op GHE\HE ~.o mo mums 30am 0
pm pmflammm mm3 AHE mm.ov H cofluomum mmoHsHHmoonmmocm

 

H coeuomsm
mmoHsaamoocmmocm mo coaumuuaflm How mmoumm<II.H mmDQHm

J

 

52

(9-0l X wda) S“

O

 

 

 

 

 

 

 

 

(2,0: x mm)

N e
' l
’0’
o
o’
o/
o/ .
I .
— o
/ i
f s’
O "I
.3 </‘
d/
W
" )éo’ —
‘4
\o‘ \
l l
O O
N

NOIlVHOdHOONI dWC)

80

4O
FRACTION NUMBER

20

53

FIGURE 2.--Glycerol Gradient Centrifugation of
Agarose Fraction I

Agarose Fraction I (0.2 ml) was layered on a
10 to 30% linear glycerol gradient (4.8 ml)
prepared in 50 mM potassium phosphate, pH 7.5,
1.0 mM dithiothreitol, and 200 mM KCl and
centrifuged at 4° for 19.1 hours at 35,000 rpm
in the SW 39 rotor of the Spinco model L-2
centrifuge. A hole was punched in the bottom
of the tube and twenty fractions were collected.
A sample (2 ul) of each fraction was assayed
for 3 S by liquid scintillation spectrometry
(6). Fractions 5 through 9 were pooled.

 

355 (cpm x Io’4)

.b

0‘

N

54

 

 

 

 

 

/"
/' 1 ‘
I \ ‘
/ .\ 0..
x . fl...“ /
5 l0 I15 1'5?

FRACTION N U MBER

55

Summary and Comments on the Purification. A sum-
mary of the purification in terms of the radioactivity
recovered in each enzyme fraction of the purification

35$-

procedure is presented in Table 1. Of the total
labeled material in the Initial Extract, 0.05% was
recovered in Glycerol Gradient Fraction I. The amount of
protein of the Glycerol Gradient Fraction I was 570 pg.
As judged by sucrose gradient centrifugation (data not
shown) and by SDS—polyacrylamide gel electrophoresis (see

Figures 3, 5, 6, and 8) the 35

S-labeled RNA polymerase
(Glycerol Gradient Fraction I) was at least 98% pure.
SDS-polyacrylamide gel electrophoresis of the material of
each enzyme fraction of the purification procedure was
performed. The results are presented in Figure 3.

A densimetric tracing of a Coomassie blue stained
SDS-polyacrylamide gel of the Initial Extract is presented
in Figure 4. Polypeptide chains which have the same
mobilities as the B' and 8 subunits of RNA polymerase
were well separated from the majority of the polypeptide
chains. Since these polypeptide chains are equal in
amount and radioactive content (data not shown), and
because of their unusual size, they are considered to be
predominately if not exclusively the 8' and 8 subunits of
RNA polymerase. As calculated from the area occupied by
8' plus 8 relative to the total area of the densimetric

tracing of the SDS-polyacrylamide gel, 8' plus B amounted

to 1.2% of the total protein of the Initial Extract.

56

TABLE l.--Summary of Purification

 

 

Enzyme Fraction 35 S Remaining
(Total cpm) (%)
Initial Extract 2.83 x 1010 100.0
High Speed Supernatant Fraction 2.38 X 1010 84.2
60% Ammonium Sulfate Fraction 1.45 X 1010 51.3
DEAE-cellulose Fraction 3.20 X 109 11.3
Phosphocellulose Fraction I 1.69 X 108 0.60
Agarose Fraction I 3.60 X 107 0.13
Glycerol Gradient Fraction I 1.32 X 107 0.05

 

The 35

S content of each enzyme fraction was deter-
mined by liquid scintillation spectrometry (6).

57

FIGURE 3.--SDS-polyacry1amide Gel Electrophoresis
of Enzyme Fractions of the Purification Procedure

Samples of the various enzyme fractions were
incubated at 100° for 10 minutes in 0.1 M sodium
phosphate, pH 7.1, 1.0% SDS, 1.0% 2-mercaptoetha-
nol and 5% glycerol. The samples (10 ug) were
layered on 11 cm SDS-polyacrylamide gels pre-
pared from an acrylamide-SDS mixture containing
3.75% acrylamide and 12.5% glycerol. Electro-
phoresis was performed at 25° for 2.5 hours at

8 ma/gel in a buffer containing 0.1 M sodium
phosphate, pH 7.1, and 0.1% SDS. The protein
was stained with Coomassie blue. The gels are
from left to right: Initial Extract, High Speed
Supernatant Fraction, 60% Ammonium Sulfate Frac-
tion, DEAR-cellulose Fraction, Phosphocellulose
Fraction I, Agarose Fraction I, and Glycerol
Gradient Fraction 1.

