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Direct measurement of the poliovirus RNA polymerase error frequency both in vitro and in vivo

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Title:
Direct measurement of the poliovirus RNA polymerase error frequency both in vitro and in vivo
Creator:
Ward, Carol D., 1960-
Publication Date:
Language:
English
Physical Description:
vi, 89 leaves : ill. ; 29 cm.

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Subjects / Keywords:
Digestion ( jstor )
DNA ( jstor )
Gels ( jstor )
Genomes ( jstor )
In vitro fertilization ( jstor )
Nucleotides ( jstor )
Oligonucleotides ( jstor )
Poliovirus ( jstor )
Ribonucleotides ( jstor )
RNA ( jstor )
DNA-Directed RNA Polymerases -- chemistry ( mesh )
Department of Immunology and Medical Microbiology thesis Ph.D ( mesh )
Dissertations, Academic -- College of Medicine -- Department of Immunology and Medical Microbiology -- UF ( mesh )
Mutation ( mesh )
Polioviruses ( mesh )
RNA Replicase ( mesh )
RNA, Viral -- biosynthesis ( mesh )
Research ( mesh )
Ribonucleotides ( mesh )
Genre:
bibliography ( marcgt )
non-fiction ( marcgt )

Notes

Thesis:
Thesis (Ph.D.)--University of Florida, 1990.
Bibliography:
Bibliography: leaves 82-88.
General Note:
Typescript.
General Note:
Vita.
Statement of Responsibility:
by Carol D. Ward.

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University of Florida
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Copyright [name of dissertation author]. Permission granted to the University of Florida to digitize, archive and distribute this item for non-profit research and educational purposes. Any reuse of this item in excess of fair use or other copyright exemptions requires permission of the copyright holder.
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25072896 ( OCLC )
AHK5373 ( NOTIS )

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Full Text












DIRECT MEASUREMENT OF THE POLIOVIRUS RNA POLYMERASE
ERROR FREQUENCY BOTH IN VITRO AND IN VIVO



















By

CAROL D. WARD


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA


1990




DIRECT MEASUREMENT OF THE
ERROR FREQUENCY BOTH
POLIOVIRUS RNA POLYMERASE
IN VITRO AND IN VIVO
By
CAROL D. WARD
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
1990


ACKNOWLEDGMENTS
There are numerous people that I would like to acknowledge who
have helped me get through graduate school. First and foremost is my
husband, Paul Kroeger, who always encouraged me and had confidence that
I would finish graduate school if that was what I really wanted. His
belief in me was unfaltering which helped me through numerous tough
times.
I would also like to acknowledge my past and present lab coworkers
for their help and perhaps more importantly for their friendship and
their sense of humor. Thanks are also extended to the members of my
committee for their suggestions and ideas. Of course my thanks go to my
mentor, Bert Flanegan, who stuck it out with me through tough times and
always tried to be encouraging. His optimism was much needed to
counterbalance my somewhat pessimistic outlook on life.
Lastly, I would like to extend my thanks to Parker Small for his
genuine concern in both the education and well-being of graduate
students. His door was always open and his insight proved invaluable.
11


TABLE OF CONTENTS
page
ACKNOWLEDGMENTS ii
ABSTRACT v
CHAPTERS
1 INTRODUCTION 1
Background 1
Fidelity of DNA Polymerases 2
Fidelity of RNA Polymerases 4
Rapid Evolution of RNA Viruses 6
Poliovirus Structure 9
Poliovirus RNA Polymerase 11
2 METHODS 14
Enzymes 14
Radiolabeled Compounds 14
Oligoribonucleotide Primers 14
Misincorporation Assays Using 3H and 32P-labeled
Nucleotides 15
Variations on Standard Double Label Experiments 15
RNA Digestion with PI Nuclease 17
High Voltage Ionophoresis 17
Determination of Apparent Km's for Ribonucleotides 17
Cell Culture 18
Purification of Poliovirion RNA 18
Oligodeoxyribonucleotides 18
Labeling of Poliovirion RNA Using [32P]P04 19
Isolation of RNA Oligonucletides with One Internal
G from 32P-labeled vRNA 19
Poliovirus Specific Transcripts 21
Isolation of 5' End-labeled RNA Oligonucleotide
with One Internal G 21
RNase T1 Digestions 23
Gel Purification of Oligonucleotides 23
5' End-labeling 25
RNA Sequencing 25
Gel Electrophoresis 26
iii


3 DETERMINATION OF POLIOVIRUS RNA POLYMERASE ERROR
FREQUENCY IN VITRO 27
Introduction 27
Results 28
Discussion 43
4 DETERMINATION OF THE POLIOVIRUS RNA POLYMERASE ERROR
FREQUENCY IN VIVO 47
Introduction 47
Results 48
Discussion 63
5 CONCLUSIONS AND PERSPECTIVES 75
Factors Affecting Poliovirus Polymerase Error
Frequency 75
Models for DNA Polymerase Base Selection 76
Evolution Rates of Poliovirus 78
Master Sequence Theory 78
RNA Genomes vs. DNA Genomes 80
Future of RNA Viruses 81
REFERENCES 82
BIOGRAPHICAL SKETCH
89


Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy
DIRECT MEASUREMENT OF THE POLIOVIRUS RNA POLYMERASE
ERROR FREQUENCY BOTH IN VITRO AND IN VIVO
by
Carol D. Ward
December, 1990
Chairman: James B. Flanegan
Major Department: Immunology and Medical Microbiology
The error frequency of the poliovirus RNA dependent RNA polymerase
was directly measured both in vivo and fn vitro. Purified polymerase
was used to copy poly(A), poly(C), and poly(I) templates in vitro. and
the error frequency was measured by determining the amount of a
noncomplementary ribonucleotide incorporated relative to the total
amount of ribonucleotide incorporated. The Michaelis constants (Km)
were determined for each ribonucleotide and were as follows: UTP = 10
/iM, ATP 7 /iM, CTP = 6 /M, and GTP = 5 /jM. Changing the relative
concentrations of the ribonucleotide substrates had a significant effect
on the error frequency. Increasing the ratio of complementary to
noncomplementary ribonucleotide substrates from 1:1 to 10:1 resulted in
a 10-fold decrease in the error frequency. Decreasing the ratio from
1:1 to 0.1:1 had no effect. Substituting Mn+2 for Mg+2 was found to
increase the error frequency 2-fold. Changes in the template, the
reaction temperature, and the noncomplementary ribonucleotide substrate
v


used in the reaction had little effect on the error frequency.
Depending on the specific reaction condition, the error frequency varied
from 8.3 X 10"5 to 4.8 X 10'3, a change of 65-fold.
The polymerase error frequency was measured in vivo at eight
different sites in the poliovirion RNA. This assay involved the
measurement of changes in specific G residues to A, C, or U. Sites were
selected that represented both conserved and variable sequences. The
error frequencies determined in these experiments were similar to the
values determined in vitro and ranged from 9 X 10"4 to 5 X 10'3. No
correlation was observed between the error frequency and the variable
vs. conserved sites in the viral genome. Thus, the high mutation rate
that is observed at specific sites in the viral genome is apparently a
result of the high error frequency of the polymerase and the selection
for these changes at the phenotypic level.
vi


CHAPTER 1
INTRODUCTION
Background
RNA versus DNA Polymerases
The central role of both RNA and DNA polymerases is to catalyze
the accurate template-directed incorporation of NTP's or dNTP's,
respectively, in a 5'->3' direction into a growing strand of nucleic acid
in accordance with Watson-Crick base pairing. Both RNA and DNA are used
for the storage of genetic information; however, all known eukaryotic
cellular organisms use DNA for the storage of their genetic information.
DNA is much more stable than RNA and is known to be replicated with high
fidelity. One can imagine that the large genomes of eukaryotic
organisms would need to be replicated with high fidelity to ensure their
perpetuation as a homogeneous species. A small number of errors may be
permitted to promote species evolution, but accuracy is important to
produce viable genome copies. Rates of evolution of cellular genes
average 10'9 substitutions per site per year, in part due to elaborate
proofreading and repair mechanisms (Li et al., 1985; Britlen, 1986). On
the other hand, viral RNA genomes are an average of only 3 to 30
kilobases in length, and RNA viruses are known to evolve at rates a
million-fold higher than their DNA hosts (Holland et al., 1982; Gojobori
and Yokoyama, 1985; Barrel, 1971). Such small genomes probably cannot
afford to invest the space needed for such elaborate repair mechanisms
1


2
and it may be more advantageous for them to be able to evolve rapidly.
As all viruses are intracellular parasites, variability is probably of
utmost importance.
Fidelity of DNA Polymerases
In Vivo Error Rates of Animal Cell DNA Polymerases
The in vivo error rates of animal cell DNA polymerases average 10'8
- 10'11 per incorporated nucleotide (Fowler et al., 1974; Drake, 1990).
This is in sharp contrast to their Tn vitro error rates which vary from
10'4 105 (Kunkel et al. 1981; Kunkel and Loeb, 1981). Many models
have been proposed to explain the kinetic mechanism by which DNA
polymerases achieve their high fidelity. All of these models involve
selection of the correct dNTP and the removal of misincorporated
nucleotides by a 3'-+5' exonuclease. Presumably, one reason for the high
error rate of these polymerases in vitro is due to the lack of any
associated exonuclease activity with the purified polymerases.
Kinetic Mechanism of Escherichia coli DNA Polymerase I
Recently much work has been done to determine the kinetic
mechanism by which DNA polymerase I from Jh_ coli achieves its high
fidelity (Kuchta et al., 1987, 1988). A three-step mechanism has been
proposed. The first step involves nucleotide discrimination from a
reduced rate of phosphodiester bond formation for incorrect nucleotides,
with a small contribution from selective dNTP binding. This allows a
discrimination level of approximately 104 106-fold. The second step
involves a slow dissociation of the incorrect DNA from the polymerase
which in conjunction with the 3'->5' exonuclease allows a discrimination
level of approximately 4 60-fold. The last step involves the slow


3
polymerization of the next correct dNTP onto the mismatch which again
allows for correction by the 3'-+5' exonuclease. This contributes to the
fidelity another 6 340-fold. Taken all together, the error rate of
the polymerase would be in the right range for what has been measured in
vivo. Also the lack of any associated exonuclease activity in vitro
would explain its higher error rate.
Factors Affectine Fidelity of DNA Polymerases
The difference in free energy between correct and incorrect base
pairings is estimated to be from one to three kcal/mol (Loeb and Kunkel,
1982). This would predict an error frequency of approximately 1 per 100
nucleotides for nonenzymatic polymerization of oligonucleotides.
Obviously DNA polymerases must enhance this fidelity even in the absence
of 3'-5' exonucleases. It has been seen that the error rates of
eukaryotic DNA polymerases are directly proportional to the ratio of
correct to incorrect nucleotide substrates (Seal et al., 1979). It is
likely, therefore, that fluctuations in nucleotide pools in vivo would
increase the error frequency of the polymerase. Metal ions have also
been seen to increase the error frequency of DNA polymerases (Sirover
and Loeb, 1977). The ions Mn2+, Co2+, Ni2+, Zn2+, and Be2+ all increase
the error frequency of DNA polymerases, although presumably in
different ways (Sirover and Loeb, 1976). While some can form a complex
with different DNA polymerases, changes in error frequencies at low Mn2+
concentrations (less than 100 M) correspond to changes in the binding
of the template and not the enzyme (Beckman et al., 1985). Sirover and
Loeb also report a positive correlation between metal ions that increase


4
the error frequency of DNA polymerases and those reported to be mutagens
in vivo (Sirover and Loeb, 1976).
Fidelity of RNA Polymerases
Measurements of RNA Polymerase Error Frequencies
Little work has been done to measure the error frequencies of RNA
polymerases. This is largely due to the lack of purified RNA
polymerases that can be used in an n vitro system. The error frequency
for vesicular stomatitis virus (VSV) polymerase has been measured in
vivo and in vitro and ranged from 1 to 4 X 10'4 (Steinhauer and Holland,
1986). These measurements were made by direct determination of the
level of substituted bases at given positions in the viral genome by
using a specific ribonuclease, RNase T1. For the 11 kilobase genome of
VSV, this would mean that every member of a plaque purified population
would differ from other genomes of that same plaque at a number of
different nucleotide positions if there was no selective pressure. With
an error frequency of 1 X 1CT4 and an 11 kilobase genome, the average
genome would have 1.1 mistakes, with 33% of the population having no
mistakes. They concluded, therefore, that the preservation of a
consensus sequence must be due to a strong biological selection for the
most fit and competitive representatives of the population.
Terminology
One must be careful when talking about fidelity of polymerases to
distinguish between evolutionary rates and error frequencies.
Evolutionary rates measure the number of mutations over time which
become fixed or dominant. Error frequencies, or mutation rates, are the
frequency of a mutation event which would be represented by the


5
misincorporation during a single round of RNA replication. Most studies
only measure viable mutation rates. Mutant frequency is the proportion
of a certain mutant appearing in an RNA population. Therefore, high
error frequencies or mutation rates are not always reflected in high
evolutionary rates. This is largely dependent upon the environmental
conditions under which a virus is replicating. Often there is a
predominating wild type sequence because the variants have no
competitive advantage over the wild type sequence. This is clearly seen
with poliovirus and other RNA viruses. While poliovirus is seen to
undergo rapid changes while replicating in humans (Kew et al., 1981;
Minor et al., 1986), it is remarkably stable in tissue culture (Parvin
et al., 1986). VSV has been seen to undergo changes during dilute
passages in tissue culture, only to revert back to wild type in later
passages (Spindler et al., 1982). Influenza virus, on the other hand,
is seen to undergo rapid changes in vitro even in conditions under which
poliovirus and VSV are seen to remain stable (Brand and Palese, 1980;
Parvin et al., 1986).
Reversion Rate of Point Mutations
The reversion rate of a point mutation is actually the measure of
a mutant frequency or the frequency of a substitution at a particular
position. A mutant frequency may reflect the mutation rate if two
conditions are met. First, the mutation must be essentially neutral
under permissive conditions, and, second, the amplification of the
mutant is completely wiped out under nonpermissive conditions.
Obviously these conditions can only be met for conditionally lethal
mutants. The reversion rate for an extracistronic point mutant of


6
bacteriophage Q£ has been calculated to be about 10"4 (Batschelet et al.,
1976; Domingo et al., 1976). While this is not a conditionally lethal
mutant, the authors were able to calculate a selective value of the
mutant compared to wild type and used this in calculating their
reversion rate.
Fidelity of Reverse Transcriptases
Retrovirus RNA dependent DNA polymerases have been purified and
used in vitro to determine their error frequencies which are quite high.
Misincorporation rates on homopolymeric templates have shown error
frequencies in the range of 10'3 to 104 (Battula and Loeb, 1974; Sirover
and Loeb, 1977; Loeb and Kunkel, 1982; Preston et al., 1988; Roberts et
al., 1988; Takeuchi et al., 1988). The error frequencies have also been
measured in vitro on heteropolymeric DNA sequences. It was seen that
the accuracy of the reverse transcriptase was dependent on the sequence
replicated and was in the range of 103 to 10"4 (Richetti and Buc, 1990).
Similarly, the frequency of point mutations in retroviruses have been
seen to arise at the same rate (Gopinathan et al., 1979; Kunkel et al.,
1981; Loeb and Kunkel, 1982; Preston et al., 1988). Reverse
transcriptases are DNA polymerases; however, there is no known
exonuclease activity associated with them and they copy RNA templates.
Retroviruses are also seen to exhibit high evolutionary rates, similar
to that of RNA viruses (Coffin et al., 1980; Gojobori and Yokoyama,
1985).
Rapid Evolution of RNA Viruses
Measurement of RNA Virus Evolution Rates
Evolution rates are defined as the rate at which viable mutations
accumulate in the genome. It has been seen that the rate of evolution


7
of RNA genomes is much higher than that seen with DNA genomes. Numerous
methods have been used to measure the evolution rates of RNA viruses.
Common methods include the rates of mutation to monoclonal antibody
resistance, measurement of changes in RNase T1 oligonucleotide maps,
drug resistance mutation rates, reversion rates of point mutations, and
direct RNA sequencing. Obviously, only viable mutant frequencies and
mutation rates are measured using these techniques.
Poliovirus Evolution Rates as Measured by Monoclonal Antibody Resistance
Poliovirus variants resistant to monoclonal antibody
neutralization have been seen to arise at the rate of 10"4 to 10"5 for
Mahoney type 1 (Emini et al., 1984a, 1984b). Similar measurements have
been made for Leon type 3 virus and the attenuated Sabin type 3 vaccine
strain derived from it (Minor et al., 1983). These variants arose at
the rate of 104 to 105 for the Sabin strain, whereas mutants from the
Leon strain arose 10 times more frequently. Most of these mutations
appear to be point mutations in the capsid proteins.
Poliovirus Evolution Rates as Measured by T1 Oligonucleotide Mapping
Poliovirus evolution rates have also been measured numerous times
by RNase T1 oligonucleotide mapping. In one study clinical isolates
were followed during a 13-month epidemic (Nottay et al., 1981). These
isolates showed continual mutation and selection during replication in
humans which resulted in the fixation of about 100 nucleotide changes,
or 1 to 2% of the genome bases. Similarly, changes in T1
oligonucleotide maps have been measured in vaccine-associated cases of
paralytic poliomyelitis (Kew et al., 1981). Most isolates appeared to
be multisite mutants which ranged from less than 10 nucleotide changes


8
to greater than 100 nucleotide changes. These changes again represent
up to 1 to 2% of the genome bases, presumably during replication in only
one or two people compared to the previously mentioned 13-month epidemic
of wild type virus.
Poliovirus Evolution Rates as Measured by Guanidine Resistance Mutations
Poliovirus mutants' resistance to guanidine have been seen to
arise at the rate of approximately 3 X 105 (Holland et al., 1973). This
may, however, represent changes at more than one site on the genome thus
underestimating the rate of a single mutation. Recently measurements
were made for the conversion of a guanidine-dependent poliovirus to
guanidine resistance (de la Torre et al., 1990). This represents a
single site reversion and was found occur at a mutation frequency of 2.5
X 10'3 to 2 X 10'4.
Poliovirus Evolution Rates as Measured by Point Mutation Reversions
Much controversy revolves around the mutation rates measured for
poliovirus by point mutation reversions. The previously mentioned point
mutation reversion of guanidine dependence to guanidine resistance
showed a high mutation frequency of 103 to 104. Another study involved
inserting 72 nucleotides into the 5' noncoding region of poliovirus
which contained an in-frame start codon (Kuge et al., 1989). This
mutant had a small plaque phenotype and reversion to large plaques was
seen to have single nucleotide changes. These revertants were seen to
arise at the rate of 102. This high number was explained by the high
viability of the large plaque phenotype mutant to the small plaque
insertion mutants and the existence of sister clones in the small
plaques. A third study argues that a single base revertant of


9
poliovirus arises at the low rate of 2.5 X 10'6 (Sedivy et al., 1987).
In this study a mutant was constructed such that a serine codon was
converted to an amber codon. The virus was then grown on suppressor
positive cells and then titered on both suppressor positive and
suppressor negative cells to determine the reversion frequency. Only
one of three possible nucleotide changes was seen to generate revertants
so the mutation rate was estimated to be approximately 105. It should
be noted, however, that the mutant virus had a 2-fold longer eclipse
period and a 10-fold lower burst size on the suppressor cell line,
probably due to incomplete suppression. Obviously this mutation was not
neutral in permissive conditions and thus is not a direct measurement of
the error frequency of the polymerase.
Poliovirus Error Frequency as Determined by Direct Sequencing
Parvin et al. sequenced a segment of the viral protein 1 (VP1)
gene of poliovirus type 1 from multiple individual virus plaques that
had all descended from a single plaque (Parvin et al., 1986). No
mutations were detected in over 95 X 103 nucleotides sequenced. A
neutral mutation rate of less than 2.1 X 106 was calculated for this
site compared to 1.5 X 10'5 for the NS gene of influenza virus. Lethal
mutations would not have been scored in this study. This low rate could
be due to the need for conservation at VP1, a lower polymerase error
frequency at this site, or due to purifying selection differences.
Poliovirus Structure
RNA Genome and Viral Proteins
The genomic RNA of poliovirus has a 3' terminal poly(A) sequence
and a 5' covalently linked protein called VPg. For the Mahoney strain


10
of type 1 poliovirus, the viral RNA (vRNA) is about 7500 bases in
length and contains one long open reading frame from base 743 to 7370.
The exact functions of the 5' and 3' noncoding regions are not known,
but they may be involved in replicase recognition and binding to
initiate RNA synthesis. The 5' noncoding region also contains sequences
that are required for ribosome binding and the initiation of
translation. The four capsid protein sequences are located at the amino
terminus of the polyprotein. The capsid proteins VP1, VP2, and VP3 have
a common structural motif (Hogle et al., 1985, 1987; Hogle and Filman,
1989) They contain a core sequence which is composed of an eight-
stranded antiparallel beta barrel with two flanking alpha helices.
Differences in the amino acid sequences of the capsid proteins of type 1
Mahoney and type 3 Leon strains have been mapped to the inner and outer
surfaces of the capsid proteins, but not to the core sequences. This
suggests a strong selective pressure for constraints imposed by protein
folding and assembly. Two proteases, 2A and 3C, are also encoded in the
long open reading frame (Hanecak et al., 1982; Toyoda et al., 1986).
These proteases are involved in the cleavage of the polyprotein into the
smaller viral gene products, with most of the cleavages performed by 3C
(Ypma-Wong et al., 1988). Other viral protein products include 3AB, a
membrane associated precursor of VPg, VPg(3B) and 3D, the viral RNA
polymerase. The roles of nonstructural viral proteins 2B and 2C in
poliovirus replication are unclear at this time. Guanidine
hydrochloride is known to block poliovirus replication. It appears that
2C is responsible for guanidine sensitivity (Anderson-Sillman et al.,
1984; Baltera and Tershak, 1989). Guanidine resistance and dependence,


11
as well as host range mutants, have also been mapped to 2C (Yin and
Lomax, 1983; Anderson-Sillman et al., 1984; Pincus and Wimmer, 1986;
Pincus et al., 1986; Baltera and Tershak, 1989; de la Torre et al.,
1990). These data suggest that 2C has a role in RNA synthesis.
Homology Between Serotypes
The homology between the three serotypes of poliovirus is 71% at
the nucleotide level (Toyoda et al., 1984). Of these substitutions, 80%
of them are silent. It is interesting that the type 3 Sabin vaccine
P3/Leon/12ab differs from its neurovirulent parent P3/Leon/37 by only 10
point mutations (Stanway et al., 1984; Almond et al., 1987a, 1987b,
Westrop et al., 1989). The recent poliomyelitis outbreak in Finland was
from a type 3 virus that had 95.5% homology with P3/Leon/37 at the amino
acid level (Hughes et al., 1986). There were 3 amino acid substitutions
and 6 amino acid substitutions, however, at two major antigenic
determining sites. These sites are normally highly conserved in wild
strains of poliovirus.
Poliovirus RNA Polymerase
Poliovirus RNA Polymerase and its Role in Replication
Poliovirus RNA replicates in the cytoplasm of infected cells using
a virus-coded RNA-dependent RNA polymerase. A soluble and template-
dependent form of the poliovirus polymerase, 30^ has been purified
from cytoplasmic extracts of infected cells (Van Dyke and Flanegan,
1980). Highly purified forms of the polymerase synthesize full-length
copies of poliovirion RNA and other polyadenylated RNAs, but only in the
presence of an oligo(U) primer (Flanegan and Baltimore, 1977; Flanegan
and Van Dyke, 1979; Tuschall et al., 1982; Van Dyke et al., 1982). The


12
requirement for a primer is unusual for RNA polymerases, but this need
can be eliminated by the addition of a cellular protein component termed
"host factor" in vitro (Dasgupta et al., 1980; Flanegan et al., 1987).
In vitro reactions require all four ribonucleotide triphosphates, Mg2+
or Mn2+, and an oligo(U) primer or host factor. In the presence of host
factor, the largest size of product RNA synthesized is twice the size of
the RNA template (Young et al., 1985). The product RNA is complementary
to and covalently linked to the template RNA (Young et al., 1985, 1986).
The amount, size distribution, and rate of synthesis of product RNA are
dependent of the Mg concentration, pH, and temperature of the in vitro
reaction conditions (Van Dyke et al., 1982). At optimal in vitro
conditions the synthesis rate is approximately 1200 nucleotides per
minute.
Replication of Homopolvmers
Synthetic homopolymers can be copied by the polio polymerase in
vitro in the presence of the correct primer. These template primers
include poly(A):oligo(U), poly(C):oligo(I), and poly(I):oligo(C)
(Tuschall et al.,1982). Poly(U):oligo(A) can only be copied to a very
small extent and no activity is noted on poly(G):oligo(C). Template
binding studies indicate that the polymerase binds to poly(G) the best,
followed by poly(U), poly(C), poly(I), and lastly poly(A), the exact
opposite order of the templates that it copies best (Oberste and
Flanegan, 1988) .
Studies on the Fidelity of Poliovirus Polymerase
Studying the fidelity of poliovirus RNA replication has a unique
advantage over studies with most other RNA animal viruses because the


13
polymerase has been purified in a soluble and template-dependent form
and can therefore be used to directly measure error frequencies in vitro
on different RNA templates. By copying synthetic homopolymers using the
purified polymerase and differentially labeled complementary and
noncomplementary ribonucleotide substrates, one can directly measure the
error frequency of the polymerase. This procedure eliminates the bias
of converting viable mutation rates to error rates of the polymerase
since lethal mutations need not be accounted for. The in vitro
procedure also has the distinct advantage of allowing one to vary the
reaction conditions and observe any changes in the error frequency.
Templates may also be varied to determine if certain base pair
mismatches are allowed more than others.


