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

MISSING IMAGE

Material Information

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

Subjects

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

Notes

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

Record Information

Source Institution:
University of Florida
Rights Management:
All applicable rights reserved by the source institution and holding location.
Resource Identifier:
aleph - 001581459
oclc - 25072896
notis - AHK5373
System ID:
AA00009078:00001


This item is only available as the following downloads:


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
















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.
















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 lonophoresis............................... 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]PO4.............. 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 Tl Digestions .................................... 23
Gel Purification of Oligonucleotides..................... 23
5' End-labeling........................................ 25
RNA Sequencing......................................... 25
Gel Electrophoresis.................................... 26









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 in 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

AM, ATP = 7 pM, CTP = 6 pM, and GTP = 5 AM. 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 Mn12 for Mg+2 was found to

increase the error frequency 2-fold. Changes in the template, the

reaction temperature, and the noncomplementary ribonucleotide substrate









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 103 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.
















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 109 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









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 in 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 E. 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 Affecting 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 Mn 2 o Co2, Ni2, Zn2+, and Be 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 pM) 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 104 (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 Tl. 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 104 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 Q8 has been calculated to be about 104 (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 10-4 (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 10"3 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 Tl 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 104 to 10s 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 10-4 to 10-s 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 Tl Oligonucleotide Mapping

Poliovirus evolution rates have also been measured numerous times

by RNase Tl 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 Tl

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 104.

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 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 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 106 (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 106 was calculated for this

site compared to 1.5 X 10-s 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,











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, 3DP', 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 Homopolymers

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 Tl 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]PO4. 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-3P]UTP, [a-32P]CTP, [a-3P]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









15

NH4HCO2.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 pl) contained 50 mM

HEPES (N-2-hydroxyethyl piperazine-N'-2-ethane sulfonic acid) pH 8.0, 3

mM MgCl2, 10 mM DTT, 2.5 Ag poly(A), 1.25 Mg oligo(U), and 3 pl Fraction

IV(HA) polymerase purified as described (Young et al.,1986). For the

correct ribonucleotide, 10 MCi of [5,6-3H]UTP (36 Ci/mmole) was added,

and for the incorrect ribonucleotide 20 uCi of either [a-32P]ATP, [a-

32P]GTP, or [-32 P]CTP was added. Unlabeled ribonucleotides (Calbiochem

Corporation) were added to make the reaction mixtures equimolar (7.2 PM)

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 ACi of [3H]UTP was used and unlabeled UTP was added

to a final concentration of 74 jiM for the correct ribonucleotide. For

the incorrect ribonucleotide, 20 pCi of [3P]CTP was used and unlabeled

CTP was added to a final concentration of 7.4 pM. Alternatively, 10 pCi

of [3H]UTP was used and 10 pCi of [32P]CTP was used making the

concentrations of correct and incorrect ribonucleotides 7.4 pM and 0.74

pM respectively. At 0.1:1 ratio of correct to incorrect

ribonucleotides, 1 pCi of [3H]UTP was used as the correct ribonucleotide

and 20 pCi of [32P]CTP was used and unlabeled CTP was added to a final

concentration of 7.4 pM.

Mg12 versus Mn+2

Another variation including substituting 0.5 mM MnCl2 for 3 mM

MgC12. 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 pM and the

incorrect ribonucleotide being at 0.74 pM.

Temperature

The error frequency was determined at 30*C, 37C, and at 420C.

This was done at equimolar concentrations and at a 10:1 ratio of correct

to incorrect ribonucleotides as described above.









17

RNA Digestion with P1 Nuclease

Product RNA was synthesized on a poly(A) template as described

above using 10 ACi each of [a-32P]UTP and 10 pCi of either [a-32P]ATP,

[a-32P]GTP, or [a-3P]CTP. The labeled product RNA was phenol:chloroform

extracted, ethanol precipitated, dissolved in 100 pj 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 py 10 mM CH3COONa.3H20, pH 6.0 containing 1.5 units of P1 nuclease

(Bethesda Research Laboratory). The sample was then heat denatured at

100C for 10 minutes, cooled to 37C, and another 1.5 units of P1

nuclease in 15 1l of 10 mM CH3COONa-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 lonophoresis

lonophoretic separation of the P1 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 pg) was copied in the presence of 10 pCi of

[32P]UTP and enough unlabeled UTP to make the final concentrations 0.8

pM, 5 pM, 10 pM, 20 pM, 40 pM, or 80 pM. The other three

ribonucleotides were also present at 500 pM. Standard reaction

conditions were used (3 mM MgC12, 50 mM HEPES pH 8.0) except that the









18

total volume was 150 pm and 30 il 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 -200C.

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.