5 8

esis
dure

 

59

.mucmwm3 smasomaoe czocx mo mpumpcmum

anew: pmcflanmump meB m3ouum map >3 pmumoflpcw munmflm3
Hmasowaoe mDOHHm> may mo mcflmco mpwummmhaom may mo Goes
lemon one .5: own an Umccmom mmz How may .mcwcflmummp
mcfl3oaaom .msan mammmfioou cufls cwcflmum mm3 cflmuoum was

.mom wH.o can .H.h mm .mumzmmocm Eswpom z H.o mcwcflmucoo

Hmmmsn m as Hmm\mE m um meson m.~ How omm um meuom
Inmm mm3 mammuozmosuomam .Houmomam wm.ma paw mpHEm
Iamuom wmh.m mcHsHmucoo musuxwfi momlmcwfimawuom cm Eoum
pwummmnm Hmm mcwfimamhomwaomnmam So HH 0 :0 command

mm3 Acflmuoum m: oav musyxfle mnu mo mamfimm alum 4
.Houmowam wm can .Hocmcumoummoumfium wo.a .mom mo.H
.H.> mm .mumnmmonm Esapom Z H.o mcwcflmucoo cowusaom

0 no a: om CH mmuscwE OH How oooa um pmumnsocfl mm3
pomuuxm HmfiuflcH mo samuoum m: ooa mcwcwmucou wnsuxflfi 4

uomnuxm HMfluHcH mo How
mpflfimamuommaomlmam am no masoche Ufluumfifimcmolu.v mmeHm

60

 

OOO'OI _)

OOO‘BI —)

 

 

 

OOO'OG —_) E

OOO'OOI ———)

OOO'OGZ ——’

ugbuo —->

 

 

um 099 lv 30stsossv

DISTANCE FROM ORIGIN

61

358 analysis of sections of this gel (Figure 5) also led

to the conclusion that 8' plus B amounted to 1.2% of the
total protein of the Initial Extract. Since 8' plus B are
63% (by weight) of the holoenzyme (0288'0), the holoenzyme
could amount to 2% of the total protein of the Initial
Extract. Accordingly, the yield of RNA polymerase in the
Glycerol Gradient Fraction I was 2.5% of the estimated
amount present in the Initial Extract.

An estimation of the percent by weight in the
Initial Extract of polypeptide chains of various molecular
weights can be made from Figures 3 and 4. The per cent of

polypeptide chains that have molecular weights greater than

25,000, 50,000, and 100,000 was 75, 31, and 9% respectively.

One-half of the polypeptide chains have molecular weights
greater than 36,000.

Characterization of 35S-labeled

RNA Polymerase
Specific enzymatic activity and specific radio-

activity of 35S-labeled RNA polymerase. The specific

enzymatic activity of 35S-labeled RNA polymerase (Glycerol

Gradient Fraction I) was 7,800 mnoles of CMP incorporated

per hour per mg of protein at 30° using gh-l DNA as the

template. With calf thymus DNA as the template, the

The specific radio-

3 x 107 cpm

Specific enzymatic activity was 3,300.

activity of 35s—labeled RNA polymerase was 2.

per mg of protein.

-—- ——-—-——-—- ——-—- —::uL—.m

62

12%

opHanxnoomHom cw mcflououm poaonmalmmm mo mommfiv
onsoooonm HoucoEHuomxme CH ponfluomoo oonuoE

onu ou mchHooom commando moz coauomnw nomo wo
uconcoo mmm one .Anooo 88 my oceauoosw mm owns
#50 mo3 v onsmflm ca pounomosm mnfloonu UHHpoEHmnop
onu mn commaoco moz nOHnS poouuxm HMflanH mo

Hom opflanwuoowaomlmom pocflmum osan owmmofioou one

vooupxm HofluwnH mo How

o©Hanmsoomaomlmom no mo mwmwamnm mmMIu.m mmame

63

 

 

OOO‘OI

OOO'QI

000'? Z

OOO'OG

OOO'OOI
OOO‘OQI
OOO'OOZ

OOO'OGZ

ugbuo

20

I
K)