CHAPTER 2
METHODS
Enzymes
Proteinase K was obtained from Boehringer Mannheim in lyophilized
form. It was dissolved and stored in 10 mM Tris*HCl, 1 mM EDTA, and 25%
glycerol at a concentration of 10 mg/ml. Ribonuclease T1 was obtained
from Calbiochem Corporation in lyophilized form. It was dissolved and
stored in 50 mM Tris at 10 u/pl.
Radiolabeled Compounds
All radiolabeled compounds were obtained from Amersham except for
[32P]P04. This compound was obtained from either ICN Biomedicals, Inc.,
or New England Nuclear. The radiolabeled nucleotide [y-32P]ATP was
obtained in aqueous solution with a specific activity of 3000 Ci/mmole.
All other radiolabeled compounds were in a 50% ethanol solution which
was vacuumed down to less than half volume before using. The
radiolabeled compounds [a-32P]UTP, [q-32P]CTP, [q-32P]ATP, and [a-32P]GTP
all had a specific activity of 410 Ci/mmole. The specific activity of
[3H]UTP ranged from 37 to 45 Ci/mmole, [3H]CTP was 20 Ci/mmole, and
[3H]GTP ranged from 10 to 15 Ci/mmole.
Oligoribonucleotides Primers
Oligoribonucleotides were generated from homopolymers as described
by Bock. Briefly, 1 2 mg of homopolymer (poly(C), poly(I), or
poly(U)) was incubated at 90C for 40 minutes in 0.5 ml 0.1 M
14


15
NH4HC02 NH40H (pH 10.0). The pH was then adjusted to 1.0 by the addition
of 0.5 ml 1 M HC1 and incubated at 20C for 20 minutes. This solution
was then neutralized by the addition of 1 ml of 1 M Tris*HCl, pH 8, and
ethanol precipitated. The oligonucleotides were then treated with calf
intestinal phosphatase (Boehringer Mannheim) as described (Maniatis et
al., 1982).
Misincorporation Assays Using 3H and 32P-labeled Nucleotides
The standard reaction mixture (final volume 30 ¡j.1) contained 50 mM
HEPES (N-2-hydroxyethyl piperazine-N2 ethane sulfonic acid) pH 8.0, 3
mM MgCl2, 10 mM DTT, 2.5 g poly(A) 1.25 /g oligo(U), and 3 /il Fraction
IV(HA) polymerase purified as described (Young et al.,1986). For the
correct ribonucleotide, 10 fid of [5,6-3H]UTP (36 Ci/mmole) was added,
and for the incorrect ribonucleotide 20 /Ci of either [q-32P]ATP, [a-
32P]GTP, or [a-32P]CTP was added. Unlabeled ribonucleotides (Calbiochem
Corporation) were added to make the reaction mixtures equimolar (7.2 yuM)
with respect to complementary and noncomplementary ribonucleotides. The
reactions were incubated for 1 h at 30C. The labeled product RNA was
precipitated in 7% trichloroacetic acid (TCA) and 1% sodium
pyrophosphate, collected on membrane filters, and counted.
Variations on Standard Double Label Experiments
Template
Variations of the misincorporation assay included substituting
poly(C):oligo(I) for the template:primer and using [3H]GTP as the
correct ribonucleotide or substituting poly(I):oligo(C) for the template
and using [3H]CTP as the correct ribonucleotide.


16
Concentration of Correct and Incorrect Ribonucleotides
All variations on the ratio of correct to incorrect
ribonucleotides were done on a poly(A) template with [32P]CTP used as
the incorrect ribonucleotide. At a 10:1 ratio of correct to incorrect
ribonucleotides, 50 /Ci of [3H]UTP was used and unlabeled UTP was added
to a final concentration of 74 /M for the correct ribonucleotide. For
the incorrect ribonucleotide, 20 /Ci of [32P]CTP was used and unlabeled
CTP was added to a final concentration of 7.4 /M. Alternatively, 10 /Ci
of [3H]UTP was used and 10 /tCi of [32P]CTP was used making the
concentrations of correct and incorrect ribonucleotides 7.4 /M and 0.74
/M respectively. At 0.1:1 ratio of correct to incorrect
ribonucleotides, 1 /Ci of [3H]UTP was used as the correct ribonucleotide
and 20 /Ci of [32P]CTP was used and unlabeled CTP was added to a final
concentration of 7.4 /M.
Mg+2 versus Mn+2
Another variation including substituting 0.5 mM MnCl2 for 3 mM
MgCl2. This was done under both equimolar concentration of correct to
incorrect ribonucleotides and at 10:1 ratio of correct to incorrect
ribonucleotides with the correct ribonucleotide being at 7.4 /M and the
incorrect ribonucleotide being at 0.74 /M.
Temperature
The error frequency was determined at 30C, 37C, and at 42C.
This was done at equimolar concentrations and at a 10:1 ratio of correct
to incorrect ribonucleotides as described above.


17
RNA Digestion with PI Nuclease
Product RNA was synthesized on a poly(A) template as described
above using 10 tCi each of [a-32P]UTP and 10 tCi of either [a-32P]ATP,
[a-32P]GTP, or [a-32P]CTP. The labeled product RNA was phenol:chloroform
extracted, ethanol precipitated, dissolved in 100 1 of 0.1 M NaCl, 1 mM
EDTA, 10 mM Tris HC1, pH 7.6, and run through a G-50 "spun column" as
described (Maniatis et al., 1982) to remove unincorporated labeled
nucleotides. The labeled RNA was ethanol precipitated and dissolved in
15 ti 10 mM CH3C00Na* 3H20, pH 6.0 containing 1.5 units of PI nuclease
(Bethesda Research Laboratory). The sample was then heat denatured at
100C for 10 minutes, cooled to 37C, and another 1.5 units of PI
nuclease in 15 tl of 10 mM CH3C00Na*3H20, pH 6.0 was added to the
sample. The sample was incubated for 1.5 h at 37C to complete the
digestion.
High Voltage Ionophoresis
Ionophoretic separation of the PI nuclease digested product RNA
was performed on Whatman 3MM paper at pH 3.5 as described (Barrel, 1971;
Rose, 1975). The 32P-labeled ribonucleoside monophosphates were located
by autoradiography, cut out from the paper and counted in 5 ml of
Aquasol-2 scintillation fluid.
Determination of Apparent Km's for Ribonucleotides
Poliovirion RNA (1 g) was copied in the presence of 10 tCi of
[32P]UTP and enough unlabeled UTP to make the final concentrations 0.8
iM, 5 tM, 10 tM, 20 tM, 40 tM, or 80 tM. The other three
ribonucleotides were also present at 500 tM. Standard reaction
conditions were used (3 mM MgCl2, 50 mM HEPES pH 8.0) except that the


18
total volume was 150 ¡i\ and 30 n 1 aliquots were taken every 10 minutes
for 30 minutes and TCA precipitated. Moles of product made were then
calculated and divided by time to determine the initial velocities. The
initial velocities and substrate concentration were then plotted on a
Lineweaver-Burk double reciprocal plot to determine the apparent Km for
UTP. The apparent Km's for the other three ribonucleotides were
determined in a similar manner.
Cell Culture
HeLa cells were maintained in suspension culture and infected with
poliovirus type 1 (Mahoney strain) as previously described
(Villa-Komaroff et al., 1974). Briefly, cells were washed with Earles
saline, pelleted, and infected with poliovirus at an MOI = 20 for 30 min
at room temperature. The cells were then diluted to 4 X 106 cells/ml in
Eagles modified minimal media with 7% sera (5% bovine calf and 2% fetal
calf). Cells were infected for 6 h at 37C, washed with Earles saline,
and frozen at -20C.
Purification of Poliovirion RNA
Poliovirion RNA (vRNA) was isolated from infected cells as
described (Young et al., 1986). Briefly, poliovirions were banded in
cesium chloride density gradients, phenol extracted three times,
chloroform extracted three times, and the vRNA ethanol precipitated.
01igodeoxvribonucleotides
All oligodeoxyribonucleotides were synthesized on an Applied
Biosystems model 380A or 380B automated DNA synthesizer, using
phosporamadite chemistry. These were then gel purified on a 20%
polyacrylamide, 7 M urea gel. A list of the oligodeoxyribonucleotides


19
used are shown in Table 2-1. These were all complementary to
poliovirion RNA at the nucleotides shown in parentheses and contained a
3' terminal poly(A)15 sequence.
Labeling of Poliovirion RNA Using F32P1PO,
HeLa cells growing in suspension culture were centrifuged and
washed three times with phosphate-free modified Eagles media buffered at
pH 7.2 with 25 mM HEPES and 10 mM TES (N-tris(hydroxymethyl)methyl-2-
aminoethane sulfonic acid). The cells were infected with poliovirus
(MOI = 20) and allowed to incubate for 30 minutes at room temperature
(approximately 25C). The cells were then diluted to 4 X 106 cells/ml
in phosphate free media with 7.0% sera (5% bovine calf sera and 2% fetal
calf sera) and dialyzed against phosphate-free Earles balanced salt
solution (116 mM NaCl, 5 mM KC1, 26 mM NaHC03, pH 7.5). After 15 min,
Actinomycin D was added at a concentration of 5 /ig/ml. At 30 min 600
/uCi/ml [32P]P04 was added. At 6 h the cells were centrifuged and washed
with Earles saline. The RNA was then purified as described (Young et
al., 1986).
Isolation of RNA Oligonucleotides with One Internal G from 32P-labeled vRNA
Poliovirion RNA (10 ig) uniformly labeled with [32p] P04 was
hybridized with 0.3 ng of DNA oligonucleotide in 30 nl of 10 mM Tris
HC1, pH 7.5, 500 mM NaCl, and 4 mM MgCl2, for 3 h at 50C. The hybrids
were digested with 6 units of RNase T1 for 2 h at 50C. The RNA was
treated with proteinase K by diluting the sample with 120 /I 0.5% SDS
buffer (100 mM NaCl, 10 mM Tris, pH 7.5, 1 mM EDTA, .5% SDS) and adding
75 ng proteinase K for 1 hour, phenol:chloroform extracted two times and
ethanol precipitated. The RNA:DNA duplex was then purified on a 20%


20
Table 2-1. List of DNA OliEonucleotides and Protected RNA Sequences
Name vRNA bases
DNA seauence 5'-*3'/RNA seauence 3'-+5'
BF8 6862 6895
CAAGTTGTTAATCATTGAGTTAAAAATTGAAGTGAAAAAAAAAAAAAAA
GUU CAACAAUUAGUAACU CAAUUUUUAACUU CAC
BF9 7269 7296
CTGATTTTAGCTAGGAATTTGTTATATTAAAAAAAAAAAAAAA
GACUAAAAUCGAUCCUUAAACAAUAUAA
BF10 5638 5671
CTTTAGAGTGATTATAGTGATTTCAAGATTGGTTAAAAAAAAAAAAAAA
GAAAUCUCACUAAUAU CACUAAAGUUCUAAC CAA
BF11 3971 3999
CTAGTTATAATAACTAGTGAGGATATGATAAAAAAAAAAAAAAA
GAU CAAUAUUAUUGAU CACUC CUAUACUA
BF25 84 109
CTAAGTTACGGGAAGGGAGTATAAAAAAAAAAAAAAAAAAA
GAUUCAAUGCCCUUCCCUCAUAUUUU
BF27 1229-1253
CTAGGTAGTGGTAGTACATATTTTGAAAAAAAAAAAAAAA
GAUCCAUCACCAUCAUGUAUAAAAC
BF30 1429-1450
CTGGTTGTTGTCAGGAGTGAAAAAAAAAAAAAAAAA
GACCAACAACAGUCCUCACUU
BF35 2757-2776
CGTGGTGGAAGCTGGGTTATAAAAAAAAAAAAAAA
GCACCACCUUCGACCCAAUA


21
polyacrylamide gel, denatured, and the RNA oligonucletide was
thenrepurifled on a 20% polyacrylamide, 7M urea gel. The RNA fragment
was then RNase T1 digested for 1 hour at 50C and the digestion products
were run on a 20% polyacrylamide, 7M urea gel. The radioactivity (cpm)
left in the RNase T1 resistant RNA fragment was then compared to the cpm
in the digestion products to determine the error frequency using an
automated gel scanner (AMBIS or Betagen). This procedure is shown
schematically in Figure 2-1.
Poliovirus Specific Transcripts
DNA plasmids pOF2612 and pOF1205 were used to generate labeled
poliovirus specific RNA transcripts. The plasmid pOF2612 contains a
full-length cDNA copy of the poliovirus genome and pOF1205 contains the
3' terminal nucleotides 6516 7440 and a poly(A)g3 sequence (Oberste,
1988a). The plasmid p0F2612 was digested with Pvu II which cuts at base
7053 in the poliovirus sequence and pOF1205 was cut with EcoRl which
cuts at the 3' end of poly(A)83. These templates were then transcribed
with SP6 polymerase and 10 pCi of [32P]UTP or [32P]ATP following the
protocol supplied by Promega Biotechnologies Inc. The RNA was DNase
treated, followed by phenol:chloroform extraction and ethanol
precipitation. The amount of RNA transcribed was calculated by TCA
precipitation and counting of a small aliquot of the sample. These
transcripts were then used in hybridization and T1 digestion experiments
as described above for [32P]P04 labeled virion RNA.
Isolation of 5' End-labeled RNA Oligonucleotide with One Internal G
Poliovirion RNA (10 /.ig) was hybridized and digested as described
above using 0.03 /ig 5'end-labeled DNA oligonucleotide and 0.27 /ig cold


22
Labeled vRNA
> DNA oligo Poly(A)
Hybridize
T1 Digest
20% Acrylamide Native Gel
A
Isolate RNA-DNA Hybrid
Denature
20% Acrylamide 7M Urea Gel
^ DNA oligo
Poly(A)
Protected RNA oligo
Purify protected RNA oligo
T1 Digest
20% Acrylamide 7M Urea Gel
T1 Resistant Oligo
T1 Digestion Products
Fig. 2-1. Schematic diagram of [32P]P04 labeled vRNA hybridization and
RNase T1 digestion assay to determine error frequency of the polymerase
at specific sites on the genome.


23
DNA oligonucleotide. The hybrid was purified on a nondenaturing 20%
polyacrylamide gel and located by autoradiography. The protected RNA
fragment was 5' end-labeled as described below, DNased with 2 units
DNase (Boehringer Mannheim) in 50 m1 of 100 mM CH3C00Na* 3H20 (pH 4.5), 5
mM MgS04, and purified on a denaturing 20% polyacrylamide, 7 M urea gel.
The protected RNA fragment was then digested with RNase T1, proteinase K
treated, and run on a 20% polyacrylamide, 7M urea gel as described
below. This procedure is shown schematically in Figure 2-2. The
digestion products were analyzed using the Betagen gel scanner.
RNase T1 Digestions
Isolated labeled RNA fragments were ethanol precipitated with 20
Mg glycogen (Boehringer Mannheim) and dissolved in 6 ¡il of 25 mM sodium
citrate (pH 3.5), 7M urea, 1 mM EDTA, 0.035% xylene cyanol and 0.035%
bromphenol blue. RNase T1 (6 units) was added and incubated at 50C for
1 h. The sample was then boiled for 1 min and another 6 units of RNase
T1 was added and incubated for another hour at 50C. Proteinase K
(1 Mg) was then added and incubated at 37C for another hour. The
reaction mixtures were then boiled, quick chilled on ice, and loaded
directly onto a 20% polyacrylamide, 7M urea gel.
Gel Purification of Oligonucleotides
All oligonucleotides that were gel purified were either located by
autoradiography or UV shadowing. The pieces were cut out of the gel,
crushed, and eluted overnight at 37C in 150 /ul to 1 ml of water
depending on the size of the gel piece. Large pieces of polyacrylamide
were removed by centrifugation in an eppendorf microfuge and the
supernatant was passed through a sterile disposable polypropelene column


24
vRNA
> DNA oligo Poly(A)
Hybridize
T1 Digest
20% Acrylamide Native Gel
"''N
Isolate RNA-DNA Hybrid
DNase
5' end label
Denature
20% Acrylamide 7M Urea Gel
Poly(A)
Protected RNA oligo
Purify protected RNA oligo
T1 Digest
Proteinase K
20% Acrylamide 7M Urea Gel
T1 Resistant Oligo
T1 Digestion Products
Fig. 2-2. Schematic diagram of 5' end-labeled vRNA hybridization and
RNase T1 digestion assay to determine the error frequency of the
polymerase at specific sites on the genome.


25
with paper disc (Isolabs). This was then ethanol precipitated with 20
Hg glycogen. Fragments eluted from very large gel pieces were extracted
three or four times with absolute ethanol to eliminate any urea.
5' End-labelinE
RNA and DNA fragments were routinely end labeled with 10 /Ci of
[7-32P]ATP using 10 units of T4 polynucleotide kinase (New England
Biolabs). When RNA pieces were to be sequenced, however, 50 tCi was
used instead for each hybridization and digestion that started with 10
Hg poliovirion RNA.
RNA Sequencing
RNase Tl-resistant oligonucleotides were gel purified and ethanol
precipitated with 15 /ig of tRNA. The RNA was divided into five aliquots
which were digested with either RNase T1 (Calbiochem Corporation), RNase
U2 (Bethesda Research Laboratories), RNase PhyM (Bethesda Research
Laboratories), RNase B. cereus (Bethesda Research Laboratory) or RNase
CL3 (Pharmacia). RNase T1 and U2 digestions conditions were 25 mM
sodium citrate (pH 3.5), 7M urea, 1 mM EDTA, 0.035% dyes, with 2
units/ml and 0.5 unit/ml RNase respectively. Digestion conditions for
PhyM were 25 mM sodium citrate (pH 5.0), 7M urea, 1 mM EDTA, 0.035%
dyes, with 100 units/ml RNase. Digestion conditions for B. cereus were
25 mM sodium citrate (pH 5.0) with 200 units/ml RNase B. cereus.
Digestion conditions for CL3 were 10 mM sodium phosphate (pH 6.5), 10 mM
EDTA, and 50 units/ml RNase. All digestions were for 15 min at 55C
except CL3 which was done at 37C. Digestions with B. cereus and CL3
were stopped by the addition of 7M urea and 0.035% dyes. All digestions


26
were stopped by freezing on dry ice. Samples were then boiled for 3 min
and quick chilled before loading onto a 20% polyacrylamide, 7M urea gel.
Gel Electrophoresis
All 20% polyacrylamide gels were at a 30:1 ratio of acrylamide to
bisacrylamide. All were run in 100 mM TrisHCl, 100 mM H3B03, and 2 mM
EDTA (pH 8.0). All long (45 cm) urea gels were prerun at 25 watts, and
all short (20 cm) urea gels were prerun at 15 watts, for a minimum of 1 h.


CHAPTER 3
DETERMINATION OF POLIOVIRUS RNA POLYMERASE ERROR FREQUENCY IN VITRO
Introduction
The poliovirus RNA polymerase has been purified from infected
cells and copies poliovirion RNA and other polyadenylated RNAs in vitro.
Synthetic homopolymers including poly(A):oligo(U), poly(C):oligo(I), and
poly(I):oligo(C) serve as template:primers for the polymerase as well.
By copying synthetic homopolymers with the purified polymerase and
differentially labeled complementary and noncomplementary substrates,
the error frequency of the polymerase can be measured directly. Another
technique that was used in this study involved copying synthetic
homopolymers in the presence of both 32P-labeled complementary and
noncomplementary ribonucleotide substrates and digesting the product RNA
with PI nuclease. PI nuclease digests the RNA to 5'-ribonucleoside
monophosphates which can be separated and counted to determine the error
frequency. Both of these techniques allow one to vary the reaction
conditions and observe how these changes affect the error frequency of
the polymerase. Reaction conditions that were looked at included the
effects of template, substrate, divalent cations, nucleotide substrate
concentration and temperature. Both of these in vitro procedures
eliminate the bias of converting viable mutation rates to error rates of
the polymerase, in contrast to what most in vivo studies measure.
27


28
The error frequency of the poliovirus polymerase has been measured
in vitro on a poly(A) template using differentially labeled
complementary and noncomplementary substrates by Mary Merchant-Stokes
(Merchant-Stokes, 1985). The error frequency was measured at pH 7 or pH
8 and at 3 mM or 7 mM MgCl2. The error frequency ranged from 0.7 X 10'3
to 5.4 X 10'3 depending on the reaction conditions. Increasing the pH
from 7 to 8 increased the error frequency of the polymerase 2 to 3 fold.
Increasing the MgCl2 concentration from 3 mM to 7 mM also increased the
error frequency about 2 fold. A correlation was seen between reaction
conditions that increased the elongation rate of the polymerase with
reaction conditions that increased the error frequency of the
polymerase.
Results
Determination of the Km for Each Ribonucleotide Substrate in the
Polymerase Reaction
One of the variables examined was the effect of nucleotide
concentration on the error frequency of the poliovirus polymerase. When
measuring the effect of nucleotide concentration, one should also take
into account the possible effect of the nucleotide concentration
relative to the Km for that particular nucleotide. It was possible that
using nucleotide concentrations that were far away from the Km might
affect the error rate. In addition, it was important to know if there
were large differences in the Km for the four different ribonucleotide
substrates as this might affect the error rate as well (See Chapter 5
for discussion of the Km Discrimination Model).


29
To determine the apparent Kms for each ribonucleotide, the initial
velocities of the polymerase reaction were measured as a function of the
concentration of each ribonucleotide substrate. This was done at a
variety of concentrations for one nucleotide (0.4 /M, 0.8 /M, 2.5 ¡i, 5
M, 10 tM, 20 /M, 40 M, and/or 80 /jM) while the other three nucleotides
were kept constant at 500 /M. The amount of ribonucleotide incorporated
in the product RNA was measured as a function of time to determine the
initial velocities (Figures 3-1 and 3-2). The Km for each nucleotide
was then determined by using a Lineweaver-Burk double-reciprocal plot of
the initial velocities vs. the substrate concentration (Figures 3-3 (UTP
and ATP) and Figure 3-4 (CTP and GTP)). The Km values for each
ribonucleotide were as follows: ATP = 10 /M, UTP = 7 M, CTP = 6 M,
and GTP = 5 pM.
Determination of the Polymerase Error Frequency by Using Differentially
Labeled Ribonucleotide Substrates
The error frequency of the poliovirus RNA polymerase was
determined by measuring the rate at which a noncomplementary
ribonucleotide substrate was incorporated into the product RNA using
synthetic homopolymeric RNAs as templates. When poly(A) was used as the
template [3H]UTP was used as the complementary substrate and 32P-labeled
ATP, GTP and CTP were used as the noncomplementary substrates. The
error frequency of the polymerase reaction was defined as the moles of
noncomplementary nucleotide incorporated divided by the total moles of
nucleotide incorporated into the product RNA.
Effect of changing the templates and substrates
The error frequency of the poliovirus polymerase was measured on
poly(A), poly(C), and poly(I) with each of the three different


30
CO
0)
o
E
D.
D
CD
i-
O
CL
i-
o
o
c
CL
5
3
[UTP]
Q .4 uM
.8 uM
a 5 uM
o 10 uM
80 uM
o
o
E
CL
-o
4)
O
CL
k_
O
o
c
a
2
<
[ATP]
.4 uM
.8 uM
5 uM
o 10 uM
80 uM
Figure 3-1. Effect of [UTP] and [ATP] on the initial velocity of the
poliovirus polymerase reaction. Initial velocities (v ) of the
polymerase were determined for five different concentrations of UTP (top
graph) or ATP (bottom graph). Standard reaction conditions were used on
vRNA with an oligo(U) primer (3 mM MgCl2, pH 8, 30C) except that the
initial reaction volume was 150 and 30 /il aliquots were removed and
counted every ten minutes.