Oligodeoxyribonucleotides

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 [32P]PO4

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 250C). 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 NaCI, 5 mM KC1, 26 mM NaHC03, pH 7.5). After 15 min,

Actinomycin D was added at a concentration of 5 ug/ml. At 30 min 600

pCi/ml [3P]PO4 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 pg) uniformly labeled with [32P]PO4 was

hybridized with 0.3 pug of DNA oligonucleotide in 30 Ml of 10 mM Tris

HC1, pH 7.5, 500 mM NaCI, and 4 mM MgC12, for 3 h at 50C. The hybrids

were digested with 6 units of RNase Tl for 2 h at 50C. The RNA was

treated with proteinase K by diluting the sample with 120 Al 0.5% SDS

buffer (100 mM NaCI, 10 mM Tris, pH 7.5, 1 mM EDTA, .5% SDS) and adding

75 pg proteinase K for 1 hour, phenol:chloroform extracted two times and

ethanol precipitated. The RNA:DNA duplex was then purified on a 20%












Table 9 -1 T c I- -fNA 014 anh ni Ann Prnta+- A DMA Q nc


Name vRNA bases

BF8 6862 6895


BF9 7269 7296


BF10 5638 5671


BF11 3971 3999


BF25 84 109


BF27 1229-1253


BF30 1429-1450


BF35 2757-2776


DNA sequence 5'-3'/RNA sequence 3'-5'


CAAGTTGTTAATCATTGAGTTAAAAATTGAAGTGAAAAAAAAAAAAAAA
GUUCAACAAUUAGUAACUCAAUUUUUAACUUCAC

CTGATTTTAGCTAGGAATTTGTTATATTAAAAAAAAAAAAAAA
GACUAAAAUCGAUCCUUAAACAAUAUAA

CTTTAGAGTGATTATAGTGATTTCAAGATTGGTTAAAAAAAAAAAAAAA
GAAAUCUCACUAAUAUCACUAAAGUUCUAACCAA

CTAGTTATAATAACTAGTGAGGATATGATAAAAAAAAAAAAAAA
GAUCAAUAUUAUUGAUCACUCCUAUACUA

CTAAGTTACGGGAAGGGAGTATAAAAAAAAAAAAAAAAAAA
GAUUCAAUGCCCUUCCCUCAUAUUUU

CTAGGTAGTGGTAGTACATATTTTGAAAAAAAAAAAAAAA
GAUCCAUCACCAUCAUGUAUAAAAC

CTGGTTGTTGTCAGGAGTGAAAAAAAAAAAAAAAAA
GACCAACAACAGUCCUCACUU

CGTGGTGGAAGCTGGGTTATAAAAAAAAAAAAAAA
GCACCACCUUCGACCCAAUA


Table 2-1 List of









21

polyacrylamide gel, denatured, and the RNA oligonucletide was

thenrepurified on a 20% polyacrylamide, 7M urea gel. The RNA fragment

was then RNase TI 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)3 sequence (Oberste,

1988a). The plasmid pOF2612 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 Tl digestion experiments

as described above for [32P]PO4 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 jig cold















Labeled vRNA


DNA oligo

Hybridize
Tl Digest
20% Acrylamide Native Gel


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
T1


Resistant Oligo

Digestion Products


Fig. 2-1. Schematic diagram of [32P]PO4 labeled vRNA hybridization and
RNase T1 digestion assay to determine error frequency of the polymerase
at specific sites on the genome.


Poly(A)









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 pl of 100 mM CH3COONa.3H20 (pH 4.5), 5

mM MgSO4, and purified on a denaturing 20% polyacrylamide, 7 M urea gel.

The protected RNA fragment was then digested with RNase Tl, 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 TI Digestions

Isolated labeled RNA fragments were ethanol precipitated with 20

pg glycogen (Boehringer Mannheim) and dissolved in 6 pl of 25 mM sodium

citrate (pH 3.5), 7M urea, 1 mM EDTA, 0.035% xylene cyanol and 0.035%

bromphenol blue. RNase Tl (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

Tl was added and incubated for another hour at 50C. Proteinase K

(1 pg) 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 37*C in 150 1l 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












vRNA


*- -


* -


DNA oligo


Hybridize
Tl Digest
20% Acrylamide Native Gel


Poly(A)


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 TI 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

pg glycogen. Fragments eluted from very large gel pieces were extracted

three or four times with absolute ethanol to eliminate any urea.

5' End-labeling

RNA and DNA fragments were routinely end labeled with 10 pCi of

[y-32P]ATP using 10 units of T4 polynucleotide kinase (New England

Biolabs). When RNA pieces were to be sequenced, however, 50 pCi was

used instead for each hybridization and digestion that started with 10

pg poliovirion RNA.

RNA Sequencing

RNase Tl-resistant oligonucleotides were gel purified and ethanol

precipitated with 15 pg of tRNA. The RNA was divided into five aliquots

which were digested with either RNase Tl (Calbiochem Corporation), RNase

U2 (Bethesda Research Laboratories), RNase PhyM (Bethesda Research

Laboratories), RNase B. cereus (Bethesda Research Laboratory) or RNase

CL3 (Pharmacia). RNase TI 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 Tris.HCl, 100 mM H3BO3, 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 Pl nuclease. P1 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.