I

2 ID

(”pl x mm) 592

 

50

4O

3O

20

I0

FRACTION NUMBER

64

Stability of 35S-labeled RNA polymerase. Another

preparation of 35S-labeled RNA polymerase which had been
purified through the Agarose gel filtration step was used
in the study of the stability of 35S-1abeled enzyme. This
preparation of enzyme had a specific enzymatic activity

of 8,700 nmoles of CMP incorporated per hour per mg of
protein using gh-l DNA as the template. The specific
radioactivity was 7 x 106 cpm per mg of protein. The

preparation of enzyme was stored at a protein concentra-

 

tion of 6 mg/ml at -20° in 50 mM Tris acetate, pH 8.0,
10 mM magnesium acetate, 0.1 mM EDTA, 1 mM dithiothreitol,

200 mM KCl, and 50% glycerol (v/v). After 90 days which
is slightly more than the 87.9 day half-life of 355, 80
to 90% of the enzymatic activity was retained.

. 35
SDS-polyacrylamide gel electrophoresis of S-

labeled RNA polymerase. The B', B, o and 0 subunits of

the 35S-labeled enzyme were resolved by SDS-polyacrylamide
gel electrophoresis (Figure 6). A densimetric tracing of
a Coomassie blue stained SDS-polyacrylamide gel is pre-
sented in Figure 7. As determined from measurements of

the area under each subunit peak, the amount of 8' plus B
and of a relative to a was 3.7 to 1.1 to 1.0, respectively.

The amount of 8' was equal to the amount of 8 (Figure 8).

As calculated from the relative amounts of the subunits

and their relative molecular weights, the subunit struc—

tural formula for the 35S-labeled enzyme was 0288'0.

65

FIGURE 6.--SDS-polyacrylamide Gel Electrophoresis
of Glycerol Gradient Fraction I

A mixture containing 30 09 protein of Glycerol
Gradient Fraction I was incubated at 100° for 10
minutes in 60 ul of a solution containing 0.1 M
sodium phosphate, pH 7.1, 1.0% SDS, 1.0% 2-
mercaptoethanol, and 5% glycerol. A 4-ul sample
of the mixture (2 pg protein) was layered on an
11 cm SDS-polyacrylamide gel prepared from an
acrylamide-SDS mixture containing 3.75% acrylamide.
Electrophoresis was performed at 25° for 4.75
hours at 8 ma/gel in a buffer (0.1 M sodium phos-
phate, pH 7.1, and 0.1% SDS). The protein was
stained with Coomassie blue.

H,

,.
-‘o..~w

67

.uuommcmuu

Hooded puowaflw o mcwms Ed omm um oboe mos mcfloonu
oeuuoefimcop one .oan owmmmEoou nuflz confloum wok
neonosm one .0 ossmflm on pcomoa on» Ce confluomon mm
memoHonQouuooHo Hom oUHanwuoomaomlQO en pouhaono
mo3 H coeuooum unofipmnw Honoohaw mo onEmm mnlm a

H noflnoouh ucowpouw Houoohao mo How opHE
IoH>HomwaomlQO no mo mneoone oeunofiflmcoall.h mmDUHm

68

 

 

 

 

 

 

I11“ 099 1V EONVBEIOSBV

DI STANCE FROM ORIGIN

69

FIGURE 8.--Densimetric Tracing of the B' and 8
Region of an SDS-polyacrylamide Gel of Glycerol
Gradient Fraction I

A 2-ug sample of Glycerol Gradient Fraction I
was analyzed by SDS-polyacrylamide gel electro-
phoresis according to the legend to Figure 6
except that electrophoresis was for 6 hours.
The gel was stained with Coomassie blue and
scanned using a Joyce-Loebel Microdensitometer.

 

70

 

 

 

 

Ecomm ._.< moz<mmomm<

DISTANCE FROM ORIGIN

71

Analysis of the 3SS content of sections of an SDS-

polyacrylamide gel is presented in Figure 9. The amount

of 358 of 8' plus B and a relative to a was 7.2 to 2.2 to

1.0, respectively. In other experiments, the amount of

358 of 8' relative to B was determined. As shown in three

35

different experiments, the S contents of B' and B were

equal (Table 2). Table 3 summarizes the data concerning

the 353 content of the subunits of RNA polymerase.