31
Cfl
o
o
E
Q.
"O
0)
*1
10
o
Q.
w
O

c
a
5
O
[CTP]
.4 uM
.8 uM
a 5 uM
10 uM
40 uM
w
0)
o
E
Q.
TJ
V
o
a
h_
o
o
c
a.
2
(5
[GTP]
o 2.5 uM
5 uM
a 10 uM
40 uM
Figure 3-2. Effect of [CTP] and [GTP] on the initial velocity of the
poliovirus RNA polymerase reaction. Initial velocities (v ) of the
polymerase were determined at five different concentration of CTP (top
graph) and four different concentrations of GTP (bottom graph).
Reaction conditions were as described in legend to Figure 3-1.
Measurements of (v0) for GTP were repeated and both values were plotted
on the Lineweaver/Burk plot shown in Figure 3-4.


32
1/[UTP]
Fig. 3-3. Determination of Km for ATP aned UTP using Lineweaver-Burke
plots. Double reciprocal plot of v'1 vs. [NTP]1 for poliovirus
polymerase where v represents the initial velocity of incorporation for
each NTP (UTP and ATP). Initial velocities were determined at each
concentration using the data shown in Figure 3-1.


33
1/[CTP]
1/[GTP]
Fig. 3-4. Determination of Km for CTP and GXP using Lineweaver-Burk
plots. Double-reciprocal plot of v'1 vs. [NTP]'1 for poliovirus
polymerase where v represents the initial velocity of incorporation for
each NTP (CTP and GTP). Initial velocities were determined at each
concentration using the data shown in Figure 3-2.


34
noncomplementary ribonucleotide substrates for each template (Table 3-1).
All measurements were made using equimolar amounts of complementary and
noncomplementary ribonucleotide substrates. There was no
significant difference between the error frequency on poly(A) and
poly(C), however the error frequency did appear to be slightly lower on
poly(I) (less than 2-fold). There appeared to be no significant
difference between the four different noncomplementary ribonucleotide
substrates. It was not possible to determine the error frequency on
poly(U) or poly(G) because of very low levels of polymerase activity on
these templates.
Effect of changing the nucleotide concentration
By far the largest factor found to affect the error frequency of
the polymerase was the concentration of ribonucleotide substrates. All
assays were done on a poly(A) template using [3H]UTP as the correct
ribonucleotide and [32P]CTP as the incorrect ribonucleotide substrate.
The UTP concentration was varied from 0.74 /M to 74 /M and the CTP
concentration from 0.74 /M to 7.4 /M. It was found that as the ratio of
correct to incorrect ribonucleotide substrate was increased from 1:1 to
10:1, the error frequency decreased approximately 10-fold. If the ratio
of correct to incorrect ribonucleotide substrates was decreased from 1:1
to 0.1:1, the error frequency remained the same (Table 3-2). It was
interesting to find that decreasing the ratio of correct to incorrect
nucleotides had no effect on the error frequency. This suggested that
the error frequency had reached a maximum value that was not farther
increased by decreasing the relative concentration of the complementary
substrate. In marked contrast, increasing the relative concentration of


35
Table 3-1. Error Frequency of the Poliovirus RNA Polymerase
Reaction3
Noncomplementary
Complementary
Errorb
Conditions
Substrate
Substrate
Frequency
Poly(A)
[32P ] ATP
[3H]UTP
3.8
+
0.6 X 10'3
3 mM MgCl2
[32P] CTP
[3H]UTP
2.9
+
0.4 X 10'3
pH 8
[32P]GTP
[3H]UTP
3.8
+
0.5 X 10'3
Poly(C)
[32P] ATP
[3H]GTP
2.9
+
1.2 X 10'3
3 mM MgCl2
[32P] CTP
[3H]GTP
4.8
+
2.0 X 10'3
pH 8
[32P]UTP
[3H]GTP
2.5
+
0.2 X 103
Poly(I)
[32P] ATP
[3H]CTP
2.2
+
0.5 X 10'3
3 mM MgCl2
[32P]GTP
[3H] CTP
1.6
+
0.4 X 10'3
pH 8
[32P ] UTP
[3H]CTP
1.9
+
0.6 X 10'3
a Reaction conditions were as described in Materials and Methods except
the final volume of the poly(A) reactions was 50 /I. In the poly(C)
and poly(I) reactions, the final volume was 30 fj.1 and the total
nucleotide concentrations were 25.6 /M and 16.6 /M respectively. The
error frequency on poly(A) was determined by Mary Merchant-Stokes
(Merchant-Stokes, 1985).
b The error frequency was defined as the pmoles of noncomplementary
nucleotide incorporated divided by the total pmole of nucleotide
incorporated into product RNA. For example, 432,802 cpm [3H]UMP
(2.02 X 104 cpm/pmole) and 8,038 cpm [32P]AMP (1.33 X 105 cpm/pmole)
were incorporated at pH 8, 3 mM MgCl2. A counting efficiency for 3H
of 0.33 was assumed.


36
Table 3-2. Effect of Nucleotide Concentration on the Error Frequency of
the Polymerase
[UTP]
[ CTP ]
[UTP]/[CTP]
Error
Frequency3
0.7 /iM
7.4 /iM
0.1
3.2
0.8 X 10'3
7.4 /iM
7.4 /iM
1
2.9
0.4 X 10'3
74.0 /iM
7.4 /xM
10
2.0
0.8 X 10'4
7.4 /iM
0.7 /iM
10
4.4
1.6 X 10'4
3 All reactions were on poly(A), 3 mM MgCl2, pH 8.


37
the complementary substrate resulted in a significant decrease in the
error frequency. There did not appear to be a direct relationship
between the error frequency and the Km's of the nucleotide substrates.
For example, when the concentration of the noncomplementary substrate
was held constant at a concentration near its Km, decreasing the
concentration of the complementary substrate from a concentration near
its Km value to one-tenth of its Km had no effect, whereas increasing
its concentration to ten times the Km value decreased the error
frequency. Thus, it appears that the ratio of the nucleotide substrate
concentrations and not their absolute concentrations has the greatest
affect on the error frequency.
Effect of MnCl
It was found that substituting .5 mM MnCl2 as the divalent cation
resulted in a 2-fold increase in poliovirus polymerase error frequency
relative to 3 mM MgCl2 (Table 3-3) All of these assays were done on a
poly(A) template using [3H]UTP as the correct ribonucleotide substrate
and [32P]CTP as the incorrect ribonucleotide substrate. This
measurement was made at both equimolar ratios of complementary to
noncomplementary ribonucleotide substrate and at a 10:1 ratio of
complementary to noncomplementary ribonucleotide substrate (7.4 /xM UTP,
0.7 /xM CTP) Again it was seen that increasing the ratio of
complementary to noncomplementary ribonucleotide substrate to 10:1
resulted in about a 9-fold decrease in the error rate.
Effect of temperature
Changes in temperature of the reaction conditions appeared to have
no effect on the error frequency of the poliovirus polymerase. (Table 3-4).


38
Table 3-3. Effect of MnCl2 on Poliovirus Polymerase Error Frequency
Nucleotide Ratio3
Compl./Noncompl,
10:1
10:1
1:1
1:1
Divalent Cationb
3 mM MgCL,
0.5 mM MnCl2
3 mM MgCl2
0.5 mM MnCl2
Error Frequency
0.8 0.1 X 10~4
1.4 0.3 X 10'4
7.2 2.0 X 10'4
12.6 2.4 X 104
Increase in
Error Frequency
1
2
9
16
3 At 10:1 ratio of complementary to noncomplementary substrates, 7.4 /xM
UTP and 0.7 /xM CTP were used respectively. At 1:1 ratio of
complementary to noncomplementary substrates, 7.4 /zM UTP and 7.4 /xM CTP
were used respectively.
b All reactions were on poly(A) at pH 8.


39
Table 3-4. Effect of Temperature on Polymerase Error Frequency
Nucleotide ratio3
Compl./Noncompl. 30C
37C
42C
1:1 7.2 2.0 X
10:1 8.3 0.8 X
io-4
IO'5
7.4 1.9 X
1.3 0.3 X
IO'4
IO4
6.8 2.2 X IO4
9.3 5.8 X IO5
3 At 1:1 ratio off complementary to noncomplementary nucleotide, 7.4 M
UTP and CTP were used. At 10:1 ratio of complementary to
noncomplementary nucleotide 7.4 M UTP and 0.7 M CTP were used. All
reactions were on poly(A), 3 mM MgCl2, pH 8.


40
The error frequency was determined at 30C, 37C, and 42C in reactions
that contained 3 mM MgCl2, pH 8. These assays were also done on a
poly(A) template at both equimolar ratios of complementary to
noncomplementary ribonucleotide substrates and at a 10:1 ratio of
complementary to noncomplementary ribonucleotide substrate (7.4 M UTP,
0.7 /M CTP) .
It should be noted that the absolute value for the error frequency
determined at a nucleotide ratio of 1:1 and at 3 mM MgCl2, pH 8 in
Tables 3-3 and 3-4 was about 4-fold lower than the values obtained under
similar conditions in Tables 3-1 and 3-2. The reason for this change in
error frequency is not clear, however all experiments were internally
controlled so that relative changes in the error frequency due to
changes in the reaction conditions were real. The change in ratio of
nucleotide concentration done during the MnCl2 and temperature variation
experiments still resulted in a 9-fold relative change in the error
frequency.
Error Frequency as Determined by Pi Nuclease Digestion
Poly(A) templates were copied with both 32P-labeled complementary
and noncomplementary ribonucleotide substrates. The product RNA was
then separated from unincorporated labeled ribonucleotides and digested
to completion with PI nuclease. PI nuclease digests RNA to
ribonucleoside 5'-monophosphates and should yield a 32P-labeled
noncomplementary ribonucleoside only if it was actually incorporated
into the product RNA (Figure 3-5). These digestion products were
separated by high voltage ionophoresis and visualized by
autoradiography (Figure 3-6). The ribonucleoside 5'-monophosphates were


5 oligoU
2' PolyA
5'
polymerase
PPP u
+ PPPC ^ 3,
5 oligo UpUpUpUpUpUpCpUpUpU
3 PolyA
iPI
Nuclease
5' 4-* ^ '!* 4-* 4-* *
oligo U pU pU pU pU pU pC pU pU pU
3/ PolyA
^ 5 PC
5 pU
5'
3'
5'
Figure 3-5. Diagram showing 5'-ribonucleoside monophosphates recovered from product RNA
digested with PI nuclease. Illustrated is the synthesis of product RNA on a poly(A) template
the presence of an oligo(U) primer by the polymerase in the presence of [^PJUTP and [32P]CTP.
The expected PI cleavage of the labeled product RNA is also shown.


42
Figure 3-6. High-voltage ionophoretic separation of PI nuclease
digestion products. RNA was synthesized in the presence of [32P]UTP and
the following 32P-labeled noncomplementary ribonucleotides: [32P]ATP
(lane 1), [32P]CTP (lane 2), [32P]GTP (lane 3). Lane 4 is a marker lane
containing 5'-ribonucleoside monophosphates UMP (pU), GMP (pG), AMP
(pA), and CMP (pC).


43
then cut out and counted to determine the error frequency. All of these
assays were done at 3 mH MgCl2, pH 8. The error frequency ranged from
2.0 X 10'3 to 4.8 X 10'3 (Table 3-5).
Discussion
The results of these experiments indicated that the error
frequency of the poliovirus RNA polymerase was affected by changes in
the in vitro reaction conditions. A number of variables
including the type of divalent cation, the ratio of complementary to
noncomplementary ribonucleotide substrates, and the type of template
copied had an effect on the error frequency of the polymerase.
Previously it was seen that the pH and the MgCl2 concentration had an
effect on the error frequency. The only variables measured that had no
effect on the error frequency were the temperature and the
noncomplementary ribonucleotide substrate used in the reaction.
The type of homopolymeric RNA template used in the reaction seemed
to have little or no effect on the error frequency of the polymerase.
This was similar to the results that have been observed with reverse
transcriptases (Battula and Loeb, 1974). It should be noted, however,
that alternating copolymers are typically copied with more fidelity by
DNA polymerases, and that the fidelity of reverse transcriptase appears
to be sequence dependent on heteropolymeric templates, but not
homopolymeric templates (Battula and Loeb, 1974; Loeb and Kunkel, 1982;
Richetti and Buc, 1990).
Changing the ribonucleotide that was added as the noncomplementary
substrate did not affect the error frequency of the poliovirus
polymerase. The molecular mechanisms involved in the selection of


44
Table 3-5. Error Frequency as Determined by PI Nuclease Digestion
Reaction
Conditions
Noncomplementary
Substrate
Complementary
Substrate
Error Frequency
Poly(A)
[32P] ATP
[32P]UTP
2.9 2.8 X 1CT3
3 mM MgCl2
[32P] CTP
[32p]utp
4.8 2.4 X 10'3
pH 8
[32P]GTP
[32P]UTP
2.0 2.4 X 10'3


45
correct versus incorrect nucleotides by polymerases is still not
understood and needs further exploration. While numerous models have
been proposed, the data does not clearly choose one model above all
others. For a discussion of these models, see Chapter 5.
Substituting MnCl2 for MgCl2 was found to increase the error
frequency of poliovirus polymerase by 2-fold. Manganese chloride
decreases the fidelity of DNA polymerases from 2 to 25 fold, depending
on the polymerase (Sirover and Loeb, 1977; Beckman et al., 1985). This
decrease in fidelity has been attributed mostly to the binding of the
Mn+2 to the template. This is thought to facilitate the formation of
noncomplementary bases pairs during polymerization by changing the
hydrogen bonding properties of the nucleotides. At higher
concentrations of Mn+2 it is also possible that interactions with the
enzyme or nucleotide substrates affect the error frequency.
The largest influence on the error frequency of the polymerase was
the relative concentrations of the ribonucleotide substrates used in the
in vitro reactions. At equimolar concentrations or less of
complementary to noncomplementary ribonucleotide substrates, the error
rate was at its highest value of about 3 x 10"3. With a 10-fold increase
in the ratio of complementary to noncomplementary ribonucleotide
substrates, the error rate correspondingly decreased 10-fold. This was
observed whether the complementary nucleotide concentration was equal to
the Km of the substrate or 10-fold above it. Therefore, the ratio of
correct to incorrect substrates appeared to be more important in
determining the error rate of the polymerase than substrate
concentration relative to the Km.


46
It was rather surprising to find that changing the temperature did
not affect the error frequency of the poliovirus polymerase. Since the
temperature is known to affect the elongation rate of the polymerase,
there does not appear to be a simple relationship between the elongation
rate of the polymerase and the error frequency of the polymerase.
Apparently, temperature does not affect the base selection process.
Measurements of the error frequency after digesting the product
RNA with PI nuclease supported the data obtained by using differentially
labeled ribonucleotide substrates. One disadvantage of the assay that
uses differentially labeled substrates is that any contaminating 32P-
labeled complementary nucleotide that might be present in the 32P-
labeled noncomplementary ribonucleotide could be incorporated and
counted as an error. The PI nuclease digestion assay eliminates this
problem as all substrates are 32P-labeled. The error rate determined in
this manner varied from 2.0 X 10'3 to 4.8 X 10'3. These numbers were in
general agreement with the error frequencies determined under the same
reaction conditions by differentially counting the 3H and 32P-labeled
product RNA.
Overall, the error frequency of the poliovirus RNA polymerase
measured in vitro ranged from 8.3 X 10'5 to 4.8 X 103, a change of 65-
fold. Thus, while any one change in reaction conditions had a
relatively small effect, together they can cause a large change in the
error frequency of the poliovirus RNA polymerase.


CHAPTER 4
DETERMINATION OF THE POLIOVIRUS RNA POLYMERASE ERROR FREQUENCY IN VIVO
Introduction
The recent work by Steinhauer and Holland (Steinhauer and Holland,
1986) shows that it is now possible to measure the error frequency of
viral RNA polymerases both in vitro and in vivo. They developed a
technique that can be modified to measure the error frequency of most
RNA viruses. This technique involves the measurement of the error
frequency at a specific nucleotide in the viral RNA. Although there are
some limitations on which nucleotides can be used in this assay, it is
possible to select specific sites from different regions of the viral
genome. It does not require that the viral RNA be infectious unlike
most other in vivo techniques that only measure viable mutation rates or
error frequencies. The viral RNA used in this assay can either be
purified from the cytoplasm of infected cells or from purified virions.
Steinhauer and Hollands measurements made with VSV using this technique
both in vivo and in vitro indicate a high error frequency of the VSV RNA
polymerase, which ranged from 1 X 10"4 to 4 X 104 (Steinhauer and
Holland, 1986).
Much controversy exists over the error frequency of the poliovirus
polymerase. While no direct measurements have been made, the
evolutionary rate of poliovirus has been measured in many different ways
with apparently conflicting results (for details, see Chapter 1). What
47


48
is now needed is a direct measurement of the error frequency of
poliovirus RNA polymerase in vivo. This was the primary objective of
the studies described in this chapter. I have adopted and modified the
technique of Steinhauer and Holland to measure the error frequency at
specific sites in purified poliovirion RNA.
Results
Purification of RNA Oligonucleotides
The basic approach used to determine the polymerase error rate in
vivo was a procedure that was a modification of the method previously
described by Steinhauer and Holland (Steinhauer and Holland, 1986).
Briefly, 32P-labeled RNA was hybridized to a synthetic DNA
oligonucleotide and digested with RNase T1. The protected RNA
oligonucleotide that was complementary to the DNA was isolated by gel
purification and then digested with RNase T1. The rate of change in the
single G residue found in the protected fragment was determined by
quantitating the amount of the protected fragment that was resistant to
digestion (for details, see Chapter 2 and figure 2-1). The technique
was initially developed on pOF1205 transcript RNA, which consists of the
3' terminal 1000 bases of poliovirus RNA. The transcript RNA was
labeled with either [32P]UMP or [32P]AMP, hybridized to a synthetic DNA
oligonucleotide, digested with RNases T1 and U2 or RNase T1 alone, and
run on a 20% polyacrylamide, 7 M urea gel. A protected oligonucleotide
of the expected size (34 nucleotides) was recovered from the [32P]UMP-
labeled RNA that was hybridized to the synthetic DNA oligonucleotides
(Figure 4-1, lanes IB and 1C). This oligonucleotide was not present
when the RNA was digested in the absence of the synthetic DNA (Figure 4-1,


49
lanes 1A and ID). When the transcript RNA was labeled with [32P]AMP,
the same protected oligonucleotide was present (Figure 4-1, lanes 2B and
2C). In this case, however, it was apparent that a ladder of labeled
poly(A) fragments was also present (Figure 4-1, lanes 2A to 2D). This
result indicated that the protected RNA oligonucleotide that was
isolated from this gel was contaminated with a poly(A) fragment of the
same size. This was confirmed in other experiments where the [32P]UMP-
labeled 34mer was further characterized. The labeled 34mer was gel
purified, digested with RNase T1, and run on a denaturing polyacrylamide
gel. Two major bands of the expected size were observed along with a
small amount of the 34mer that was resistant to digestion (Figure 4-2).
The 34mer was isolated from the gel and 5'-end labeled with [y-32P]ATP
and polynucleotide kinase. The sequence was then determined by
enzymatic sequencing procedures as described in Chapter 2 (Figure 4-3).
It was clear that the Tl-resistant 34mer was in fact contaminated with a
large amount of poly(A). This would be an obvious problem in any
experiments where the 3' terminal poly(A) sequence actually was labeled
(for example, [32P] P04-labeled virion RNA). This problem was not dealt
with by Steinhauer and Holland since VSV RNA is not polyadenylated.
Avoiding the coisolation of poly(A) with the protected RNA
oligonucleotide was solved by initially isolating the RNA-DNA duplex on
a nondenaturing 20% polyacrylamide gel (Figure 4-4). The RNA-DNA duplex
comigrates on this gel with poly(A) fragments which are much longer than
the protected RNA oligonucleotide which is part of this duplex. The
duplex was isolated from the nondenaturing gel, denatured with urea, and
run on a 20% polyacrylamide, 7 M urea gel to separate the protected RNA


50
1A IB 1C ID 2A 2B 2C 2D 3C
Figure 4-1. Isolation of RNA oligonucleotide protected from BF8 after
RNAse T1 digestion. 32P-labeled poliovirus specific RNA was transcribed
from pOF1205 DNA. The RNA was labeled with either [32P]UMP (lanes 1A -
ID) or [32P]AMP (lanes 2A 2D). THe RNA was either directly digested
with RNase T1 and U2 (lanes A) or RNase T1 (lanes D) or was first
hybridized with synthetic DNA oligonucleotide BF8 and then digested with
RNases T1 and U2 (lanes B) or RNase T1 (lanes C). Poliovirion RNA
labeled in vivo with [32P]P04, hybridized with BF8, and digested with
RNase T1 is shown in lane 3C.


51
12 3 4 5 6
Figure 4-2. Final RNase TI digestion products of RNA oligonucleotide
protected by BF8. The oligonucleotide was isolated from a gel similar
to that shown in Figure 4-1 where pOF1205 transcript RNA was labeled
with [32P]UMP. The initial hybridization and RNase T1 digestion
conditions were as follows: 37C (lane 1), 55C (lane 3), 50C (lane
4), 45C (lane 5), 40C (lane 6). The RNA in lane 2 was hybridized at
55C and then digested at 37C. All final T1 digestions were done at
55C.


52
O
+
O O < <
4>
"O
"O
<0
3
+
O O < <
Figure 4-3.
BF8. First
Figure 4-2.
lanes 3, 4,
RNA sequencing of TI resistant oligonucleotide protected by
set of digests and ladder (five lanes) are from lane 2 in
Second set of digests (last four lanes) are from pooling
5, and 6 in Figure 4-2.


53
Figure 4-4. RNA hybridization and TI digestion products run on a native
20% polyacrylamide gel. Lanes 1 and 2 are [32P]UMP labeled pOF1205
transcripts, lane 3 is [32P]P04 labeled vRNA, and lane 4 contains
[32P]AMP labeled pOF1205 transcripts. Lane 1 was not hybridized with
BF8.


54
oligonucleotide from the DNA oligonucleotide and any contaminating
poly(A) (Figure 4-5). It should be noted that the DNA was engineered to
have a 15 base long poly(A) tail so that it would separate from the RNA
oligonucleotide on a denaturing gel. In earlier studies, I found that
it was not possible to quantitatively remove all of the DNA from this
duplex by a simple digestion with DNase. Thus, the two-step gel
purification procedure was adopted. The purified RNA oligonucleotide
was isolated from the second gel and then digested with RNAse T1 to
determine the error rate. The error rate was defined as the
radioactivity (cpm) in the RNase T1 resistant band relative to the
radioactivity in the two oligonucleotides that were the RNase T1
digestion products.
Polymerase Error Frequency Determined by Using i32P1PC^-Labeled
Poliovirion RNA
The error frequency has been measured at two sites in the
poliovirus genome using the procedure described above and [32P]P04-
labeled RNA. The two sites were at nucleotide 6883 in the 3Dpo1 coding
sequence and at nucleotide 5648 in the 3Cpro coding sequence. DNA
oligonucleotides BF8 and BF10 were used to isolate RNA oligonucleotides
that contained these two sites (see Table 2-1 for the exact sequences).
The RNA oligonucleotide protected by BF8 was digested to completion with
RNase T1 and was analyzed by gel electrophoresis (Figure 4-6). A very
small but detectable amount of the oligonucleotide was resistant to
digestion (Figure 4-6, lane 1). The radioactivity (cpm) recovered in
this resistant band represented 4.3 X 10'3 of the total radioactivity
recovered in all three bands (i.e., the two major bands representing the
digestion products and the resistant band). Thus, the polymerase error


55
1 2 3
Figure 4-5. Separation of RNA oligonucleotide protected by BF8 from
contaminating poly(A) by gel purification. The RNA-DNA duplex was first
isolated by gel purification on a nondenaturing 20% polyacrylamide gel.
The duplex was then denatured and run on a 20% polyacrylamide, 7 M urea
gel. Poliovirion RNA was labeled in vivo with [32P]P04 (lane 1) SP6
polymerase transcripts of p0F1205 DNA were labeled with either [32P]UMP
(lane 2) or [32P]AMP (lane 3).


56
1 2
Figure 4-6. Final RNase T1 digestion products of RNA oligonucleotide
protected by BF8 run on a 20% polyacrylamide, 7 M urea gel. The
protected oligonucleotide from poliovirion RNA labeled with [32P]P04 was
run after RNase T1 digestion (lane 1). BF8 protects a 34mer and digests
to a 22mer and a 12mer. Lane 2 shows undigested marker.


57
frequency at this site (i.e., nucleotide 6883) was 4.3 X 10'3 (Table 4-
1). The error frequency determined using BF10 was about the same with a
value of 0.9 X 10'3 (Table 4-1).
The major drawback to this approach to determine the error
frequency was the relatively low specific radioactivity of the labeled
virion RNA and the large amount of radioactivity (50 mCi) that was
required to label the vRNA synthesized in infected cells. For these
reasons, a second approach was used to determine the in vivo error
frequency of the poliovirus RNA polymerase.
Polymerase Error Frequency Determined Using 5' End-Labeled
Oligonucleotides from Poliovirion RNA
The error frequency at eight different sites in the poliovirus
genome were determined using the 5' end-labeling technique. These sites
were located in constant and variable regions of the poliovirus genome
(illustrated in Figure 4-7). The technique used is summarized briefly
here, for details see Chapter 2. Poliovirion RNA was hybridized to a 5'
end-labeled DNA oligonucleotide and digested with RNase T1. This hybrid
was then purified on a nondenaturing 20% polyacrylamide native gel
(Figure 4-8). The hybrid was then 5' end-labeled with [y-32P]ATP and
polynucleotide kinase, treated with DNase and run on a denaturing 20%
polyacrylamide, 7 M urea gel (Figure 4-9). The band representing the
protected RNA oligonucleotide was isolated from the gel, digested with
RNase Tl, and run on a 20% polyacrylamide, 7 M urea gel. The digestion
products were quantitated by an AMBIS or Betagen gel scanner.
The final digestion products for the RNA oligonucleotides
protected by BF8 and BF10 are shown in Figure 4-10. The final digestion


58
Table 4-1. Poliovirus Error Frequency at Specific Sites
Labeling Method0
Protecting DNA oligonucleotide3 [32P]P04 5'-end label
BF8: 6862-6895 3D conserved 3.2 X 10"3
4.7
X
103
BF9:
7269-7296
3D
conserved
4.4
X
103
BF10:
5638-5671
3C
conserved
0.7 X 103
CM
on
X
103
BF11:
3971-3999
2B
conserved
3.5
X
103
BF25:
84-109
5'NC
conserved
3.8
X
103
BF27:
1229-1253
VP2
conserved
5.0
X
103
BF30:
1429-1450
VP2
variable
4.6
X
CO
b
11
BF35:
2757-2776
VP1
variable
3.2
X
10'3
aNumbers of protecting DNA oligonucleotide represent the nucleotides
protected in the poliovirus genome. 3D, 3C, 2B, VP2, and VP1 refer to
the genes encoded at these sites on the poliovirus genome. 5'NC refers
to the 5' non coding region. Conserved and variable refer to whether
these sites on the genome are known to change (variable) or not
(conserved).
bNumbers indicate corrected values based upon RNA sequencing and
redigesting with RNase T1 and are an average of at least 3 experiments.


59
VPg
Polyprotein
P'NC
Poly(A)
o
C
VP4 3BVPg
VP2
VP3
VP1
O
Q-
<
01
2B
2C
3A
3Cpro
3Dpo1
hs. co
'M-
CD
CO
CO CD
CO CO
CO
00
'<*
co co
CM M"
h-
O)
CO
00 CM
T- T-
CM
CO
ID
cd r-
C V
V
c
c
c c
Figure 4-7. Schematic diagram of the poliovirus genome with the various
sites examined by RNase T1 digestion indicated. C represents constant
regions on the genome and V represents variable regions on the genome.


60
Figure 4-8. Isolation of RNA oligonucleotides protected from BF27 and
BF10 after RNase T1 digestion. DNA oligonucletides BF27 and BF10 were
5'-end labeled and either run directly on the gel (Lanes 1 and 6,
respectively), or hybridized with virion RNA and digested with RNase T1.
BF27 hybridization and digestion products were run in lanes 2 and 3.
BF10 hybridization and digestion products were run in lanes 4 and 5.


61
12 3 4
Poly(A)
{
DNA
Figure 4-9. Isolation of 5'-end labeled protected RNA oligonucleotides.
RNA/DNA hybrids were isolated from a gel as shown in figure 4-8, 5'-end
labeled, digested with DNase, denatured with urea, and run on a 20%
polyacrylamide, 7M urea gel. Note the coisolation of poly(A) as well as
residual DNA that was not completely DNased.


Figure 4-10. Final RNase TI digestion products of 5'-end labeled
protected oligonucleotides. Lane 1 is from an oligoribonucleotide
protected by BF8 and lane 2 is from an oligoribonucleotide protected by
BF10. Lanes 3 and 4 are undigested markers.


63
products for the RNA oligonucleotides protected by BF9, 11, 25, 27, 30
and 35 are shown in Figures 4-11 through 4-16. All duplicate lanes
shown in Figures 4-11 through 4-16 represent reactions with various RNA
preparations that were separately hybridized and digested. All of these
assays were repeated at least three times for each oligonucleotide and
the average error frequency was determined (Table 4-1). This average
error frequency takes into account a correction factor that reduced the
error frequency by 25%. The RNAse T1 resistant oligonucleotides
protected by BF8 and BF11 were isolated from the gel and their
nucleotide sequence was determined (Figures 4-17 and 4-18,
respectively). While both of these sequencing gels clearly show the
presence of the three other nucleotides besides G, there is still a G
band present which represented approximately 25% of the total
radioactivity in the sequencing bands. In other experiments, the RNase
T1 resistant oligonucleotides were redigested with RNase T1. The amount
of RNase T1 resistant fragment that could be redigested with T1 ranged
from 10 to 50%. For this reason an average correction factor of 25% was
used.
Discussion
A modification of the technique developed by Steinhauer and
Holland was used to measure the in vivo error rate of poliovirus
polymerase. There were many unexpected problems encountered during the
modification of this technique. The first unforeseen problem was the
coisolation of contaminating poly(A) sequences with the RNA-DNA duplex.
The average size of poly(A) on poliovirion RNA is 75 100 nucleotides
and the heteroduplexes isolated were significantly smaller, 20 49


64
AAUAUAACAAAUUCCUAGCUAAAAUCAG 28mer
^T1 Digest
*AAUAUAACAAAUUCCUAG CUAAAAUCAG
18mer 10mer
1 2 3
4 5
28
Figure 4-11. Final RNase T1 digestion products of 5'-end labeled
oligonucleotides protected by BF9. Lane 1 is undigested marker.
BF9 protects a 28mer and digests to a 18mer (labeled) and a lOmer
(unlabeled). Lanes 2-5 are all separate reactions.


65
AUCAUAUCCUCACUAGUUAUUAUAACUAG 29mer
^T1 Digest
AUCAUAUCCUCACUAG
16mer
UUAUUAUAACUAG
13mer
Figure 4-12. Final RNase T1 digestion products from 5'-end labeled
oligonucleotide protected by BF11. Lane 4 is undigested marker.
BF11 protects a 29mer and digests to a 16mer (labeled) and a 13mer
(unlabeled). Lanes 1-3 are all separate reactions.


66
UUUUAUACUCCCUUCCCGUAACUUAG 26mer
^T1 Digest
UUUUAUACUCCCUUCCCG UAACUUAG
18mer 8mer
1 2 3 4 5
26
1 8
Figure 4-13. Final RNase T1 digestion products of 5'-end labeled
oligonucleotide protected by BF25. Lane 1 is undigested marker.
BF25 protects a 26mer and digests to a 18mer (labeled) and a 8mer
(unlabeled). Lanes 2-5 are all separate reactions.


67
CAAAAUAUG UACUACCACUACCUAG 25mer
^T1 Digest
CAAAAUAUG UACUACCACUACCUAG
9mer 16mer
Figure 4-14. Final RNase T1 digestion products of 5'-end labeled
oligonucleotide protected by BF27. BF27 protects a 25mer and digests to
a 9mer (labeled) and a 16mer (unlabeled). Lanes 1-4 are all separte
reactions.


68
*UUCACUCCUG AC AAC AACCAG 21mer
^T1 Digest
UUCACUCCUG ACAACAACCAG
10mer 11mer
Figure 4-15. Final RNase T1 digestion products of 5'-end labeled
oligonucleotide protected by BF30. Lane 1 is undigested marker,
protects a 21mer and digests to a lOmer (labeled) and a llmer
(unlabeled). Lanes 2 and 3 are separate reactions.
BF30


69
*AUAACCCAGCUUCCACCACG 20mer
^T1 Digest
AUAACCCAG CUUCCACCACG
9mer 11mer
1 2
20
3
Figure 4-16. Final RNase T1 digestion products of 5'-end labeled
oligonucleotide protected by BF35. Lane 3 is undigested marker,
protects a 20mer and digests to a 9mer (labeled) and a llmer
(unlabeled). Lanes 1 and 2 are separate reactions.
BF35


70
Figure 4-17. Enzymatic sequencing of the oligoribonucleotide protected
by BF8 before (right four lanes) and after RNAse T1 digestion (left four
lanes). The arrow marks the change in the single G residue to the other
three ribonucleotides.


71
Figure 4-18. Enzymatic sequencing of the oligoribonucleotide protected
by BF11 before (right five lanes) and after RNase T1 digestion (left
five lanes). The arrow marks the change in the single G residue to the
other three ribonucleotides.


72
nucleotides in length. Apparantly the poly(A) sequence in poliovirion
RNA is much more heterogeneous in length than previously thought.
Although only a small fraction of the virion RNA molecules contain short
poly(A) sequences, the amount present is still relatively large compared
to the amount of the RNase T1 resistant oligonucleotide which is
recovered in these experiments.
A second problem encountered was the quantitative removal of all
of the protecting DNA oligonucleotide. Original experiments used a DNA
oligonucleotide of the same length as the protected RNA oligonucleotide.
The hybrids were digested with DNase to remove the DNA and then digested
with RNase T1. These RNA oligonucleotides could not be digested to
completion as a result of residual DNA oligonucleotide. This problem
and the preceding problem of coisolating contaminating poly(A) sequences
were solved by engineering a poly(A)15 tail on the protecting DNA
oligonucleotide. A two step purification was then used to isolate the
heteroduplex on the first gel, and then a second gel was used to
separate the RNA from the protecting DNA oligonucleotide and any
contaminating poly(A) derived from the virion RNA.
One final problem encountered was an apparent gel shifting of the
RNase T1 digestion products. Unexpected high bands appeared on the
final RNase T1 digestion gels which could not be accounted for. They
were resistant to DNase and additional RNase T1 treatment. These bands
were apparently caused by contaminating proteins since they disappeared
with the addition of a proteinase K digestion step.
The error frequency of the poliovirus RNA polymerase was
determined at eight different sites on the poliovirus genome and ranged


73
from 0.9 to 5.0 X 103. These values are dependent on the complete
digestion of protected RNA oligonucleotide by RNase T1. Therefore, it
was important to determine how efficient this digestion was. For this
reason RNase T1 resistant oligonucleotides were sequenced when possible.
A large limiting factor on sequencing, however, was the small amount of
radioactivity in these RNase T1 resistant bands. When sequencing could
not be done, the RNase T1 resistant bands were redigested with RNase T1
and quantitated to determine the percent that could be redigested. This
percentage varied from 10 50%. Sequencing showed approximately 25% of
the RNase T1 resistant band contained a G residue For these reasons a
correction factor of 25% was used. With this correction factor taken
into account, the error frequency determined in vivo was in agreement
with the numbers determined in vitro.
The error frequency did not vary significantly between the eight
different sites measured on the poliovirus genome. Two of the sites
examined are known to mutate rapidly under selective pressure. Site
1439 (BF30) is located within the E-F loop of VP2. This site has been
observed to change rapidly when the virus is grown in the presence of
monoclonal antibodies which resulted in amino acid changes from aspartic
acid to asparagine and histidine (Page et al., 1988). Site 2765 (BF35)
is located in the B-C loop of VP1 in which host range mutants have been
mapped (Murray et al., 1988). This G is not conserved between serotypes
1 and 3 which results in an amino acid change from alanine to proline.
The loop regions of the capsid proteins in general are less conserved
between the the three serotype of poliovirus. The other six sites are
relatively conserved between the three serotypes of poliovirus. Sites


74
6883 (BF8) and 7286 (BF9) are located within ID*501 and any change in
these G residues would result in amino acid changes from methionine to
isoleucine and from alanine to serine, proline or threonine,
respectively. Site 5648 (BF10) is located within 3Cpr0 and a change in
this G residue would result in a change from glutamic acid to lysine,
glutamine, or a stop codon. Site 3986 (BF11) is located within 2B and
would result in an amino acid change from valine to isoleucine, leucine,
or phenylalanine. Site 101 is located in the 5' non-coding region of
poliovirus. This site, as well as the four other conserved sites
mentioned so far, appears to be very important in the replication of
poliovirus. The sixth conserved site, 1236 (BF27) is located within /3
barrel B of VP2. All of the p barrels of the capsid proteins are highly
conserved between the serotypes of poliovirus. A change in this G
residue would result in an amino acid change from methionine to
isoleucine.
While changes at these G residues would cause relatively minor
changes in some cases (BF35, from one nonpolar amino acid to another)
and relatively major changes in others (BF10, from a negatively charged
amino acid to either a positively charged amino acid, a polar uncharged
amino acid or a stop codon), there was no significant difference in the
error frequency determined at these different sites. They ranged from
.9 X 10'3 to 5 X 10'3, approximately a five-fold difference. These small
differences did not correlate with conserved and variable regions of the
viral genome. Thus, it appears that the error frequency of the
poliovirus polymerase is relatively constant at G sites across the
poliovirus genome.


CHAPTER 5
CONCLUSIONS AND PERSPECTIVES
Factors Affecting Poliovirus Polymerase Error Frequency
I found that a number of factors can affect the poliovirus RNA
polymerase error frequency which ranged in vitro from 8 X 10'5 to
5 X 10'3. These factors included the type of divalent cation, the
relative concentrations of correct and incorrect nucleotide substrates,
and the type of template used. The specific ribonucleotide that was
used as the incorrect substrate and changes in the temperature had
little or no effect on the error frequency of the polymerase. How these
factors exert their effect on nucleotide selection by the polymerase is
unknown at this time. The error frequency of the poliovirus polymerase
measured in vivo was also found to be similar to the in vitro values and
ranged from 9 X 104 to 5 X 10'3. No significant difference was found in
the error frequency at eight different sites in the poliovirus genome.
Two sites were selected because one is known to rapidly mutate when
grown under certain environmental conditions (in the presence of
monoclonal antibody) and the other is known to vary between the three
serotypes of poliovirus. The other six sites were selected as conserved
sites since they are unchanged between the three serotypes of
poliovirus. Overall, my results indicated that there was no significant
difference in the error frequency between the variable and the conserved
75


76
sites. This suggests that the variation observed in nature is due to
selection at the phenotypic level.
Models for DNA Polymerase Base Selection
There are several models that have been proposed to explain how
DNA polymerases select the correct incoming nucleotide. The first of
these models, the "Km Discrimination Model" (Goodman et al., 1977),
proposes that the difference in free energy between correct and
incorrect base pairings is increased in the presence of polymerases.
This model assumes that the rates for binding are the same for both
correct and incorrect nucleoside triphosphates, and that the
discrimination is based on differences in the dissociation rates. This
theory would then predict that the error rate is proportional to the
differences in the Km for the correct and incorrect nucleotides.
A second model, the "Conformation Model", proposes that the
polymerase changes in conformation with each nucleotide addition step,
which affects base selection. Most polymerases appear to have one site
for the binding of all four nucleotides. Therefore, one would predict
that these polymerases must have a mechanism to accommodate the
different structures of the nucleotides and base pairs. This model
predicts that a difference in Vmax for the correct and incorrect
nucleotides would be proportional to the error rate of the polymerase
(Watanabe and Goodman, 1982) .
Finally a third model, the "Energy Relay Model" (Hopfield, 1980),
proposes that the energy released by phosphate bond cleavage is used by
the polymerase to proofread the insertion of the next nucleotide. This
model would predict that the incorporation of the first nucleotide is


77
more error prone than the following nucleotides. This model was not
supported by the use of reversion frequency assays performed by Abbotts
and Loeb using mammalian DNA polymerases (Abbotts and Loeb, 1984). Nor
was it supported by Kuchta (Kuchta et al., 1988) using an elongation
assay with DNA polymerase I.
Because no large differences were seen between the
misincorporation rates of the four different ribonucleotides both on
homopolymeric templates in vitro and on heteropolymeric templates in
vivo. it is not clear that one of these models is greatly supported or
negated over another. It should be pointed out however that Michaelis
constants can vary significantly at specific sites and that these were
not measured. Factors that were found to affect the error frequency of
the poliovirus polymerase certainly may affect the conformation of the
polymerase and its intereaction with the nucleotide substrates.
However, it was also clear in sequencing the T1 resistant
oligonucleotides that all four nucleotides were present and one
incorrect base did not predominate over the others as one might expect
with the conformation model or the Km Discrimination Model. However,
RNA sequencing may not be quantitative enough to make this distinction.
It is possible that with the new capability to make synthetic RNA
oligonucleotides in the laboratory, that more sensitive measurements can
be made to determine the error frequency of poliovirus polymerase at
specific sequences utilizing the elongation assay developed by Bossalis
(Boosalis et al., 1987). This method involves elongating a 5' end-
labeled primer with the separate addition of the four different


78
nucleotide substrates and assaying for elongation by gel
electrophoresis.
Evolution Rates of Poliovirus
It is interesting to note that poliovirus is a very stable virus
when grown in tissue culture, yet it is known to change very rapidly
when it replicates in humans (for details see Chapter 1). The fact that
there are only three serotypes of poliovirus is probably a reflection of
having only a few functionally distinct neutralization sites, rather
than a reflection of phenotypic stability of any single antigenic site.
While these neutralization sites are in general very stable, any change
in the genome at these neutralization sites may result in an outbreak of
poliomyelitis as was seen with the relatively recent (1984) outbreak in
Finland. The changes in the neutralization sites, however, were not
solely responsible for this outbreak. In fact, the virus was
neutralized by antisera that was specific for that serotype. However,
the Finnish population used killed poliovirus vaccine, which overall
results in lower antibody titers than the attenuated poliovirus vaccine.
So it appeared that it was the combination of low antibody titers and
changes in some of the neutralization sites that resulted in the
poliomyelitis outbreak.
Master Sequence Theory
An important question then is why poliovirus is so stable when
grown under certain conditions and so quick to mutate under others. One
theory is that RNA viruses consist not of a single genotype but of a
distribution of related genotypes (Domingo et al., 1978, 1985). There
is a distribution around one or several degenerate master sequences


79
which are efficiently replicated. Under some conditions, the fraction
of master sequences relative to the total population is low and mutants
dominate the population. Mutants of high reproductive power relative to
the master sequence may modify the population drastically. This appears
to be the case when poliovirus is replicating in the human body. In
contrast, when poliovirius is grown in tissue culture, a single master
sequence is maintained. While the wild type is not characterized by a
single sequence, the wild type appears to have an unambiguous consensus
sequence which is probably identical to the master sequence. My data
clearly demonstrated that the poliovirus genome does change while
replicating in tissue culture at least at the eight different sites
measured for a small percentage of the population. However, sequencing
of the viable poliovirus population would have undoubtedly shown the
maintenance of the same consensus sequence that has been propagated in
the laboratory for years. It would thus appear that every nucleotide in
the poliovirus genome is selected for in one way or another. Selection
may be at the level of RNA structure as it relates to RNA packaging and
recognition by the replication machinery, as well as at the protein
level. As the poliovirus genome is initially translated as a large
polyprotein, the structure of this polyprotein may be very important for
cleavage by the viral proteases and any disturbance in this structure
may affect proteolytic processing. Also there is evidence that many of
the viral replicative proteins may only function in cis (Bernstein et
al.,1986) which may limit the perpetuation of mutant RNA sequences and
their corresponding mutant viral proteins.


80
RNA Genomes vs. DNA Genomes
It has already been mentioned that in general RNA genomes are of
limited length relative to DNA genomes. The error threshold for
maintenance of genetic information would be expected to correlate with
the sequence length, hence allowing a higher error frequency for RNA
polymerases. It should also be mentioned that RNA genomes are only
found as host dependent cellular parasites. Much of their replication
strategy therefore involves using host cell machinery, not their own.
RNA genomes also have much shorter replication times which contribute to
their higher evolution rate.
While DNA genomes are copied with a much higher fidelity than RNA
genomes, they have other strategies which allow for their evolution.
Recombination is one such strategy utilized by DNA genomes and to a
lesser extent can be utilized by some RNA genomes, including poliovirus.
One potentially interesting question is the evolution of RNA
viruses compared to DNA viruses. Do RNA viruses evolve independently of
most host cell functions whereas DNA viruses coevolve with the host
cell? Certainly some of the DNA viruses and the retroviruses which
utilize a DNA polymerase can integrate and be maintained and replicated
by the host cells replicative machinery. Perhaps it is therefore to
their advantage to evolve at a slower rate similar to their host cell.
Diversity may be much more advantageous for RNA viruses compared to DNA
viruses. Certainly the generation of defective interfering particles
and the large ratio of particle to infectious virions for RNA viruses
suggests that diversity is of great importance.


81
Future of RNA Viruses
Perhaps the most important lesson to be learned from the growing
accumulation of knowledge regarding the high error frequency of RNA
polymerases is to expect the continual emergence from time to time of
new diseases due to the continual evolution of RNA viruses. While RNA
viruses generally have a defined method of transmission and are
associated with a particular disease, there may be appearances of new
strains with markedly different host range, tissue tropism, disease
patterns, and virulence. This may be particularly true when a virus can
find a new niche, whether it be a new host, a new tissue, or a new
vector. We have witnessed the emergence of such new diseases. Acute
hemorrhagic conjunctivitis is one example of a new disease that emerged
in the 1960's and is caused by a picornavirus. And of course AIDS is
the latest example of such a new disease, which emerged in the 1970's
and is caused by a retrovirus which utilizes both RNA and DNA in its
replication strategy.