72

=.AmHom o©HEoH>HooeHom CH mceoeoem
poHoannmmm Ho hmmmflv ouspoooum HoecoEHHomxm=
:H poneuomop poneoE one oe manHoooo UonHoco
mm3 coHeomHH nooo Ho ecoecoo mmm one .Anomo
as my mcoHeooHe om oenH e50 wm3 m ousmem CH
poecomoem mneooue UHHeoEHmnop one an ponHoco
moz noenB H noeeooum.enoepouo HouoohHw Ho Hom
ooHEMHmuoomHomlmom pocemem oan onmmEoou one

H coHeoon ecoepouw HouooeHw Ho How
owesmHmsommHOQImom cm mo memmHmss mmm:u.m mmooem

73

 

 

mmmZDZ 20:04.1“.

 

 

 

 

1
00
(t-Ol x de) 392

 

 

 

 

74

TABLE 2.——3SS Content of 8' Relative to B

 

 

Experiment 355 in B' 358 in B 358 Content of 8'
(cpm) (cpm) Relative to B
1 800 790 1.01
2 1490 1380 V 1.08

3 1510 1650 0.92

 

75

 

 

 

o.H o.H m.o m.o a
o.H o.H m.o m.o a
w.v o.m N.~ H.H o
N.e o.m m.m m.H m
m.e o.m m.m m.H .m
ceonu opeemom meHGDQSm
useom pecansm no a ooo.ee muecsnsm mueasnsm
mom ecoecou Mom ecoecou Ho ecoecou Ho eanoz an eecsndm
Mmm o>HemHom mmm o>HeMHom mmm o>HeMHom enDOEd o>HeMHom opeemommHom
e 0e o>eeMHom o pnm .m ..m e0 enoecoo m ||.m mqmne

mm

DISCUSSION

35S-labeled Pseudomonas putida RNA polymerase

 

(0288'0) was purified from labeled cells grown on minimal
growth medium containing 35S-labeled Sulfate. The speci-
fic radioactivity of the pure enzyme was 2.3 X 107 cpm per
mg protein. As a result of this high specific radioacti-
vity, the radioactive assay used for protein quantitation
was sensitive to 0.01 pg protein. As calculated from the

35

known specific radioactivity of the S-labeled sulfate

used in the growth medium, one sulfur atom in approxi-
mately 20,000 was radioactive. If each RNA polymerase
molecule contains 400 sulfur atoms, one enzyme molecule

in every 50 contains a 358 atom.

The amount of 8' plus B in the Initial Extract was
1.2% by weight as determined by SDS-polyacrylamide gel
electrophoresis. From the percent by weight of 0288'0 in
the Initial Extract, the molecular weight of 0288'0 (5.06
105 daltons), and assuming that the Coomassie blue stain
reacts with the polypeptides of the Initial Extract equi-

valently it was calculated that each bacterial cell con-

tains approximately 5,000 molecules of aZBB'G.

This preparation of 35S-labeled RNA polymerase is

being used in the study of the release of the a subunit

76

77

from the holoenzyme (0288'0) during the RNA synthetic
process (7 and unpublished results). The results of this
study have shown that c is released following the binding
of the holoenzyme to poly(A), poly(U), or to native P.
putida DNA. The a subunit is not released following the
binding of the holoenzyme to poly[d(A-T)], native gh-l DNA;
denatured gh-l or denatured P. putida DNA. Following the
initiation of RNA synthesis on native gh-l DNA as the
template, 0 is released from the enzyme. This result
provides the first direct experimental evidence for the
in zit£g_release of O as described in the sigma cycle

proposed by Travers and Burgess (8).

REFERENCES

Lowry, O. H., Rosenbrough, N. J., Farr, A. L., and
Randall, R. J., J. Biol. Chem., 193, 265 (1951).

 

Goff, C. G., and Weber, K., Cold Spring Harbor Sym-
posia on Quantitative Biology, 35, Cold Spring
Harbor Laboratory, New York, 1970, p. 101.

Ihler, G. M., Biochim. Biophys. Acta, 213, 523 (1970).

Stevens, A., Biochem. Biophys. Res. Comm., 41, 367
(1970).

Tishler, P. V., and Epstein, C. J., Anal. Biochem.,
22, 89 (1968).

Bray, G. A., Anal. Biochem., l, 279 (1960).

Gerard, G. F., Johnson, J. C., and Boezi, J. A.,
Fed. Proceedings, (1971) in press.

Travers, A. A., and Burgess, R. R., Nature, 2 2, 537
(1969).

78

 

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