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BIOGRAPHICAL SKETCH
I was born December 2, 1960, in Washington, DC, the fifth and last
child of John and Ruth Ward. I lived in Silver Spring, Maryland, for 15
years where I attended Holiday Park Elementary School, Newport Junior
High School, and one year at Albert Einstein High School. I moved in my
junior year to Minnesota where I attended Fridly Senior High. The cold
quickly drove my family back to the Washington, DC, area where I
attended George Mason University from 1978 to 1983. I foolishly got
married while I was an undergraduate student, which ended in divorce
while I was a graduate student. Luckily, I met another graduate
student, Paul Kroeger, whom I married in 1987. We had one child in
1988, Alan Scott Kroeger, and are expecting our second child at the end
of this year.
89


I certify that I
conforms to acceptable
adequate, in scope and
Doctor of Philosophy.
I certify that I
conforms to acceptable
adequate, in scope and
Doctor of Philosophy.
I certify that I
conforms to acceptable
adequate, in scope and
Doctor of Philosophy.
I certify that I
conforms to acceptable
adequate, in scope and
Doctor of Philosophy.
I certify that I
conforms to acceptable
adequate, in scope and
Doctor of Philosophy.
have read this study and that in my opinion it
standards of scholarly presentation and is fully
quality, as a dissertation for the degree of
nes B. Flanegan,/(Jhair
Professor of Immunology and
Medical Microbiology
have read this study and that in my opinion it
standards of scholarly presentation and is fully
quality, as a dissertation for the degree of
^
Ernest Hiebert
Professor of Plant Pathology
have read this study and that in my opinion it
standards of scholarly presentation and is fully
quality, as a dissertation for the degree of
Lindsey Hath-Fletcher
Professor of Pathology and
Laboratory Medicine
have read this study and that in my opinion it
standards of scholarly presentation and is fully
quality, as a dissertation for the degree of
Richard Moyer
Professor of Immunbligy and
Medical Microbiology
have read this study and that in my opinion it
standards of scholarly presentation and is fully
quality, as a dissertation for the degree of
S
on
Sue Moyer
Professor of Immunology and
Medical Microbiology


This dissertation was submitted to the Graduate Faculty of the
College of Medicine and to the Graduate School and was accepted as
partial fulfillment of the requirements for the degree of Doctor of
Philosophy.
December, 1990
a
2!
Lslxa^
Dean,
Colleg
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f- Medicine
Dean, Graduate School


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81,9(56,7< 2) )/25,'$


UNIVERSITY OF FLORIDA
3 1262 08554 4939


DIRECT MEASUREMENT OF THE
ERROR FREQUENCY BOTH
POLIOVIRUS RNA POLYMERASE
IN VITRO AND IN VIVO
By
CAROL D. WARD
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
1990

ACKNOWLEDGMENTS
There are numerous people that I would like to acknowledge who
have helped me get through graduate school. First and foremost is my
husband, Paul Kroeger, who always encouraged me and had confidence that
I would finish graduate school if that was what I really wanted. His
belief in me was unfaltering which helped me through numerous tough
times.
I would also like to acknowledge my past and present lab coworkers
for their help and perhaps more importantly for their friendship and
their sense of humor. Thanks are also extended to the members of my
committee for their suggestions and ideas. Of course my thanks go to my
mentor, Bert Flanegan, who stuck it out with me through tough times and
always tried to be encouraging. His optimism was much needed to
counterbalance my somewhat pessimistic outlook on life.
Lastly, I would like to extend my thanks to Parker Small for his
genuine concern in both the education and well-being of graduate
students. His door was always open and his insight proved invaluable.
11

TABLE OF CONTENTS
page
ACKNOWLEDGMENTS ii
ABSTRACT v
CHAPTERS
1 INTRODUCTION 1
Background 1
Fidelity of DNA Polymerases 2
Fidelity of RNA Polymerases 4
Rapid Evolution of RNA Viruses 6
Poliovirus Structure 9
Poliovirus RNA Polymerase 11
2 METHODS 14
Enzymes 14
Radiolabeled Compounds 14
Oligoribonucleotide Primers 14
Misincorporation Assays Using 3H and 32P-labeled
Nucleotides 15
Variations on Standard Double Label Experiments 15
RNA Digestion with PI Nuclease 17
High Voltage Ionophoresis 17
Determination of Apparent Km's for Ribonucleotides 17
Cell Culture 18
Purification of Poliovirion RNA 18
Oligodeoxyribonucleotides 18
Labeling of Poliovirion RNA Using [32P]P04 19
Isolation of RNA Oligonucletides with One Internal
G from 32P-labeled vRNA 19
Poliovirus Specific Transcripts 21
Isolation of 5' End-labeled RNA Oligonucleotide
with One Internal G 21
RNase T1 Digestions 23
Gel Purification of Oligonucleotides 23
5' End-labeling 25
RNA Sequencing 25
Gel Electrophoresis 26
iii

3 DETERMINATION OF POLIOVIRUS RNA POLYMERASE ERROR
FREQUENCY IN VITRO 27
Introduction 27
Results 28
Discussion 43
4 DETERMINATION OF THE POLIOVIRUS RNA POLYMERASE ERROR
FREQUENCY IN VIVO 47
Introduction 47
Results 48
Discussion 63
5 CONCLUSIONS AND PERSPECTIVES 75
Factors Affecting Poliovirus Polymerase Error
Frequency 75
Models for DNA Polymerase Base Selection 76
Evolution Rates of Poliovirus 78
Master Sequence Theory 78
RNA Genomes vs. DNA Genomes 80
Future of RNA Viruses 81
REFERENCES 82
BIOGRAPHICAL SKETCH
89

Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy
DIRECT MEASUREMENT OF THE POLIOVIRUS RNA POLYMERASE
ERROR FREQUENCY BOTH IN VITRO AND IN VIVO
by
Carol D. Ward
December, 1990
Chairman: James B. Flanegan
Major Department: Immunology and Medical Microbiology
The error frequency of the poliovirus RNA dependent RNA polymerase
was directly measured both in vivo and fn vitro. Purified polymerase
was used to copy poly(A), poly(C), and poly(I) templates in vitro. and
the error frequency was measured by determining the amount of a
noncomplementary ribonucleotide incorporated relative to the total
amount of ribonucleotide incorporated. The Michaelis constants (Km)
were determined for each ribonucleotide and were as follows: UTP = 10
/iM, ATP — 7 /iM, CTP = 6 /¿M, and GTP = 5 /jM. Changing the relative
concentrations of the ribonucleotide substrates had a significant effect
on the error frequency. Increasing the ratio of complementary to
noncomplementary ribonucleotide substrates from 1:1 to 10:1 resulted in
a 10-fold decrease in the error frequency. Decreasing the ratio from
1:1 to 0.1:1 had no effect. Substituting Mn+2 for Mg+2 was found to
increase the error frequency 2-fold. Changes in the template, the
reaction temperature, and the noncomplementary ribonucleotide substrate
v

used in the reaction had little effect on the error frequency.
Depending on the specific reaction condition, the error frequency varied
from 8.3 X 10"5 to 4.8 X 10'3, a change of 65-fold.
The polymerase error frequency was measured in vivo at eight
different sites in the poliovirion RNA. This assay involved the
measurement of changes in specific G residues to A, C, or U. Sites were
selected that represented both conserved and variable sequences. The
error frequencies determined in these experiments were similar to the
values determined in vitro and ranged from 9 X 10"4 to 5 X 10'3. No
correlation was observed between the error frequency and the variable
vs. conserved sites in the viral genome. Thus, the high mutation rate
that is observed at specific sites in the viral genome is apparently a
result of the high error frequency of the polymerase and the selection
for these changes at the phenotypic level.
vi

CHAPTER 1
INTRODUCTION
Background
RNA versus DNA Polymerases
The central role of both RNA and DNA polymerases is to catalyze
the accurate template-directed incorporation of NTP's or dNTP's,
respectively, in a 5'->3' direction into a growing strand of nucleic acid
in accordance with Watson-Crick base pairing. Both RNA and DNA are used
for the storage of genetic information; however, all known eukaryotic
cellular organisms use DNA for the storage of their genetic information.
DNA is much more stable than RNA and is known to be replicated with high
fidelity. One can imagine that the large genomes of eukaryotic
organisms would need to be replicated with high fidelity to ensure their
perpetuation as a homogeneous species. A small number of errors may be
permitted to promote species evolution, but accuracy is important to
produce viable genome copies. Rates of evolution of cellular genes
average 10'9 substitutions per site per year, in part due to elaborate
proofreading and repair mechanisms (Li et al., 1985; Britlen, 1986). On
the other hand, viral RNA genomes are an average of only 3 to 30
kilobases in length, and RNA viruses are known to evolve at rates a
million-fold higher than their DNA hosts (Holland et al., 1982; Gojobori
and Yokoyama, 1985; Barrel, 1971). Such small genomes probably cannot
afford to invest the space needed for such elaborate repair mechanisms
1

2
and it may be more advantageous for them to be able to evolve rapidly.
As all viruses are intracellular parasites, variability is probably of
utmost importance.
Fidelity of DNA Polymerases
In Vivo Error Rates of Animal Cell DNA Polymerases
The in vivo error rates of animal cell DNA polymerases average 10'8
- 10'11 per incorporated nucleotide (Fowler et al., 1974; Drake, 1990).
This is in sharp contrast to their Tn vitro error rates which vary from
10'4 - 10’5 (Kunkel et al. , 1981; Kunkel and Loeb, 1981). Many models
have been proposed to explain the kinetic mechanism by which DNA
polymerases achieve their high fidelity. All of these models involve
selection of the correct dNTP and the removal of misincorporated
nucleotides by a 3'-+5' exonuclease. Presumably, one reason for the high
error rate of these polymerases in vitro is due to the lack of any
associated exonuclease activity with the purified polymerases.
Kinetic Mechanism of Escherichia coli DNA Polymerase I
Recently much work has been done to determine the kinetic
mechanism by which DNA polymerase I from Jh_ coli achieves its high
fidelity (Kuchta et al., 1987, 1988). A three-step mechanism has been
proposed. The first step involves nucleotide discrimination from a
reduced rate of phosphodiester bond formation for incorrect nucleotides,
with a small contribution from selective dNTP binding. This allows a
discrimination level of approximately 104 - 106-fold. The second step
involves a slow dissociation of the incorrect DNA from the polymerase
which in conjunction with the 3'->5' exonuclease allows a discrimination
level of approximately 4 - 60-fold. The last step involves the slow

3
polymerization of the next correct dNTP onto the mismatch which again
allows for correction by the 3'-+5' exonuclease. This contributes to the
fidelity another 6 - 340-fold. Taken all together, the error rate of
the polymerase would be in the right range for what has been measured in
vivo. Also the lack of any associated exonuclease activity in vitro
would explain its higher error rate.
Factors Affectine Fidelity of DNA Polymerases
The difference in free energy between correct and incorrect base
pairings is estimated to be from one to three kcal/mol (Loeb and Kunkel,
1982). This would predict an error frequency of approximately 1 per 100
nucleotides for nonenzymatic polymerization of oligonucleotides.
Obviously DNA polymerases must enhance this fidelity even in the absence
of 3'-+5' exonucleases. It has been seen that the error rates of
eukaryotic DNA polymerases are directly proportional to the ratio of
correct to incorrect nucleotide substrates (Seal et al., 1979). It is
likely, therefore, that fluctuations in nucleotide pools in vivo would
increase the error frequency of the polymerase. Metal ions have also
been seen to increase the error frequency of DNA polymerases (Sirover
and Loeb, 1977). The ions Mn2+, Co2+, Ni2+, Zn2+, and Be2+ all increase
the error frequency of DNA polymerases, although presumably in
different ways (Sirover and Loeb, 1976). While some can form a complex
with different DNA polymerases, changes in error frequencies at low Mn2+
concentrations (less than 100 ¿¿M) correspond to changes in the binding
of the template and not the enzyme (Beckman et al., 1985). Sirover and
Loeb also report a positive correlation between metal ions that increase

4
the error frequency of DNA polymerases and those reported to be mutagens
in vivo (Sirover and Loeb, 1976).
Fidelity of RNA Polymerases
Measurements of RNA Polymerase Error Frequencies
Little work has been done to measure the error frequencies of RNA
polymerases. This is largely due to the lack of purified RNA
polymerases that can be used in an in vitro system. The error frequency
for vesicular stomatitis virus (VSV) polymerase has been measured in
vivo and in vitro and ranged from 1 to 4 X 10'4 (Steinhauer and Holland,
1986). These measurements were made by direct determination of the
level of substituted bases at given positions in the viral genome by
using a specific ribonuclease, RNase T1. For the 11 kilobase genome of
VSV, this would mean that every member of a plaque purified population
would differ from other genomes of that same plaque at a number of
different nucleotide positions if there was no selective pressure. With
an error frequency of 1 X 1CT4 and an 11 kilobase genome, the average
genome would have 1.1 mistakes, with 33% of the population having no
mistakes. They concluded, therefore, that the preservation of a
consensus sequence must be due to a strong biological selection for the
most fit and competitive representatives of the population.
Terminology
One must be careful when talking about fidelity of polymerases to
distinguish between evolutionary rates and error frequencies.
Evolutionary rates measure the number of mutations over time which
become fixed or dominant. Error frequencies, or mutation rates, are the
frequency of a mutation event which would be represented by the

5
misincorporation during a single round of RNA replication. Most studies
only measure viable mutation rates. Mutant frequency is the proportion
of a certain mutant appearing in an RNA population. Therefore, high
error frequencies or mutation rates are not always reflected in high
evolutionary rates. This is largely dependent upon the environmental
conditions under which a virus is replicating. Often there is a
predominating wild type sequence because the variants have no
competitive advantage over the wild type sequence. This is clearly seen
with poliovirus and other RNA viruses. While poliovirus is seen to
undergo rapid changes while replicating in humans (Kew et al., 1981;
Minor et al., 1986), it is remarkably stable in tissue culture (Parvin
et al., 1986). VSV has been seen to undergo changes during dilute
passages in tissue culture, only to revert back to wild type in later
passages (Spindler et al., 1982). Influenza virus, on the other hand,
is seen to undergo rapid changes in vitro even in conditions under which
poliovirus and VSV are seen to remain stable (Brand and Palese, 1980;
Parvin et al., 1986).
Reversion Rate of Point Mutations
The reversion rate of a point mutation is actually the measure of
a mutant frequency or the frequency of a substitution at a particular
position. A mutant frequency may reflect the mutation rate if two
conditions are met. First, the mutation must be essentially neutral
under permissive conditions, and, second, the amplification of the
mutant is completely wiped out under nonpermissive conditions.
Obviously these conditions can only be met for conditionally lethal
mutants. The reversion rate for an extracistronic point mutant of

6
bacteriophage Q£ has been calculated to be about 10"4 (Batschelet et al.,
1976; Domingo et al., 1976). While this is not a conditionally lethal
mutant, the authors were able to calculate a selective value of the
mutant compared to wild type and used this in calculating their
reversion rate.
Fidelity of Reverse Transcriptases
Retrovirus RNA dependent DNA polymerases have been purified and
used in vitro to determine their error frequencies which are quite high.
Misincorporation rates on homopolymeric templates have shown error
frequencies in the range of 10'3 to 104 (Battula and Loeb, 1974; Sirover
and Loeb, 1977; Loeb and Kunkel, 1982; Preston et al., 1988; Roberts et
al., 1988; Takeuchi et al., 1988). The error frequencies have also been
measured in vitro on heteropolymeric DNA sequences. It was seen that
the accuracy of the reverse transcriptase was dependent on the sequence
replicated and was in the range of 103 to 10"4 (Richetti and Buc, 1990).
Similarly, the frequency of point mutations in retroviruses have been
seen to arise at the same rate (Gopinathan et al., 1979; Kunkel et al.,
1981; Loeb and Kunkel, 1982; Preston et al., 1988). Reverse
transcriptases are DNA polymerases; however, there is no known
exonuclease activity associated with them and they copy RNA templates.
Retroviruses are also seen to exhibit high evolutionary rates, similar
to that of RNA viruses (Coffin et al., 1980; Gojobori and Yokoyama,
1985).
Rapid Evolution of RNA Viruses
Measurement of RNA Virus Evolution Rates
Evolution rates are defined as the rate at which viable mutations
accumulate in the genome. It has been seen that the rate of evolution

7
of RNA genomes is much higher than that seen with DNA genomes. Numerous
methods have been used to measure the evolution rates of RNA viruses.
Common methods include the rates of mutation to monoclonal antibody
resistance, measurement of changes in RNase T1 oligonucleotide maps,
drug resistance mutation rates, reversion rates of point mutations, and
direct RNA sequencing. Obviously, only viable mutant frequencies and
mutation rates are measured using these techniques.
Poliovirus Evolution Rates as Measured by Monoclonal Antibody Resistance
Poliovirus variants resistant to monoclonal antibody
neutralization have been seen to arise at the rate of 10"4 to 10"5 for
Mahoney type 1 (Emini et al., 1984a, 1984b). Similar measurements have
been made for Leon type 3 virus and the attenuated Sabin type 3 vaccine
strain derived from it (Minor et al., 1983). These variants arose at
the rate of 104 to 10’5 for the Sabin strain, whereas mutants from the
Leon strain arose 10 times more frequently. Most of these mutations
appear to be point mutations in the capsid proteins.
Poliovirus Evolution Rates as Measured by T1 Oligonucleotide Mapping
Poliovirus evolution rates have also been measured numerous times
by RNase T1 oligonucleotide mapping. In one study clinical isolates
were followed during a 13-month epidemic (Nottay et al., 1981). These
isolates showed continual mutation and selection during replication in
humans which resulted in the fixation of about 100 nucleotide changes,
or 1 to 2% of the genome bases. Similarly, changes in T1
oligonucleotide maps have been measured in vaccine-associated cases of
paralytic poliomyelitis (Kew et al., 1981). Most isolates appeared to
be multisite mutants which ranged from less than 10 nucleotide changes

8
to greater than 100 nucleotide changes. These changes again represent
up to 1 to 2% of the genome bases, presumably during replication in only
one or two people compared to the previously mentioned 13-month epidemic
of wild type virus.
Poliovirus Evolution Rates as Measured by Guanidine Resistance Mutations
Poliovirus mutants' resistance to guanidine have been seen to
arise at the rate of approximately 3 X 10‘5 (Holland et al., 1973). This
may, however, represent changes at more than one site on the genome thus
underestimating the rate of a single mutation. Recently measurements
were made for the conversion of a guanidine-dependent poliovirus to
guanidine resistance (de la Torre et al., 1990). This represents a
single site reversion and was found occur at a mutation frequency of 2.5
X 10'3 to 2 X 10'4.
Poliovirus Evolution Rates as Measured by Point Mutation Reversions
Much controversy revolves around the mutation rates measured for
poliovirus by point mutation reversions. The previously mentioned point
mutation reversion of guanidine dependence to guanidine resistance
showed a high mutation frequency of 10’3 to 10’4. Another study involved
inserting 72 nucleotides into the 5' noncoding region of poliovirus
which contained an in-frame start codon (Kuge et al., 1989). This
mutant had a small plaque phenotype and reversion to large plaques was
seen to have single nucleotide changes. These revertants were seen to
arise at the rate of 10’2. This high number was explained by the high
viability of the large plaque phenotype mutant to the small plaque
insertion mutants and the existence of sister clones in the small
plaques. A third study argues that a single base revertant of

9
poliovirus arises at the low rate of 2.5 X 10'6 (Sedivy et al., 1987).
In this study a mutant was constructed such that a serine codon was
converted to an amber codon. The virus was then grown on suppressor
positive cells and then titered on both suppressor positive and
suppressor negative cells to determine the reversion frequency. Only
one of three possible nucleotide changes was seen to generate revertants
so the mutation rate was estimated to be approximately 10’5. It should
be noted, however, that the mutant virus had a 2-fold longer eclipse
period and a 10-fold lower burst size on the suppressor cell line,
probably due to incomplete suppression. Obviously this mutation was not
neutral in permissive conditions and thus is not a direct measurement of
the error frequency of the polymerase.
Poliovirus Error Frequency as Determined by Direct Sequencing
Parvin et al. sequenced a segment of the viral protein 1 (VP1)
gene of poliovirus type 1 from multiple individual virus plaques that
had all descended from a single plaque (Parvin et al., 1986). No
mutations were detected in over 95 X 103 nucleotides sequenced. A
neutral mutation rate of less than 2.1 X 10’6 was calculated for this
site compared to 1.5 X 10'5 for the NS gene of influenza virus. Lethal
mutations would not have been scored in this study. This low rate could
be due to the need for conservation at VP1, a lower polymerase error
frequency at this site, or due to purifying selection differences.
Poliovirus Structure
RNA Genome and Viral Proteins
The genomic RNA of poliovirus has a 3' terminal poly(A) sequence
and a 5' covalently linked protein called VPg. For the Mahoney strain

10
of type 1 poliovirus, the viral RNA (vRNA) is about 7500 bases in
length and contains one long open reading frame from base 743 to 7370.
The exact functions of the 5' and 3' noncoding regions are not known,
but they may be involved in replicase recognition and binding to
initiate RNA synthesis. The 5' noncoding region also contains sequences
that are required for ribosome binding and the initiation of
translation. The four capsid protein sequences are located at the amino
terminus of the polyprotein. The capsid proteins VP1, VP2, and VP3 have
a common structural motif (Hogle et al., 1985, 1987; Hogle and Filman,
1989) . They contain a core sequence which is composed of an eight-
stranded antiparallel beta barrel with two flanking alpha helices.
Differences in the amino acid sequences of the capsid proteins of type 1
Mahoney and type 3 Leon strains have been mapped to the inner and outer
surfaces of the capsid proteins, but not to the core sequences. This
suggests a strong selective pressure for constraints imposed by protein
folding and assembly. Two proteases, 2A and 3C, are also encoded in the
long open reading frame (Hanecak et al., 1982; Toyoda et al., 1986).
These proteases are involved in the cleavage of the polyprotein into the
smaller viral gene products, with most of the cleavages performed by 3C
(Ypma-Wong et al., 1988). Other viral protein products include 3AB, a
membrane associated precursor of VPg, VPg(3B) and 3D, the viral RNA
polymerase. The roles of nonstructural viral proteins 2B and 2C in
poliovirus replication are unclear at this time. Guanidine
hydrochloride is known to block poliovirus replication. It appears that
2C is responsible for guanidine sensitivity (Anderson-Sillman et al.,
1984; Baltera and Tershak, 1989). Guanidine resistance and dependence,

11
as well as host range mutants, have also been mapped to 2C (Yin and
Lomax, 1983; Anderson-Sillman et al., 1984; Pincus and Wimmer, 1986;
Pincus et al., 1986; Baltera and Tershak, 1989; de la Torre et al.,
1990). These data suggest that 2C has a role in RNA synthesis.
Homology Between Serotypes
The homology between the three serotypes of poliovirus is 71% at
the nucleotide level (Toyoda et al., 1984). Of these substitutions, 80%
of them are silent. It is interesting that the type 3 Sabin vaccine
P3/Leon/12ab differs from its neurovirulent parent P3/Leon/37 by only 10
point mutations (Stanway et al., 1984; Almond et al., 1987a, 1987b,
Westrop et al., 1989). The recent poliomyelitis outbreak in Finland was
from a type 3 virus that had 95.5% homology with P3/Leon/37 at the amino
acid level (Hughes et al., 1986). There were 3 amino acid substitutions
and 6 amino acid substitutions, however, at two major antigenic
determining sites. These sites are normally highly conserved in wild
strains of poliovirus.
Poliovirus RNA Polymerase
Poliovirus RNA Polymerase and its Role in Replication
Poliovirus RNA replicates in the cytoplasm of infected cells using
a virus-coded RNA-dependent RNA polymerase. A soluble and template-
dependent form of the poliovirus polymerase, 30^ , has been purified
from cytoplasmic extracts of infected cells (Van Dyke and Flanegan,
1980). Highly purified forms of the polymerase synthesize full-length
copies of poliovirion RNA and other polyadenylated RNAs, but only in the
presence of an oligo(U) primer (Flanegan and Baltimore, 1977; Flanegan
and Van Dyke, 1979; Tuschall et al., 1982; Van Dyke et al., 1982). The

12
requirement for a primer is unusual for RNA polymerases, but this need
can be eliminated by the addition of a cellular protein component termed
"host factor" in vitro (Dasgupta et al., 1980; Flanegan et al., 1987).
In vitro reactions require all four ribonucleotide triphosphates, Mg2+
or Mn2+, and an oligo(U) primer or host factor. In the presence of host
factor, the largest size of product RNA synthesized is twice the size of
the RNA template (Young et al., 1985). The product RNA is complementary
to and covalently linked to the template RNA (Young et al., 1985, 1986).
The amount, size distribution, and rate of synthesis of product RNA are
dependent of the Mg concentration, pH, and temperature of the in vitro
reaction conditions (Van Dyke et al., 1982). At optimal in vitro
conditions the synthesis rate is approximately 1200 nucleotides per
minute.
Replication of Homopolvmers
Synthetic homopolymers can be copied by the polio polymerase in
vitro in the presence of the correct primer. These template primers
include poly(A):oligo(U), poly(C):oligo(I), and poly(I):oligo(C)
(Tuschall et al.,1982). Poly(U):oligo(A) can only be copied to a very
small extent and no activity is noted on poly(G):oligo(C). Template
binding studies indicate that the polymerase binds to poly(G) the best,
followed by poly(U), poly(C), poly(I), and lastly poly(A), the exact
opposite order of the templates that it copies best (Oberste and
Flanegan, 1988) .
Studies on the Fidelity of Poliovirus Polymerase
Studying the fidelity of poliovirus RNA replication has a unique
advantage over studies with most other RNA animal viruses because the

13
polymerase has been purified in a soluble and template-dependent form
and can therefore be used to directly measure error frequencies in vitro
on different RNA templates. By copying synthetic homopolymers using the
purified polymerase and differentially labeled complementary and
noncomplementary ribonucleotide substrates, one can directly measure the
error frequency of the polymerase. This procedure eliminates the bias
of converting viable mutation rates to error rates of the polymerase
since lethal mutations need not be accounted for. The in vitro
procedure also has the distinct advantage of allowing one to vary the
reaction conditions and observe any changes in the error frequency.
Templates may also be varied to determine if certain base pair
mismatches are allowed more than others.

CHAPTER 2
METHODS
Enzymes
Proteinase K was obtained from Boehringer Mannheim in lyophilized
form. It was dissolved and stored in 10 mM Tris*HCl, 1 mM EDTA, and 25%
glycerol at a concentration of 10 mg/ml. Ribonuclease T1 was obtained
from Calbiochem Corporation in lyophilized form. It was dissolved and
stored in 50 mM Tris at 10 u/¿¿l.
Radiolabeled Compounds
All radiolabeled compounds were obtained from Amersham except for
[32P]P04. This compound was obtained from either ICN Biomedicals, Inc.,
or New England Nuclear. The radiolabeled nucleotide [y-32P]ATP was
obtained in aqueous solution with a specific activity of 3000 Ci/mmole.
All other radiolabeled compounds were in a 50% ethanol solution which
was vacuumed down to less than half volume before using. The
radiolabeled compounds [a-32P]UTP, [q-32P]CTP, [a-32P]ATP, and [a-32P]GTP
all had a specific activity of 410 Ci/mmole. The specific activity of
[3H]UTP ranged from 37 to 45 Ci/mmole, [3H]CTP was 20 Ci/mmole, and
[3H]GTP ranged from 10 to 15 Ci/mmole.
Oligoribonucleotides Primers
Oligoribonucleotides were generated from homopolymers as described
by Bock. Briefly, 1 - 2 mg of homopolymer (poly(C), poly(I), or
poly(U)) was incubated at 90°C for 40 minutes in 0.5 ml 0.1 M
14

15
NH4HC02• NH40H (pH 10.0). The pH was then adjusted to 1.0 by the addition
of 0.5 ral 1 M HC1 and incubated at 20°C for 20 minutes. This solution
was then neutralized by the addition of 1 ml of 1 M Tris*HCl, pH 8, and
ethanol precipitated. The oligonucleotides were then treated with calf
intestinal phosphatase (Boehringer Mannheim) as described (Maniatis et
al., 1982).
Misincorporation Assays Using 3H and 32P-labeled Nucleotides
The standard reaction mixture (final volume 30 ¡j.1) contained 50 mM
HEPES (N-2-hydroxyethyl piperazine-N2 - ethane sulfonic acid) pH 8.0, 3
mM MgCl2, 10 mM DTT, 2.5 ¿¿g poly(A) , 1.25 /¿g oligo(U), and 3 ¿¿1 Fraction
IV(HA) polymerase purified as described (Young et al.,1986). For the
correct ribonucleotide, 10 fid of [5,6-3H]UTP (36 Ci/mmole) was added,
and for the incorrect ribonucleotide 20 /¿Ci of either [q-32P]ATP, [a-
32P]GTP, or [a-32P]CTP was added. Unlabeled ribonucleotides (Calbiochem
Corporation) were added to make the reaction mixtures equimolar (7.2 /iM)
with respect to complementary and noncomplementary ribonucleotides. The
reactions were incubated for 1 h at 30°C. The labeled product RNA was
precipitated in 7% trichloroacetic acid (TCA) and 1% sodium
pyrophosphate, collected on membrane filters, and counted.
Variations on Standard Double Label Experiments
Template
Variations of the misincorporation assay included substituting
poly(C):oligo(I) for the template:primer and using [3H]GTP as the
correct ribonucleotide or substituting poly(I):oligo(C) for the template
and using [3H]CTP as the correct ribonucleotide.

16
Concentration of Correct and Incorrect Ribonucleotides
All variations on the ratio of correct to incorrect
ribonucleotides were done on a poly(A) template with [32P]CTP used as
the incorrect ribonucleotide. At a 10:1 ratio of correct to incorrect
ribonucleotides, 50 /¿Ci of [3H]UTP was used and unlabeled UTP was added
to a final concentration of 74 /¿M for the correct ribonucleotide. For
the incorrect ribonucleotide, 20 /¿Ci of [32P]CTP was used and unlabeled
CTP was added to a final concentration of 7.4 /¿M. Alternatively, 10 /¿Ci
of [3H]UTP was used and 10 /tCi of [32P]CTP was used making the
concentrations of correct and incorrect ribonucleotides 7.4 /¿M and 0.74
/¿M respectively. At 0.1:1 ratio of correct to incorrect
ribonucleotides, 1 /¿Ci of [3H]UTP was used as the correct ribonucleotide
and 20 /¿Ci of [32P]CTP was used and unlabeled CTP was added to a final
concentration of 7.4 /¿M.
Mg+2 versus Mn+2
Another variation including substituting 0.5 mM MnCl2 for 3 mM
MgCl2. This was done under both equimolar concentration of correct to
incorrect ribonucleotides and at 10:1 ratio of correct to incorrect
ribonucleotides with the correct ribonucleotide being at 7.4 /¿M and the
incorrect ribonucleotide being at 0.74 /¿M.
Temperature
The error frequency was determined at 30°C, 37°C, and at 42°C.
This was done at equimolar concentrations and at a 10:1 ratio of correct
to incorrect ribonucleotides as described above.

17
RNA Digestion with PI Nuclease
Product RNA was synthesized on a poly(A) template as described
above using 10 ¿tCi each of [a-32P]UTP and 10 ¿tCi of either [a-32P]ATP,
[a-32P]GTP, or [a-32P]CTP. The labeled product RNA was phenol:chloroform
extracted, ethanol precipitated, dissolved in 100 ¿¿1 of 0.1 M NaCl, 1 mM
EDTA, 10 mM Tris HC1, pH 7.6, and run through a G-50 "spun column" as
described (Maniatis et al., 1982) to remove unincorporated labeled
nucleotides. The labeled RNA was ethanol precipitated and dissolved in
15 ¿ti 10 mM CH3C00Na* 3H20, pH 6.0 containing 1.5 units of PI nuclease
(Bethesda Research Laboratory). The sample was then heat denatured at
100°C for 10 minutes, cooled to 37°C, and another 1.5 units of PI
nuclease in 15 ¿tl of 10 mM CH3C00Na*3H20, pH 6.0 was added to the
sample. The sample was incubated for 1.5 h at 37°C to complete the
digestion.
High Voltage Ionophoresis
Ionophoretic separation of the PI nuclease digested product RNA
was performed on Whatman 3MM paper at pH 3.5 as described (Barrel, 1971;
Rose, 1975). The 32P-labeled ribonucleoside monophosphates were located
by autoradiography, cut out from the paper and counted in 5 ml of
Aquasol-2 scintillation fluid.
Determination of Apparent Km's for Ribonucleotides
Poliovirion RNA (1 ¿íg) was copied in the presence of 10 ¿tCi of
[32P]UTP and enough unlabeled UTP to make the final concentrations 0.8
¿iM, 5 ¿tM, 10 ¿tM, 20 ¿tM, 40 ¿tM, or 80 ¿tM. The other three
ribonucleotides were also present at 500 ¿tM. Standard reaction
conditions were used (3 mM MgCl2, 50 mM HEPES pH 8.0) except that the

18
total volume was 150 ¡i\ and 30 n 1 aliquots were taken every 10 minutes
for 30 minutes and TCA precipitated. Moles of product made were then
calculated and divided by time to determine the initial velocities. The
initial velocities and substrate concentration were then plotted on a
Lineweaver-Burk double reciprocal plot to determine the apparent Km for
UTP. The apparent Km's for the other three ribonucleotides were
determined in a similar manner.
Cell Culture
HeLa cells were maintained in suspension culture and infected with
poliovirus type 1 (Mahoney strain) as previously described
(Villa-Komaroff et al., 1974). Briefly, cells were washed with Earles
saline, pelleted, and infected with poliovirus at an MOI = 20 for 30 min
at room temperature. The cells were then diluted to 4 X 106 cells/ml in
Eagles modified minimal media with 7% sera (5% bovine calf and 2% fetal
calf). Cells were infected for 6 h at 37°C, washed with Earles saline,
and frozen at -20°C.
Purification of Poliovirion RNA
Poliovirion RNA (vRNA) was isolated from infected cells as
described (Young et al., 1986). Briefly, poliovirions were banded in
cesium chloride density gradients, phenol extracted three times,
chloroform extracted three times, and the vRNA ethanol precipitated.
01igodeoxvribonucleotides
All oligodeoxyribonucleotides were synthesized on an Applied
Biosystems model 380A or 380B automated DNA synthesizer, using
phosporamadite chemistry. These were then gel purified on a 20%
polyacrylamide, 7 M urea gel. A list of the oligodeoxyribonucleotides

19
used are shown in Table 2-1. These were all complementary to
poliovirion RNA at the nucleotides shown in parentheses and contained a
3' terminal poly(A)15 sequence.
Labeling of Poliovirion RNA Using F32P1PO,
HeLa cells growing in suspension culture were centrifuged and
washed three times with phosphate-free modified Eagles media buffered at
pH 7.2 with 25 mM HEPES and 10 mM TES (N-tris(hydroxymethyl)methyl-2-
aminoethane sulfonic acid). The cells were infected with poliovirus
(MOI = 20) and allowed to incubate for 30 minutes at room temperature
(approximately 25°C). The cells were then diluted to 4 X 106 cells/ml
in phosphate free media with 7.0% sera (5% bovine calf sera and 2% fetal
calf sera) and dialyzed against phosphate-free Earles balanced salt
solution (116 mM NaCl, 5 mM KC1, 26 mM NaHC03, pH 7.5). After 15 min,
Actinomycin D was added at a concentration of 5 /¿g/ml. At 30 min 600
/uCi/ml [32P]P04 was added. At 6 h the cells were centrifuged and washed
with Earles saline. The RNA was then purified as described (Young et
al., 1986).
Isolation of RNA Oligonucleotides with One Internal G from 32P-labeled vRNA
Poliovirion RNA (10 ¿ig) uniformly labeled with [32p] P04 was
hybridized with 0.3 ng of DNA oligonucleotide in 30 n\ of 10 mM Tris
HC1, pH 7.5, 500 mM NaCl, and 4 mM MgCl2, for 3 h at 50°C. The hybrids
were digested with 6 units of RNase T1 for 2 h at 50°C. The RNA was
treated with proteinase K by diluting the sample with 120 /¿I 0.5% SDS
buffer (100 mM NaCl, 10 mM Tris, pH 7.5, 1 mM EDTA, .5% SDS) and adding
75 ng proteinase K for 1 hour, phenol:chloroform extracted two times and
ethanol precipitated. The RNA:DNA duplex was then purified on a 20%

20
Table 2-1. List of DNA OliEonucleotides and Protected RNA Sequences
Name vRNA bases
DNA seauence 5'-*3'/RNA seauence 3'-+5'
BF8 6862 - 6895
CAAGTTGTTAATCATTGAGTTAAAAATTGAAGTGAAAAAAAAAAAAAAA
GUU CAACAAUUAGUAACU CAAUUUUUAACUU CAC
BF9 7269 - 7296
CTGATTTTAGCTAGGAATTTGTTATATTAAAAAAAAAAAAAAA
GACUAAAAUCGAUCCUUAAACAAUAUAA
BF10 5638 - 5671
CTTTAGAGTGATTATAGTGATTTCAAGATTGGTTAAAAAAAAAAAAAAA
GAAAUCUCACUAAUAU CACUAAAGUUCUAAC CAA
BF11 3971 - 3999
CTAGTTATAATAACTAGTGAGGATATGATAAAAAAAAAAAAAAA
GAU CAAUAUUAUUGAU CACUC CUAUACUA
BF25 84 - 109
CTAAGTTACGGGAAGGGAGTATAAAAAAAAAAAAAAAAAAA
GAUUCAAUGCCCUUCCCUCAUAUUUU
BF27 1229-1253
CTAGGTAGTGGTAGTACATATTTTGAAAAAAAAAAAAAAA
GAUCCAUCACCAUCAUGUAUAAAAC
BF30 1429-1450
CTGGTTGTTGTCAGGAGTGAAAAAAAAAAAAAAAAA
GACCAACAACAGUCCUCACUU
BF35 2757-2776
CGTGGTGGAAGCTGGGTTATAAAAAAAAAAAAAAA
GCACCACCUUCGACCCAAUA

21
polyacrylamide gel, denatured, and the RNA oligonucletide was
thenrepurifled on a 20% polyacrylamide, 7M urea gel. The RNA fragment
was then RNase T1 digested for 1 hour at 50°C and the digestion products
were run on a 20% polyacrylamide, 7M urea gel. The radioactivity (cpm)
left in the RNase T1 resistant RNA fragment was then compared to the cpm
in the digestion products to determine the error frequency using an
automated gel scanner (AMBIS or Betagen). This procedure is shown
schematically in Figure 2-1.
Poliovirus Specific Transcripts
DNA plasmids pOF2612 and pOF1205 were used to generate labeled
poliovirus specific RNA transcripts. The plasmid pOF2612 contains a
full-length cDNA copy of the poliovirus genome and pOF1205 contains the
3' terminal nucleotides 6516 - 7440 and a poly(A)83 sequence (Oberste,
1988a). The plasmid p0F2612 was digested with Pvu II which cuts at base
7053 in the poliovirus sequence and pOF1205 was cut with EcoRl which
cuts at the 3' end of poly(A)83. These templates were then transcribed
with SP6 polymerase and 10 /¿Ci of [32P]UTP or [32P]ATP following the
protocol supplied by Promega Biotechnologies Inc. The RNA was DNase
treated, followed by phenol:chloroform extraction and ethanol
precipitation. The amount of RNA transcribed was calculated by TCA
precipitation and counting of a small aliquot of the sample. These
transcripts were then used in hybridization and T1 digestion experiments
as described above for [32P]P04 labeled virion RNA.
Isolation of 5' End-labeled RNA Oligonucleotide with One Internal G
Poliovirion RNA (10 pg) was hybridized and digested as described
above using 0.03 pg 5'end-labeled DNA oligonucleotide and 0.27 ng cold

22
Labeled vRNA
> DNA oligo Poly(A)
Hybridize
T1 Digest
20% Acrylamide Native Gel
A
Isolate RNA-DNA Hybrid
Denature
20% Acrylamide 7M Urea Gel
^ DNA oligo
Poly(A)
Protected RNA oligo
Purify protected RNA oligo
T1 Digest
20% Acrylamide 7M Urea Gel
T1 Resistant Oligo
T1 Digestion Products
Fig. 2-1. Schematic diagram of [32P]P04 labeled vRNA hybridization and
RNase T1 digestion assay to determine error frequency of the polymerase
at specific sites on the genome.

23
DNA oligonucleotide. The hybrid was purified on a nondenaturing 20%
polyacrylamide gel and located by autoradiography. The protected RNA
fragment was 5' end-labeled as described below, DNased with 2 units
DNase (Boehringer Mannheim) in 50 m1 of 100 mM CH3C00Na* 3H20 (pH 4.5), 5
mM MgS04, and purified on a denaturing 20% polyacrylamide, 7 M urea gel.
The protected RNA fragment was then digested with RNase T1, proteinase K
treated, and run on a 20% polyacrylamide, 7M urea gel as described
below. This procedure is shown schematically in Figure 2-2. The
digestion products were analyzed using the Betagen gel scanner.
RNase T1 Digestions
Isolated labeled RNA fragments were ethanol precipitated with 20
Mg glycogen (Boehringer Mannheim) and dissolved in 6 ¡xl of 25 mM sodium
citrate (pH 3.5), 7M urea, 1 mM EDTA, 0.035% xylene cyanol and 0.035%
bromphenol blue. RNase T1 (6 units) was added and incubated at 50°C for
1 h. The sample was then boiled for 1 min and another 6 units of RNase
T1 was added and incubated for another hour at 50°C. Proteinase K
(1 Mg) was then added and incubated at 37°C for another hour. The
reaction mixtures were then boiled, quick chilled on ice, and loaded
directly onto a 20% polyacrylamide, 7M urea gel.
Gel Purification of Oligonucleotides
All oligonucleotides that were gel purified were either located by
autoradiography or UV shadowing. The pieces were cut out of the gel,
crushed, and eluted overnight at 37°C in 150 /ul to 1 ml of water
depending on the size of the gel piece. Large pieces of polyacrylamide
were removed by centrifugation in an eppendorf microfuge and the
supernatant was passed through a sterile disposable polypropelene column

24
vRNA
>» DNA oligo Poly(A)
Hybridize
T1 Digest
20% Acrylamide Native Gel
Isolate RNA-DNA Hybrid
DNase
5' end label
Denature
20% Acrylamide 7M Urea Gel
Poly(A)
Protected RNA oligo
Purify protected RNA oligo
T1 Digest
Proteinase K
20% Acrylamide 7M Urea Gel
T1 Resistant Oligo
T1 Digestion Products
Fig. 2-2. Schematic diagram of 5' end-labeled vRNA hybridization and
RNase T1 digestion assay to determine the error frequency of the
polymerase at specific sites on the genome.

25
with paper disc (Isolabs). This was then ethanol precipitated with 20
Hg glycogen. Fragments eluted from very large gel pieces were extracted
three or four times with absolute ethanol to eliminate any urea.
5' End-labelinE
RNA and DNA fragments were routinely end labeled with 10 /iCi of
[7-32P]ATP using 10 units of T4 polynucleotide kinase (New England
Biolabs). When RNA pieces were to be sequenced, however, 50 ¿tCi was
used instead for each hybridization and digestion that started with 10
Hg poliovirion RNA.
RNA Sequencing
RNase Tl-resistant oligonucleotides were gel purified and ethanol
precipitated with 15 /ig of tRNA. The RNA was divided into five aliquots
which were digested with either RNase T1 (Calbiochem Corporation), RNase
U2 (Bethesda Research Laboratories), RNase PhyM (Bethesda Research
Laboratories), RNase B. cereus (Bethesda Research Laboratory) or RNase
CL3 (Pharmacia). RNase T1 and U2 digestions conditions were 25 mM
sodium citrate (pH 3.5), 7M urea, 1 mM EDTA, 0.035% dyes, with 2
units/ml and 0.5 unit/ml RNase respectively. Digestion conditions for
PhyM were 25 mM sodium citrate (pH 5.0), 7M urea, 1 mM EDTA, 0.035%
dyes, with 100 units/ml RNase. Digestion conditions for B. cereus were
25 mM sodium citrate (pH 5.0) with 200 units/ml RNase B. cereus.
Digestion conditions for CL3 were 10 mM sodium phosphate (pH 6.5), 10 mM
EDTA, and 50 units/ml RNase. All digestions were for 15 min at 55°C
except CL3 which was done at 37°C. Digestions with B. cereus and CL3
were stopped by the addition of 7M urea and 0.035% dyes. All digestions

26
were stopped by freezing on dry ice. Samples were then boiled for 3 min
and quick chilled before loading onto a 20% polyacrylamide, 7M urea gel.
Gel Electrophoresis
All 20% polyacrylamide gels were at a 30:1 ratio of acrylamide to
bisacrylamide. All were run in 100 mM Tris•HCl, 100 mM H3B03, and 2 mM
EDTA (pH 8.0). All long (45 cm) urea gels were prerun at 25 watts, and
all short (20 cm) urea gels were prerun at 15 watts, for a minimum of 1 h.

CHAPTER 3
DETERMINATION OF POLIOVIRUS RNA POLYMERASE ERROR FREQUENCY IN VITRO
Introduction
The poliovirus RNA polymerase has been purified from infected
cells and copies poliovirion RNA and other polyadenylated RNAs in vitro.
Synthetic homopolymers including poly(A):oligo(U), poly(C):oligo(I), and
poly(I):oligo(C) serve as template:primers for the polymerase as well.
By copying synthetic homopolymers with the purified polymerase and
differentially labeled complementary and noncomplementary substrates,
the error frequency of the polymerase can be measured directly. Another
technique that was used in this study involved copying synthetic
homopolymers in the presence of both 32P-labeled complementary and
noncomplementary ribonucleotide substrates and digesting the product RNA
with PI nuclease. PI nuclease digests the RNA to 5'-ribonucleoside
monophosphates which can be separated and counted to determine the error
frequency. Both of these techniques allow one to vary the reaction
conditions and observe how these changes affect the error frequency of
the polymerase. Reaction conditions that were looked at included the
effects of template, substrate, divalent cations, nucleotide substrate
concentration and temperature. Both of these in vitro procedures
eliminate the bias of converting viable mutation rates to error rates of
the polymerase, in contrast to what most in vivo studies measure.
27

28
The error frequency of the poliovirus polymerase has been measured
in vitro on a poly(A) template using differentially labeled
complementary and noncomplementary substrates by Mary Merchant-Stokes
(Merchant-Stokes, 1985). The error frequency was measured at pH 7 or pH
8 and at 3 mM or 7 mM MgCl2. The error frequency ranged from 0.7 X 10'3
to 5.4 X 10'3 depending on the reaction conditions. Increasing the pH
from 7 to 8 increased the error frequency of the polymerase 2 to 3 fold.
Increasing the MgCl2 concentration from 3 mM to 7 mM also increased the
error frequency about 2 fold. A correlation was seen between reaction
conditions that increased the elongation rate of the polymerase with
reaction conditions that increased the error frequency of the
polymerase.
Results
Determination of the Km for Each Ribonucleotide Substrate in the
Polymerase Reaction
One of the variables examined was the effect of nucleotide
concentration on the error frequency of the poliovirus polymerase. When
measuring the effect of nucleotide concentration, one should also take
into account the possible effect of the nucleotide concentration
relative to the Km for that particular nucleotide. It was possible that
using nucleotide concentrations that were far away from the Km might
affect the error rate. In addition, it was important to know if there
were large differences in the Km for the four different ribonucleotide
substrates as this might affect the error rate as well (See Chapter 5
for discussion of the Km Discrimination Model).

29
To determine the apparent Kms for each ribonucleotide, the initial
velocities of the polymerase reaction were measured as a function of the
concentration of each ribonucleotide substrate. This was done at a
variety of concentrations for one nucleotide (0.4 /¿M, 0.8 /iM, 2.5 /¿M, 5
/iM, 10 /tM, 20 /iM, 40 /iM, and/or 80 /iM) while the other three nucleotides
were kept constant at 500 /iM. The amount of ribonucleotide incorporated
in the product RNA was measured as a function of time to determine the
initial velocities (Figures 3-1 and 3-2). The Km for each nucleotide
was then determined by using a Lineweaver-Burk double-reciprocal plot of
the initial velocities vs. the substrate concentration (Figures 3-3 (UTP
and ATP) and Figure 3-4 (CTP and GTP)). The Km values for each
ribonucleotide were as follows: ATP = 10 /iM, UTP = 7 /iM, CTP = 6 /iM,
and GTP = 5 /iM.
Determination of the Polymerase Error Frequency by Using Differentially
Labeled Ribonucleotide Substrates
The error frequency of the poliovirus RNA polymerase was
determined by measuring the rate at which a noncomplementary
ribonucleotide substrate was incorporated into the product RNA using
synthetic homopolymeric RNAs as templates. When poly(A) was used as the
template [3H]UTP was used as the complementary substrate and 32P-labeled
ATP, GTP and CTP were used as the noncomplementary substrates. The
error frequency of the polymerase reaction was defined as the moles of
noncomplementary nucleotide incorporated divided by the total moles of
nucleotide incorporated into the product RNA.
Effect of changing the templates and substrates
The error frequency of the poliovirus polymerase was measured on
poly(A), poly(C), and poly(I) with each of the three different

30
CO
0)
o
E
Q.
â– D
CD
O
CL
V-
O
o
c
CL
5
3
[UTP]
Q .4 uM
♦ .8 uM
a 5 uM
o 10 uM
â–  80 uM
m
O
o
E
CL
â– o
4)
O
CL
k_
O
o
c
a
2
<
[ATP]
â–¡ .4 uM
♦ .8 uM
â–¡ 5 uM
o 10 uM
â–  80 uM
Figure 3-1. Effect of [UTP] and [ATP] on the initial velocity of the
poliovirus polymerase reaction. Initial velocities (v ) of the
polymerase were determined for five different concentrations of UTP (top
graph) or ATP (bottom graph). Standard reaction conditions were used on
vRNA with an oligo(U) primer (3 mM MgCl2, pH 8, 30°C) except that the
initial reaction volume was 150 and 30 /il aliquots were removed and
counted every ten minutes.

31
Cfl
o
o
E
Q.
"O
0)
10
o
Q.
w
O
Ü
c
a
5
O
[CTP]
â–¡ .4 uM
♦ .8 uM
â–¡ 5 uM
o 10 uM
â–  40 uM
w
0)
o
E
Q.
TJ
V
o
a
h_
o
o
c
a.
2
(5
[GTP]
o 2.5 uM
♦ 5 uM
a 10 uM
o 40 uM
Figure 3-2. Effect of [CTP] and [GTP] on the initial velocity of the
poliovirus RNA polymerase reaction. Initial velocities (v ) of the
polymerase were determined at five different concentration of CTP (top
graph) and four different concentrations of GTP (bottom graph).
Reaction conditions were as described in legend to Figure 3-1.
Measurements of (v0) for GTP were repeated and both values were plotted
on the Lineweaver/Burk plot shown in Figure 3-4.

32
1/[UTP]
Fig. 3-3. Determination of Km for ATP aned UTP using Lineweaver-Burke
plots. Double reciprocal plot of v'1 vs. [NTP]‘1 for poliovirus
polymerase where v represents the initial velocity of incorporation for
each NTP (UTP and ATP). Initial velocities were determined at each
concentration using the data shown in Figure 3-1.

33
1/[CTP]
1/[GTP]
Fig. 3-4. Determination of Km for CTP and GTP using Lineweaver-Burk
plots. Double-reciprocal plot of v'1 vs. [NTP]'1 for poliovirus
polymerase where v represents the initial velocity of incorporation for
each NTP (CTP and GTP). Initial velocities were determined at each
concentration using the data shown in Figure 3-2.

34
noncomplementary ribonucleotide substrates for each template (Table 3-1).
All measurements were made using equimolar amounts of complementary and
noncomplementary ribonucleotide substrates. There was no
significant difference between the error frequency on poly(A) and
poly(C), however the error frequency did appear to be slightly lower on
poly(I) (less than 2-fold). There appeared to be no significant
difference between the four different noncomplementary ribonucleotide
substrates. It was not possible to determine the error frequency on
poly(U) or poly(G) because of very low levels of polymerase activity on
these templates.
Effect of changing the nucleotide concentration
By far the largest factor found to affect the error frequency of
the polymerase was the concentration of ribonucleotide substrates. All
assays were done on a poly(A) template using [3H]UTP as the correct
ribonucleotide and [32P]CTP as the incorrect ribonucleotide substrate.
The UTP concentration was varied from 0.74 /¿M to 74 /¿M and the CTP
concentration from 0.74 /¿M to 7.4 /¿M. It was found that as the ratio of
correct to incorrect ribonucleotide substrate was increased from 1:1 to
10:1, the error frequency decreased approximately 10-fold. If the ratio
of correct to incorrect ribonucleotide substrates was decreased from 1:1
to 0.1:1, the error frequency remained the same (Table 3-2). It was
interesting to find that decreasing the ratio of correct to incorrect
nucleotides had no effect on the error frequency. This suggested that
the error frequency had reached a maximum value that was not farther
increased by decreasing the relative concentration of the complementary
substrate. In marked contrast, increasing the relative concentration of

35
Table 3-1. Error Frequency of the Poliovirus RNA Polymerase
Reaction8
Noncomplementary
Complementary
Errorb
Conditions
Substrate
Substrate
Frequency
Poly(A)
[32P ] ATP
[3H]UTP
3.8
+
0.6 X 10'3
3 mM MgCl2
[32P] CTP
[3H]UTP
2.9
+
0.4 X 10'3
pH 8
[32P]GTP
[3H]UTP
3.8
+
0.5 X 10'3
Poly(C)
[32P ] ATP
[3H]GTP
2.9
+
1.2 X 10'3
3 mM MgCl2
[32P] CTP
[3H]GTP
4.8
+
2.0 X 10'3
pH 8
[32P]UTP
[3H]GTP
2.5
+
0.2 X 103
Poly(I)
[32P] ATP
[3H]CTP
2.2
+
0.5 X 10'3
3 mM MgCl2
[32P]GTP
[3H] CTP
1.6
+
0.4 X 10'3
pH 8
[32P ] UTP
[3H]CTP
1.9
+
0.6 X 10'3
a Reaction conditions were as described in Materials and Methods except
the final volume of the poly(A) reactions was 50 /i 1. In the poly(C)
and poly(I) reactions, the final volume was 30 /¿I and the total
nucleotide concentrations were 25.6 /¿M and 16.6 /¿M respectively. The
error frequency on poly(A) was determined by Mary Merchant-Stokes
(Merchant-Stokes, 1985).
b The error frequency was defined as the pmoles of noncomplementary
nucleotide incorporated divided by the total pmole of nucleotide
incorporated into product RNA. For example, 432,802 cpm [3H]UMP
(2.02 X 104 cpm/pmole) and 8,038 cpm [32P]AMP (1.33 X 105 cpm/pmole)
were incorporated at pH 8, 3 mM MgCl2. A counting efficiency for 3H
of 0.33 was assumed.

36
Table 3-2. Effect of Nucleotide Concentration on the Error Frequency of
the Polymerase
[UTP]
[ CTP ]
[UTP]/[CTP]
Error
Frequency3
0.7 /tM
7.4 /iM
0.1
3.2 ±
0.8 X 10'3
7.4 /iM
7.4 /iM
1
2.9 ±
0.4 X 10'3
74.0 /iM
7.4 /xM
10
2.0 ±
0.8 X 10'4
7.4 /iM
0.7 /iM
10
4.4 ±
1.6 X 10'4
3 All reactions were on poly(A), 3 mM MgCl2, pH 8.

37
the complementary substrate resulted in a significant decrease in the
error frequency. There did not appear to be a direct relationship
between the error frequency and the Km's of the nucleotide substrates.
For example, when the concentration of the noncomplementary substrate
was held constant at a concentration near its Km, decreasing the
concentration of the complementary substrate from a concentration near
its Km value to one-tenth of its Km had no effect, whereas increasing
its concentration to ten times the Km value decreased the error
frequency. Thus, it appears that the ratio of the nucleotide substrate
concentrations and not their absolute concentrations has the greatest
affect on the error frequency.
Effect of MnCl„
It was found that substituting .5 mM MnCl2 as the divalent cation
resulted in a 2-fold increase in poliovirus polymerase error frequency
relative to 3 mM MgCl2 (Table 3-3) . All of these assays were done on a
poly(A) template using [3H]UTP as the correct ribonucleotide substrate
and [32P]CTP as the incorrect ribonucleotide substrate. This
measurement was made at both equimolar ratios of complementary to
noncomplementary ribonucleotide substrate and at a 10:1 ratio of
complementary to noncomplementary ribonucleotide substrate (7.4 /xM UTP,
0.7 /xM CTP) . Again it was seen that increasing the ratio of
complementary to noncomplementary ribonucleotide substrate to 10:1
resulted in about a 9-fold decrease in the error rate.
Effect of temperature
Changes in temperature of the reaction conditions appeared to have
no effect on the error frequency of the poliovirus polymerase. (Table 3-4).

38
Table 3-3. Effect of MnCl2 on Poliovirus Polymerase Error Frequency
Nucleotide Ratio3
Compl./Noncompl,
10:1
10:1
1:1
1:1
Divalent Cationb
3 mM MgCL,
0.5 mM MnCl2
3 mM MgCl2
0.5 mM MnCl2
Error Frequency
0.8 ± 0.1 X 10~4
1.4 ± 0.3 X 10'4
7.2 ± 2.0 X 10'4
12.6 ± 2.4 X 10'4
Increase in
Error Frequency
1
2
9
16
3 At 10:1 ratio of complementary to noncomplementary substrates, 7.4 /zM
UTP and 0.7 /zM CTP were used respectively. At 1:1 ratio of
complementary to noncomplementary substrates, 7.4 /zM UTP and 7.4 /zM CTP
were used respectively.
b All reactions were on poly(A) at pH 8.

39
Table 3-4. Effect of Temperature on Polymerase Error Frequency
Nucleotide ratio3
Compl./Noncompl. 30°C
37°C
42°C
1:1 7.2 ± 2.0 X
10:1 8.3 ± 0.8 X
io-4
IO'5
7.4 ± 1.9 X
1.3 ± 0.3 X
IO'4
IO4
6.8 ± 2.2 X IO’4
9.3 ± 5.8 X IO’5
3 At 1:1 ratio off complementary to noncomplementary nucleotide, 7.4 ¿¿M
UTP and CTP were used. At 10:1 ratio of complementary to
noncomplementary nucleotide 7.4 ¿¿M UTP and 0.7 ¿¿M CTP were used. All
reactions were on poly(A), 3 mM MgCl2, pH 8.

40
The error frequency was determined at 30°C, 37°C, and 42°C in reactions
that contained 3 mM MgCl2, pH 8. These assays were also done on a
poly(A) template at both equimolar ratios of complementary to
noncomplementary ribonucleotide substrates and at a 10:1 ratio of
complementary to noncomplementary ribonucleotide substrate (7.4 ¿¿M UTP,
0.7 /xM CTP) .
It should be noted that the absolute value for the error frequency
determined at a nucleotide ratio of 1:1 and at 3 mM MgCl2, pH 8 in
Tables 3-3 and 3-4 was about 4-fold lower than the values obtained under
similar conditions in Tables 3-1 and 3-2. The reason for this change in
error frequency is not clear, however all experiments were internally
controlled so that relative changes in the error frequency due to
changes in the reaction conditions were real. The change in ratio of
nucleotide concentration done during the MnCl2 and temperature variation
experiments still resulted in a 9-fold relative change in the error
frequency.
Error Frequency as Determined by Pi Nuclease Digestion
Poly(A) templates were copied with both 32P-labeled complementary
and noncomplementary ribonucleotide substrates. The product RNA was
then separated from unincorporated labeled ribonucleotides and digested
to completion with PI nuclease. PI nuclease digests RNA to
ribonucleoside 5'-monophosphates and should yield a 32P-labeled
noncomplementary ribonucleoside only if it was actually incorporated
into the product RNA (Figure 3-5). These digestion products were
separated by high voltage ionophoresis and visualized by
autoradiography (Figure 3-6). The ribonucleoside 5'-monophosphates were

5 oligoU
2> PolyA
5'
polymerase
PPP u
+ PPPC ^ 3,
5 oligo UpUpUpUpUpUpCpUpUpU
3» PolyA
iPI
Nuclease
5' 'i-* 4-* ^ * '!•* 4-* 4-* ¿ *
oligo U pU pU pU pU pU pC pU pU pU
3/ PolyA
i
5' *
5 pc
5' *
5 PU
5'
3'
5'
Figure 3-5. Diagram showing 5'-ribonucleoside monophosphates recovered from product RNA
digested with PI nuclease. Illustrated is the synthesis of product RNA on a poly(A) template
the presence of an oligo(U) primer by the polymerase in the presence of [^PJUTP and [32P]CTP.
The expected PI cleavage of the labeled product RNA is also shown.

42
• pG
pA
pC
% ori
Figure 3-6. High-voltage ionophoretic separation of PI nuclease
digestion products. RNA was synthesized in the presence of [32P]UTP and
the following 32P-labeled noncomplementary ribonucleotides: [32P]ATP
(lane 1), [32P]CTP (lane 2), [32P]GTP (lane 3). Lane 4 is a marker lane
containing 5'-ribonucleoside monophosphates UMP (pU), GMP (pG), AMP
(pA), and CMP (pC).

43
then cut out and counted to determine the error frequency. All of these
assays were done at 3 mH MgCl2, pH 8. The error frequency ranged from
2.0 X 10'3 to 4.8 X 10'3 (Table 3-5).
Discussion
The results of these experiments indicated that the error
frequency of the poliovirus RNA polymerase was affected by changes in
the in vitro reaction conditions. A number of variables
including the type of divalent cation, the ratio of complementary to
noncomplementary ribonucleotide substrates, and the type of template
copied had an effect on the error frequency of the polymerase.
Previously it was seen that the pH and the MgCl2 concentration had an
effect on the error frequency. The only variables measured that had no
effect on the error frequency were the temperature and the
noncomplementary ribonucleotide substrate used in the reaction.
The type of homopolymeric RNA template used in the reaction seemed
to have little or no effect on the error frequency of the polymerase.
This was similar to the results that have been observed with reverse
transcriptases (Battula and Loeb, 1974). It should be noted, however,
that alternating copolymers are typically copied with more fidelity by
DNA polymerases, and that the fidelity of reverse transcriptase appears
to be sequence dependent on heteropolymeric templates, but not
homopolymeric templates (Battula and Loeb, 1974; Loeb and Kunkel, 1982;
Richetti and Buc, 1990).
Changing the ribonucleotide that was added as the noncomplementary
substrate did not affect the error frequency of the poliovirus
polymerase. The molecular mechanisms involved in the selection of

44
Table 3-5. Error Frequency as Determined by PI Nuclease Digestion
Reaction
Conditions
Noncomplementary
Substrate
Complementary
Substrate
Error Frequency
Poly(A)
[32P] ATP
[32P]UTP
2.9 ± 2.8 X 10'3
3 mM MgCl2
f 32p ] CTP
[32p]utp
4.8 ± 2.4 X 10'3
pH 8
[32P]GTP
[32P]UTP
2.0 ± 2.4 X 10'3

45
correct versus incorrect nucleotides by polymerases is still not
understood and needs further exploration. While numerous models have
been proposed, the data does not clearly choose one model above all
others. For a discussion of these models, see Chapter 5.
Substituting MnCl2 for MgCl2 was found to increase the error
frequency of poliovirus polymerase by 2-fold. Manganese chloride
decreases the fidelity of DNA polymerases from 2 to 25 fold, depending
on the polymerase (Sirover and Loeb, 1977; Beckman et al., 1985). This
decrease in fidelity has been attributed mostly to the binding of the
Mn+2 to the template. This is thought to facilitate the formation of
noncomplementary bases pairs during polymerization by changing the
hydrogen bonding properties of the nucleotides. At higher
concentrations of Mn+2 it is also possible that interactions with the
enzyme or nucleotide substrates affect the error frequency.
The largest influence on the error frequency of the polymerase was
the relative concentrations of the ribonucleotide substrates used in the
in vitro reactions. At equimolar concentrations or less of
complementary to noncomplementary ribonucleotide substrates, the error
rate was at its highest value of about 3 x 10"3. With a 10-fold increase
in the ratio of complementary to noncomplementary ribonucleotide
substrates, the error rate correspondingly decreased 10-fold. This was
observed whether the complementary nucleotide concentration was equal to
the Km of the substrate or 10-fold above it. Therefore, the ratio of
correct to incorrect substrates appeared to be more important in
determining the error rate of the polymerase than substrate
concentration relative to the Km.

46
It was rather surprising to find that changing the temperature did
not affect the error frequency of the poliovirus polymerase. Since the
temperature is known to affect the elongation rate of the polymerase,
there does not appear to be a simple relationship between the elongation
rate of the polymerase and the error frequency of the polymerase.
Apparently, temperature does not affect the base selection process.
Measurements of the error frequency after digesting the product
RNA with PI nuclease supported the data obtained by using differentially
labeled ribonucleotide substrates. One disadvantage of the assay that
uses differentially labeled substrates is that any contaminating 32P-
labeled complementary nucleotide that might be present in the 32P-
labeled noncomplementary ribonucleotide could be incorporated and
counted as an error. The PI nuclease digestion assay eliminates this
problem as all substrates are 32P-labeled. The error rate determined in
this manner varied from 2.0 X 10'3 to 4.8 X 10'3. These numbers were in
general agreement with the error frequencies determined under the same
reaction conditions by differentially counting the 3H and 32P-labeled
product RNA.
Overall, the error frequency of the poliovirus RNA polymerase
measured in vitro ranged from 8.3 X 10'5 to 4.8 X 10‘3, a change of 65-
fold. Thus, while any one change in reaction conditions had a
relatively small effect, together they can cause a large change in the
error frequency of the poliovirus RNA polymerase.

CHAPTER 4
DETERMINATION OF THE POLIOVIRUS RNA POLYMERASE ERROR FREQUENCY IN VIVO
Introduction
The recent work by Steinhauer and Holland (Steinhauer and Holland,
1986) shows that it is now possible to measure the error frequency of
viral RNA polymerases both in vitro and in vivo. They developed a
technique that can be modified to measure the error frequency of most
RNA viruses. This technique involves the measurement of the error
frequency at a specific nucleotide in the viral RNA. Although there are
some limitations on which nucleotides can be used in this assay, it is
possible to select specific sites from different regions of the viral
genome. It does not require that the viral RNA be infectious unlike
most other in vivo techniques that only measure viable mutation rates or
error frequencies. The viral RNA used in this assay can either be
purified from the cytoplasm of infected cells or from purified virions.
Steinhauer and Hollands measurements made with VSV using this technique
both in vivo and in vitro indicate a high error frequency of the VSV RNA
polymerase, which ranged from 1 X 10"4 to 4 X 10‘4 (Steinhauer and
Holland, 1986).
Much controversy exists over the error frequency of the poliovirus
polymerase. While no direct measurements have been made, the
evolutionary rate of poliovirus has been measured in many different ways
with apparently conflicting results (for details, see Chapter 1). What
47

48
is now needed is a direct measurement of the error frequency of
poliovirus RNA polymerase in vivo. This was the primary objective of
the studies described in this chapter. I have adopted and modified the
technique of Steinhauer and Holland to measure the error frequency at
specific sites in purified poliovirion RNA.
Results
Purification of RNA Oligonucleotides
The basic approach used to determine the polymerase error rate in
vivo was a procedure that was a modification of the method previously
described by Steinhauer and Holland (Steinhauer and Holland, 1986).
Briefly, 32P-labeled RNA was hybridized to a synthetic DNA
oligonucleotide and digested with RNase T1. The protected RNA
oligonucleotide that was complementary to the DNA was isolated by gel
purification and then digested with RNase T1. The rate of change in the
single G residue found in the protected fragment was determined by
quantitating the amount of the protected fragment that was resistant to
digestion (for details, see Chapter 2 and figure 2-1). The technique
was initially developed on pOF1205 transcript RNA, which consists of the
3' terminal 1000 bases of poliovirus RNA. The transcript RNA was
labeled with either [32P]UMP or [32P]AMP, hybridized to a synthetic DNA
oligonucleotide, digested with RNases T1 and U2 or RNase T1 alone, and
run on a 20% polyacrylamide, 7 M urea gel. A protected oligonucleotide
of the expected size (34 nucleotides) was recovered from the [32P]UMP-
labeled RNA that was hybridized to the synthetic DNA oligonucleotides
(Figure 4-1, lanes IB and 1C). This oligonucleotide was not present
when the RNA was digested in the absence of the synthetic DNA (Figure 4-1,

49
lanes 1A and ID). When the transcript RNA was labeled with [32P]AMP,
the same protected oligonucleotide was present (Figure 4-1, lanes 2B and
2C). In this case, however, it was apparent that a ladder of labeled
poly(A) fragments was also present (Figure 4-1, lanes 2A to 2D). This
result indicated that the protected RNA oligonucleotide that was
isolated from this gel was contaminated with a poly(A) fragment of the
same size. This was confirmed in other experiments where the [32P]UMP-
labeled 34mer was further characterized. The labeled 34mer was gel
purified, digested with RNase T1, and run on a denaturing polyacrylamide
gel. Two major bands of the expected size were observed along with a
small amount of the 34mer that was resistant to digestion (Figure 4-2).
The 34mer was isolated from the gel and 5'-end labeled with [y-32P]ATP
and polynucleotide kinase. The sequence was then determined by
enzymatic sequencing procedures as described in Chapter 2 (Figure 4-3).
It was clear that the Tl-resistant 34mer was in fact contaminated with a
large amount of poly(A). This would be an obvious problem in any
experiments where the 3' terminal poly(A) sequence actually was labeled
(for example, [32P] P04-labeled virion RNA). This problem was not dealt
with by Steinhauer and Holland since VSV RNA is not polyadenylated.
Avoiding the coisolation of poly(A) with the protected RNA
oligonucleotide was solved by initially isolating the RNA-DNA duplex on
a nondenaturing 20% polyacrylamide gel (Figure 4-4). The RNA-DNA duplex
comigrates on this gel with poly(A) fragments which are much longer than
the protected RNA oligonucleotide which is part of this duplex. The
duplex was isolated from the nondenaturing gel, denatured with urea, and
run on a 20% polyacrylamide, 7 M urea gel to separate the protected RNA

50
1A IB 1C ID 2A 2B 2C 2D 3C
Figure 4-1. Isolation of RNA oligonucleotide protected from BF8 after
RNAse T1 digestion. 32P-labeled poliovirus specific RNA was transcribed
from pOF1205 DNA. The RNA was labeled with either [32P]UMP (lanes 1A -
ID) or [32P]AMP (lanes 2A - 2D). THe RNA was either directly digested
with RNase T1 and U2 (lanes A) or RNase T1 (lanes D) or was first
hybridized with synthetic DNA oligonucleotide BF8 and then digested with
RNases T1 and U2 (lanes B) or RNase T1 (lanes C). Poliovirion RNA
labeled in vivo with [32P]P04, hybridized with BF8, and digested with
RNase T1 is shown in lane 3C.

51
12 3 4 5 6
Figure 4-2. Final RNase TI digestion products of RNA oligonucleotide
protected by BF8. The oligonucleotide was isolated from a gel similar
to that shown in Figure 4-1 where pOF1205 transcript RNA was labeled
with [32P]UMP. The initial hybridization and RNase T1 digestion
conditions were as follows: 37°C (lane 1), 55°C (lane 3), 50°C (lane
4), 45°C (lane 5), 40°C (lane 6). The RNA in lane 2 was hybridized at
55°C and then digested at 37°C. All final T1 digestions were done at
55°C.

52
O
+
O O < <
4>
X)
"O
<0
3
+
O O < <
Figure 4-3.
BF8. First
Figure 4-2.
lanes 3, 4,
RNA sequencing of TI resistant oligonucleotide protected by
set of digests and ladder (five lanes) are from lane 2 in
Second set of digests (last four lanes) are from pooling
5, and 6 in Figure 4-2.

53
Figure 4-4. RNA hybridization and TI digestion products run on a native
20% polyacrylamide gel. Lanes 1 and 2 are [32P]UMP labeled pOF1205
transcripts, lane 3 is [32P]P04 labeled vRNA, and lane 4 contains
[32P]AMP labeled pOF1205 transcripts. Lane 1 was not hybridized with
BF8.

54
oligonucleotide from the DNA oligonucleotide and any contaminating
poly(A) (Figure 4-5). It should be noted that the DNA was engineered to
have a 15 base long poly(A) tail so that it would separate from the RNA
oligonucleotide on a denaturing gel. In earlier studies, I found that
it was not possible to quantitatively remove all of the DNA from this
duplex by a simple digestion with DNase. Thus, the two-step gel
purification procedure was adopted. The purified RNA oligonucleotide
was isolated from the second gel and then digested with RNAse T1 to
determine the error rate. The error rate was defined as the
radioactivity (cpm) in the RNase T1 resistant band relative to the
radioactivity in the two oligonucleotides that were the RNase T1
digestion products.
Polymerase Error Frequency Determined by Using i32P1PC^-Labeled
Poliovirion RNA
The error frequency has been measured at two sites in the
poliovirus genome using the procedure described above and [32P]P04-
labeled RNA. The two sites were at nucleotide 6883 in the 3Dpo1 coding
sequence and at nucleotide 5648 in the 3Cpro coding sequence. DNA
oligonucleotides BF8 and BF10 were used to isolate RNA oligonucleotides
that contained these two sites (see Table 2-1 for the exact sequences).
The RNA oligonucleotide protected by BF8 was digested to completion with
RNase T1 and was analyzed by gel electrophoresis (Figure 4-6). A very
small but detectable amount of the oligonucleotide was resistant to
digestion (Figure 4-6, lane 1). The radioactivity (cpm) recovered in
this resistant band represented 4.3 X 10'3 of the total radioactivity
recovered in all three bands (i.e., the two major bands representing the
digestion products and the resistant band). Thus, the polymerase error

55
1 2 3
Figure 4-5. Separation of RNA oligonucleotide protected by BF8 from
contaminating poly(A) by gel purification. The RNA-DNA duplex was first
isolated by gel purification on a nondenaturing 20% polyacrylamide gel.
The duplex was then denatured and run on a 20% polyacrylamide, 7 M urea
gel. Poliovirion RNA was labeled in vivo with [32P]P04 (lane 1) SP6
polymerase transcripts of p0F1205 DNA were labeled with either [32P]UMP
(lane 2) or [32P]AMP (lane 3).

56
1 2
Figure 4-6. Final RNase T1 digestion products of RNA oligonucleotide
protected by BF8 run on a 20% polyacrylamide, 7 M urea gel. The
protected oligonucleotide from poliovirion RNA labeled with [32P]P04 was
run after RNase T1 digestion (lane 1). BF8 protects a 34mer and digests
to a 22mer and a 12mer. Lane 2 shows undigested marker.

57
frequency at this site (i.e., nucleotide 6883) was 4.3 X 10'3 (Table 4-
1). The error frequency determined using BF10 was about the same with a
value of 0.9 X 10'3 (Table 4-1).
The major drawback to this approach to determine the error
frequency was the relatively low specific radioactivity of the labeled
virion RNA and the large amount of radioactivity (50 mCi) that was
required to label the vRNA synthesized in infected cells. For these
reasons, a second approach was used to determine the in vivo error
frequency of the poliovirus RNA polymerase.
Polymerase Error Frequency Determined Using 5' End-Labeled
Oligonucleotides from Poliovirion RNA
The error frequency at eight different sites in the poliovirus
genome were determined using the 5' end-labeling technique. These sites
were located in constant and variable regions of the poliovirus genome
(illustrated in Figure 4-7). The technique used is summarized briefly
here, for details see Chapter 2. Poliovirion RNA was hybridized to a 5'
end-labeled DNA oligonucleotide and digested with RNase T1. This hybrid
was then purified on a nondenaturing 20% polyacrylamide native gel
(Figure 4-8). The hybrid was then 5' end-labeled with [y-32P]ATP and
polynucleotide kinase, treated with DNase and run on a denaturing 20%
polyacrylamide, 7 M urea gel (Figure 4-9). The band representing the
protected RNA oligonucleotide was isolated from the gel, digested with
RNase Tl, and run on a 20% polyacrylamide, 7 M urea gel. The digestion
products were quantitated by an AMBIS or Betagen gel scanner.
The final digestion products for the RNA oligonucleotides
protected by BF8 and BF10 are shown in Figure 4-10. The final digestion

58
Table 4-1. Poliovirus Error Frequency at Specific Sites
Labeling Method0
Protecting DNA oligonucleotide3 [32P]P04 5'-end label
BF8: 6862-6895 3D conserved 3.2 X 10"3
4.7
X
10'3
BF9:
7269-7296
3D
conserved
4.4
X
10‘3
BF10:
5638-5671
3C
conserved
0.7 X 10'3
CM
on
X
10'3
BF11:
3971-3999
2B
conserved
3.5
X
10'3
BF25:
84-109
5'NC
conserved
3.8
X
CO
ó
t-H
BF27:
1229-1253
VP2
conserved
5.0
X
10'3
BF30:
1429-1450
VP2
variable
4.6
X
co
b
1—1
BF35:
2757-2776
VP1
variable
3.2
X
10'3
aNumbers of protecting DNA oligonucleotide represent the nucleotides
protected in the poliovirus genome. 3D, 3C, 2B, VP2, and VP1 refer to
the genes encoded at these sites on the poliovirus genome. 5'NC refers
to the 5' non coding region. Conserved and variable refer to whether
these sites on the genome are known to change (variable) or not
(conserved).
bNumbers indicate corrected values based upon RNA sequencing and
redigesting with RNase T1 and are an average of at least 3 experiments.

59
VPg —
Polyprotein
P'NC
Poly(A)
VP4
iBvpg
VP2
VP3
VP1
O
Q-
<
_C\L
2B
2C
3A
3Cpro
3Dpo'
T-
oo
''t
CO
00
CO CO
o
CO
CO
CD
00
co co
T-
C\J
â– 't
h-
O)
CO
00 CVJ
T-
T—
CvJ
CO
ir>
co r-
c
c
V
V
c
c
c c
Figure 4-7. Schematic diagram of the poliovirus genome with the various
sites examined by RNase T1 digestion indicated. C represents constant
regions on the genome and V represents variable regions on the genome.

60
Figure 4-8. Isolation of RNA oligonucleotides protected from BF27 and
BF10 after RNase T1 digestion. DNA oligonucletides BF27 and BF10 were
5'-end labeled and either run directly on the gel (Lanes 1 and 6,
respectively), or hybridized with virion RNA and digested with RNase T1.
BF27 hybridization and digestion products were run in lanes 2 and 3.
BF10 hybridization and digestion products were run in lanes 4 and 5.

61
12 3 4
Poly(A)
{
DNA
Figure 4-9. Isolation of 5'-end labeled protected RNA oligonucleotides.
RNA/DNA hybrids were isolated from a gel as shown in figure 4-8, 5'-end
labeled, digested with DNase, denatured with urea, and run on a 20%
polyacrylamide, 7M urea gel. Note the coisolation of poly(A) as well as
residual DNA that was not completely DNased.

62
Figure 4-10. Final RNase TI digestion products of 5'-end labeled
protected oligonucleotides. Lane 1 is from an oligoribonucleotide
protected by BF8 and lane 2 is from an oligoribonucleotide protected by
BF10. Lanes 3 and 4 are undigested markers.

63
products for the RNA oligonucleotides protected by BF9, 11, 25, 27, 30
and 35 are shown in Figures 4-11 through 4-16. All duplicate lanes
shown in Figures 4-11 through 4-16 represent reactions with various RNA
preparations that were separately hybridized and digested. All of these
assays were repeated at least three times for each oligonucleotide and
the average error frequency was determined (Table 4-1). This average
error frequency takes into account a correction factor that reduced the
error frequency by 25%. The RNAse T1 resistant oligonucleotides
protected by BF8 and BF11 were isolated from the gel and their
nucleotide sequence was determined (Figures 4-17 and 4-18,
respectively). While both of these sequencing gels clearly show the
presence of the three other nucleotides besides G, there is still a G
band present which represented approximately 25% of the total
radioactivity in the sequencing bands. In other experiments, the RNase
T1 resistant oligonucleotides were redigested with RNase T1. The amount
of RNase T1 resistant fragment that could be redigested with T1 ranged
from 10 to 50%. For this reason an average correction factor of 25% was
used.
Discussion
A modification of the technique developed by Steinhauer and
Holland was used to measure the in vivo error rate of poliovirus
polymerase. There were many unexpected problems encountered during the
modification of this technique. The first unforeseen problem was the
coisolation of contaminating poly(A) sequences with the RNA-DNA duplex.
The average size of poly(A) on poliovirion RNA is 75 - 100 nucleotides
and the heteroduplexes isolated were significantly smaller, 20 - 49

64
•AAUAUAACAAAUUCCUAGCUAAAAUCAG 28mer
^T1 Digest
*AAUAUAACAAAUUCCUAG CUAAAAUCAG
18mer 10mer
1 2 3
4 5
28
Figure 4-11. Final RNase T1 digestion products of 5'-end labeled
oligonucleotides protected by BF9. Lane 1 is undigested marker.
BF9 protects a 28mer and digests to a 18mer (labeled) and a lOmer
(unlabeled). Lanes 2-5 are all separate reactions.

65
‘AUCAUAUCCUCACUAGUUAUUAUAACUAG 29mer
^T1 Digest
‘AUCAUAUCCUCACUAG
16mer
UUAUUAUAACUAG
13mer
Figure 4-12. Final RNase T1 digestion products from 5'-end labeled
oligonucleotide protected by BF11. Lane 4 is undigested marker.
BF11 protects a 29mer and digests to a 16mer (labeled) and a 13mer
(unlabeled). Lanes 1-3 are all separate reactions.

66
‘UUUUAUACUCCCUUCCCGUAACUUAG 26mer
^T1 Digest
‘UUUUAUACUCCCUUCCCG UAACUUAG
18mer 8mer
1 2 3 4 5
26
1 8
Figure 4-13. Final RNase T1 digestion products of 5'-end labeled
oligonucleotide protected by BF25. Lane 1 is undigested marker.
BF25 protects a 26mer and digests to a 18mer (labeled) and a 8mer
(unlabeled). Lanes 2-5 are all separate reactions.

67
‘CAAAAUAUG UACUACCACUACCUAG 25mer
^T1 Digest
‘CAAAAUAUG UACUACCACUACCUAG
9mer 16mer
Figure 4-14. Final RNase T1 digestion products of 5'-end labeled
oligonucleotide protected by BF27. BF27 protects a 25mer and digests to
a 9mer (labeled) and a 16mer (unlabeled). Lanes 1-4 are all separte
reactions.

68
*UUCACUCCUG AC AAC AACCAG 21mer
^T1 Digest
‘UUCACUCCUG ACAACAACCAG
10mer 11mer
1 2
Figure 4-15. Final RNase T1 digestion products of 5'-end labeled
oligonucleotide protected by BF30. Lane 1 is undigested marker,
protects a 21mer and digests to a lOmer (labeled) and a llmer
(unlabeled). Lanes 2 and 3 are separate reactions.
BF30

69
*AUAACCCAGCUUCCACCACG 20mer
^T1 Digest
‘AUAACCCAG CUUCCACCACG
9mer 11mer
1 2
20
3
Figure 4-16. Final RNase T1 digestion products of 5'-end labeled
oligonucleotide protected by BF35. Lane 3 is undigested marker,
protects a 20mer and digests to a 9mer (labeled) and a llmer
(unlabeled). Lanes 1 and 2 are separate reactions.
BF35

70
Figure 4-17. Enzymatic sequencing of the oligoribonucleotide protected
by BF8 before (right four lanes) and after RNAse T1 digestion (left four
lanes). The arrow marks the change in the single G residue to the other
three ribonucleotides.

71
Figure 4-18. Enzymatic sequencing of the oligoribonucleotide protected
by BF11 before (right five lanes) and after RNase T1 digestion (left
five lanes). The arrow marks the change in the single G residue to the
other three ribonucleotides.

72
nucleotides in length. Apparantly the poly(A) sequence in poliovirion
RNA is much more heterogeneous in length than previously thought.
Although only a small fraction of the virion RNA molecules contain short
poly(A) sequences, the amount present is still relatively large compared
to the amount of the RNase T1 resistant oligonucleotide which is
recovered in these experiments.
A second problem encountered was the quantitative removal of all
of the protecting DNA oligonucleotide. Original experiments used a DNA
oligonucleotide of the same length as the protected RNA oligonucleotide.
The hybrids were digested with DNase to remove the DNA and then digested
with RNase T1. These RNA oligonucleotides could not be digested to
completion as a result of residual DNA oligonucleotide. This problem
and the preceding problem of coisolating contaminating poly(A) sequences
were solved by engineering a poly(A)15 tail on the protecting DNA
oligonucleotide. A two step purification was then used to isolate the
heteroduplex on the first gel, and then a second gel was used to
separate the RNA from the protecting DNA oligonucleotide and any
contaminating poly(A) derived from the virion RNA.
One final problem encountered was an apparent gel shifting of the
RNase T1 digestion products. Unexpected high bands appeared on the
final RNase T1 digestion gels which could not be accounted for. They
were resistant to DNase and additional RNase T1 treatment. These bands
were apparently caused by contaminating proteins since they disappeared
with the addition of a proteinase K digestion step.
The error frequency of the poliovirus RNA polymerase was
determined at eight different sites on the poliovirus genome and ranged

73
from 0.9 to 5.0 X 10‘3. These values are dependent on the complete
digestion of protected RNA oligonucleotide by RNase T1. Therefore, it
was important to determine how efficient this digestion was. For this
reason RNase T1 resistant oligonucleotides were sequenced when possible.
A large limiting factor on sequencing, however, was the small amount of
radioactivity in these RNase T1 resistant bands. When sequencing could
not be done, the RNase T1 resistant bands were redigested with RNase T1
and quantitated to determine the percent that could be redigested. This
percentage varied from 10 - 50%. Sequencing showed approximately 25% of
the RNase T1 resistant band contained a G residue For these reasons a
correction factor of 25% was used. With this correction factor taken
into account, the error frequency determined in vivo was in agreement
with the numbers determined in vitro.
The error frequency did not vary significantly between the eight
different sites measured on the poliovirus genome. Two of the sites
examined are known to mutate rapidly under selective pressure. Site
1439 (BF30) is located within the E-F loop of VP2. This site has been
observed to change rapidly when the virus is grown in the presence of
monoclonal antibodies which resulted in amino acid changes from aspartic
acid to asparagine and histidine (Page et al., 1988). Site 2765 (BF35)
is located in the B-C loop of VP1 in which host range mutants have been
mapped (Murray et al., 1988). This G is not conserved between serotypes
1 and 3 which results in an amino acid change from alanine to proline.
The loop regions of the capsid proteins in general are less conserved
between the the three serotype of poliovirus. The other six sites are
relatively conserved between the three serotypes of poliovirus. Sites

74
6883 (BF8) and 7286 (BF9) are located within BD*501 and any change in
these G residues would result in amino acid changes from methionine to
isoleucine and from alanine to serine, proline or threonine,
respectively. Site 5648 (BF10) is located within 3Cpr0 and a change in
this G residue would result in a change from glutamic acid to lysine,
glutamine, or a stop codon. Site 3986 (BF11) is located within 2B and
would result in an amino acid change from valine to isoleucine, leucine,
or phenylalanine. Site 101 is located in the 5' non-coding region of
poliovirus. This site, as well as the four other conserved sites
mentioned so far, appears to be very important in the replication of
poliovirus. The sixth conserved site, 1236 (BF27) is located within /3
barrel B of VP2. All of the p barrels of the capsid proteins are highly
conserved between the serotypes of poliovirus. A change in this G
residue would result in an amino acid change from methionine to
isoleucine.
While changes at these G residues would cause relatively minor
changes in some cases (BF35, from one nonpolar amino acid to another)
and relatively major changes in others (BF10, from a negatively charged
amino acid to either a positively charged amino acid, a polar uncharged
amino acid or a stop codon), there was no significant difference in the
error frequency determined at these different sites. They ranged from
.9 X 10'3 to 5 X 10'3, approximately a five-fold difference. These small
differences did not correlate with conserved and variable regions of the
viral genome. Thus, it appears that the error frequency of the
poliovirus polymerase is relatively constant at G sites across the
poliovirus genome.

CHAPTER 5
CONCLUSIONS AND PERSPECTIVES
Factors Affecting Poliovirus Polymerase Error Frequency
I found that a number of factors can affect the poliovirus RNA
polymerase error frequency which ranged in vitro from 8 X 10'5 to
5 X 10'3. These factors included the type of divalent cation, the
relative concentrations of correct and incorrect nucleotide substrates,
and the type of template used. The specific ribonucleotide that was
used as the incorrect substrate and changes in the temperature had
little or no effect on the error frequency of the polymerase. How these
factors exert their effect on nucleotide selection by the polymerase is
unknown at this time. The error frequency of the poliovirus polymerase
measured in vivo was also found to be similar to the in vitro values and
ranged from 9 X 10‘4 to 5 X 10'3. No significant difference was found in
the error frequency at eight different sites in the poliovirus genome.
Two sites were selected because one is known to rapidly mutate when
grown under certain environmental conditions (in the presence of
monoclonal antibody) and the other is known to vary between the three
serotypes of poliovirus. The other six sites were selected as conserved
sites since they are unchanged between the three serotypes of
poliovirus. Overall, my results indicated that there was no significant
difference in the error frequency between the variable and the conserved
75

76
sites. This suggests that the variation observed in nature is due to
selection at the phenotypic level.
Models for DNA Polymerase Base Selection
There are several models that have been proposed to explain how
DNA polymerases select the correct incoming nucleotide. The first of
these models, the "Km Discrimination Model" (Goodman et al., 1977),
proposes that the difference in free energy between correct and
incorrect base pairings is increased in the presence of polymerases.
This model assumes that the rates for binding are the same for both
correct and incorrect nucleoside triphosphates, and that the
discrimination is based on differences in the dissociation rates. This
theory would then predict that the error rate is proportional to the
differences in the Km for the correct and incorrect nucleotides.
A second model, the "Conformation Model", proposes that the
polymerase changes in conformation with each nucleotide addition step,
which affects base selection. Most polymerases appear to have one site
for the binding of all four nucleotides. Therefore, one would predict
that these polymerases must have a mechanism to accommodate the
different structures of the nucleotides and base pairs. This model
predicts that a difference in Vmax for the correct and incorrect
nucleotides would be proportional to the error rate of the polymerase
(Watanabe and Goodman, 1982) .
Finally a third model, the "Energy Relay Model" (Hopfield, 1980),
proposes that the energy released by phosphate bond cleavage is used by
the polymerase to proofread the insertion of the next nucleotide. This
model would predict that the incorporation of the first nucleotide is

77
more error prone than the following nucleotides. This model was not
supported by the use of reversion frequency assays performed by Abbotts
and Loeb using mammalian DNA polymerases (Abbotts and Loeb, 1984). Nor
was it supported by Kuchta (Kuchta et al., 1988) using an elongation
assay with DNA polymerase I.
Because no large differences were seen between the
misincorporation rates of the four different ribonucleotides both on
homopolymeric templates in vitro and on heteropolymeric templates in
vivo. it is not clear that one of these models is greatly supported or
negated over another. It should be pointed out however that Michaelis
constants can vary significantly at specific sites and that these were
not measured. Factors that were found to affect the error frequency of
the poliovirus polymerase certainly may affect the conformation of the
polymerase and its intereaction with the nucleotide substrates.
However, it was also clear in sequencing the T1 resistant
oligonucleotides that all four nucleotides were present and one
incorrect base did not predominate over the others as one might expect
with the conformation model or the Km Discrimination Model. However,
RNA sequencing may not be quantitative enough to make this distinction.
It is possible that with the new capability to make synthetic RNA
oligonucleotides in the laboratory, that more sensitive measurements can
be made to determine the error frequency of poliovirus polymerase at
specific sequences utilizing the elongation assay developed by Bossalis
(Boosalis et al., 1987). This method involves elongating a 5' end-
labeled primer with the separate addition of the four different

78
nucleotide substrates and assaying for elongation by gel
electrophoresis.
Evolution Rates of Poliovirus
It is interesting to note that poliovirus is a very stable virus
when grown in tissue culture, yet it is known to change very rapidly
when it replicates in humans (for details see Chapter 1). The fact that
there are only three serotypes of poliovirus is probably a reflection of
having only a few functionally distinct neutralization sites, rather
than a reflection of phenotypic stability of any single antigenic site.
While these neutralization sites are in general very stable, any change
in the genome at these neutralization sites may result in an outbreak of
poliomyelitis as was seen with the relatively recent (1984) outbreak in
Finland. The changes in the neutralization sites, however, were not
solely responsible for this outbreak. In fact, the virus was
neutralized by antisera that was specific for that serotype. However,
the Finnish population used killed poliovirus vaccine, which overall
results in lower antibody titers than the attenuated poliovirus vaccine.
So it appeared that it was the combination of low antibody titers and
changes in some of the neutralization sites that resulted in the
poliomyelitis outbreak.
Master Sequence Theory
An important question then is why poliovirus is so stable when
grown under certain conditions and so quick to mutate under others. One
theory is that RNA viruses consist not of a single genotype but of a
distribution of related genotypes (Domingo et al., 1978, 1985). There
is a distribution around one or several degenerate master sequences

79
which are efficiently replicated. Under some conditions, the fraction
of master sequences relative to the total population is low and mutants
dominate the population. Mutants of high reproductive power relative to
the master sequence may modify the population drastically. This appears
to be the case when poliovirus is replicating in the human body. In
contrast, when poliovirius is grown in tissue culture, a single master
sequence is maintained. While the wild type is not characterized by a
single sequence, the wild type appears to have an unambiguous consensus
sequence which is probably identical to the master sequence. My data
clearly demonstrated that the poliovirus genome does change while
replicating in tissue culture at least at the eight different sites
measured for a small percentage of the population. However, sequencing
of the viable poliovirus population would have undoubtedly shown the
maintenance of the same consensus sequence that has been propagated in
the laboratory for years. It would thus appear that every nucleotide in
the poliovirus genome is selected for in one way or another. Selection
may be at the level of RNA structure as it relates to RNA packaging and
recognition by the replication machinery, as well as at the protein
level. As the poliovirus genome is initially translated as a large
polyprotein, the structure of this polyprotein may be very important for
cleavage by the viral proteases and any disturbance in this structure
may affect proteolytic processing. Also there is evidence that many of
the viral replicative proteins may only function in cis (Bernstein et
al.,1986) which may limit the perpetuation of mutant RNA sequences and
their corresponding mutant viral proteins.

80
RNA Genomes vs. DNA Genomes
It has already been mentioned that in general RNA genomes are of
limited length relative to DNA genomes. The error threshold for
maintenance of genetic information would be expected to correlate with
the sequence length, hence allowing a higher error frequency for RNA
polymerases. It should also be mentioned that RNA genomes are only
found as host dependent cellular parasites. Much of their replication
strategy therefore involves using host cell machinery, not their own.
RNA genomes also have much shorter replication times which contribute to
their higher evolution rate.
While DNA genomes are copied with a much higher fidelity than RNA
genomes, they have other strategies which allow for their evolution.
Recombination is one such strategy utilized by DNA genomes and to a
lesser extent can be utilized by some RNA genomes, including poliovirus.
One potentially interesting question is the evolution of RNA
viruses compared to DNA viruses. Do RNA viruses evolve independently of
most host cell functions whereas DNA viruses coevolve with the host
cell? Certainly some of the DNA viruses and the retroviruses which
utilize a DNA polymerase can integrate and be maintained and replicated
by the host cells replicative machinery. Perhaps it is therefore to
their advantage to evolve at a slower rate similar to their host cell.
Diversity may be much more advantageous for RNA viruses compared to DNA
viruses. Certainly the generation of defective interfering particles
and the large ratio of particle to infectious virions for RNA viruses
suggests that diversity is of great importance.

81
Future of RNA Viruses
Perhaps the most important lesson to be learned from the growing
accumulation of knowledge regarding the high error frequency of RNA
polymerases is to expect the continual emergence from time to time of
new diseases due to the continual evolution of RNA viruses. While RNA
viruses generally have a defined method of transmission and are
associated with a particular disease, there may be appearances of new
strains with markedly different host range, tissue tropism, disease
patterns, and virulence. This may be particularly true when a virus can
find a new niche, whether it be a new host, a new tissue, or a new
vector. We have witnessed the emergence of such new diseases. Acute
hemorrhagic conjunctivitis is one example of a new disease that emerged
in the 1960's and is caused by a picornavirus. And of course AIDS is
the latest example of such a new disease, which emerged in the 1970's
and is caused by a retrovirus which utilizes both RNA and DNA in its
replication strategy.

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17846-17856.

BIOGRAPHICAL SKETCH
I was born December 2, 1960, in Washington, DC, the fifth and last
child of John and Ruth Ward. I lived in Silver Spring, Maryland, for 15
years where I attended Holiday Park Elementary School, Newport Junior
High School, and one year at Albert Einstein High School. I moved in my
junior year to Minnesota where I attended Fridly Senior High. The cold
quickly drove my family back to the Washington, DC, area where I
attended George Mason University from 1978 to 1983. I foolishly got
married while I was an undergraduate student, which ended in divorce
while I was a graduate student. Luckily, I met another graduate
student, Paul Kroeger, whom I married in 1987. We had one child in
1988, Alan Scott Kroeger, and are expecting our second child at the end
of this year.
89

I certify that I
conforms to acceptable
adequate, in scope and
Doctor of Philosophy.
I certify that I
conforms to acceptable
adequate, in scope and
Doctor of Philosophy.
I certify that I
conforms to acceptable
adequate, in scope and
Doctor of Philosophy.
I certify that I
conforms to acceptable
adequate, in scope and
Doctor of Philosophy.
I certify that I
conforms to acceptable
adequate, in scope and
Doctor of Philosophy.
have read this study and that in my opinion it
standards of scholarly presentation and is fully
quality, as a dissertation for the degree of
nes B. Flanegan,/(Jhair
Professor of Immunology and
Medical Microbiology
have read this study and that in my opinion it
standards of scholarly presentation and is fully
quality, as a dissertation for the degree of
Ernest Hiebert
Professor of Plant Pathology
have read this study and that in my opinion it
standards of scholarly presentation and is fully
quality, as a dissertation for the degree of
Lindsey Hu£t>Fletcher
Professor of Pathology and
Laboratory Medicine
have read this study and that in my opinion it
standards of scholarly presentation and is fully
quality, as a dissertation for the degree of
Richard Moyer
Professor of Immunbligy and
Medical Microbiology
have read this study and that in my opinion it
standards of scholarly presentation and is fully
quality, as a dissertation for the degree of
S
on
Sue Moyer
Professor of Immunology and
Medical Microbiology

This dissertation was submitted to the Graduate Faculty of the
College of Medicine and to the Graduate School and was accepted as
partial fulfillment of the requirements for the degree of Doctor of
Philosophy.
December, 1990
!5Wl
Lsu^av^ ^\.
Dean,
Colleg
e o
f- Medicine
Dean, Graduate School

UNIVERSITY OF FLORIDA
3 1262 08554 4939