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 AM, 0.8 pM, 2.5 pM, 5

pM, 10 pM, 20 pM, 40 pM, and/or 80 pM) while the other three nucleotides

were kept constant at 500 pM. 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 pM, UTP = 7 pM, CTP = 6 pM,

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














4
S UTP
0
E
0 3


S[UTP]
2
o 0 .4 uM
.8uM
0
1 luM
a 1 5UM
.............. 0 10uM
80 uM


0 10 20 30 40
Time (min)





1.2-
*ATP
o 1.0

0.8
V [ATP]
| 0*.6-
0] .4uM
0
.8uM
0 0.4- a 5uM

0 10 uM
0.2- 80 uM

0.0 I I
0 10 20 30 40
Time (min)




Figure 3-1. Effect of [UTP] and [ATP] on the initial velocity of the
poliovirus polymerase reaction. Initial velocities (vo) 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 MgC1,, pH 8, 30C) except that the
initial reaction volume was 150 pl and 30 4l aliquots were removed and
counted every ten minutes.
















E 0.8
C-

0.6 -
0.6* [CTP]
0 5 .4 uM
0.4 < .8 uM
a 0 Ua 5 uM
0.2- 0o 10 UM
a S 40 uM
0.0

0 10 20 30
Time (min)








1.0- GTP


E 0.8


S 0.6-
[GTP]
0.4- 0 2.5 uM
S* 5 uM
5 10 uM
0.2* 40 uM

0.0 ,I
0 10 20 30
Time (min)


Figure 3-2. Effect of [CTP] and [GTP] on the initial velocity of the
poliovirus RNA polymerase reaction. Initial velocities (vo) 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 (vo) for GTP were repeated and both values were plotted
on the Lineweaver/Burk plot shown in Figure 3-4.










32





600- Km= 7 uM

500-

400-

> 300 -
'I-
200-

100-

0-o

-100 I I
-1 0 1 2 3
1/[ATP]






300-
Km =10 uM
250 -

200-

> 150

"- 100-

50- 0

0

-50 = I II
-1 0 1 2 3
1/[UTP]







Fig. 3-3. Determination of Km for ATP aned UTP using Lineweaver-Burke
plots. Double reciprocal plot of v'1 vs. [NTP]f1 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.


















300-


200-


100-


0 ---


-100 -t -- -- -- -- -- --
-1 0 1 2 3
1/[CTP]






200 Km= 5 uM


150-


100- [


50- f
0



-50 1 I I
-0.5 0.0 0.5 1.0
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 pM to 74 pM and the CTP

concentration from 0.74 pM to 7.4 pM. 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












Table 3-1. Error Frequency of the Poliovirus RNA Polymerase


Reaction 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 103

Poly(C) [32P]ATP [3H]GTP 2.9 1.2 X 10-3
3 mM MgC12 [32P]CTP [3H]GTP 4.8 2.0 X 10-3
pH 8 [32PJUTP [3H]GTP 2.5 0.2 X 10-3

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 pl. In the poly(C)
and poly(I) reactions, the final volume was 30 4l and the total
nucleotide concentrations were 25.6 AM and 16.6 AM 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.
















Table 3-2. Effect of Nucleotide Concentration on the Error Frequency of
the Polymerase


[UTP] [CTP] [UTP]/[CTP] Error Frequencya


0.7 pM 7.4 pM 0.1 3.2 0.8 X 10-3

7.4 pM 7.4 pM 1 2.9 0.4 X 10-3


74.0 AM 7.4 pM 10 2.0 0.8 X 10-4

7.4 pM 0.7 AM 10 4.4 1.6 X 104


a All reactions were on poly(A), 3 mM MgC2,, 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 MnCl2

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 AM UTP,

0.7 AM 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).
















Table 3-3. Effect of MnCl2 on Poliovirus Polymerase Error Frequency


Nucleotide Ratio'


Increase in


Compl./Noncompl. Divalent Cationb Error Frequency Error Frequency
10:1 3 mM MgCl2 0.8 0.1 X 104 1

10:1 0.5 mM MnC12 1.4 0.3 X 10-4 2


1:1 3 mM MgCl2 7.2 2.0 X 10-4 9

1:1 0.5 mM MnCl2 12.6 2.4 X 10-4 16


a At 10:1 ratio of complementary to noncomplementary substrates, 7.4 UM
UTP and 0.7 uM CTP were used respectively. At 1:1 ratio of
complementary to noncomplementary substrates, 7.4 pM UTP and 7.4 pM
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 ratio
Compl./Noncompl. 300C 370C 42C


1:1 7.2 2.0 X 10-4 7.4 1.9 X 104 6.8 2.2 X 10-4
10:1 8.3 0.8 X 10"5 1.3 0.3 X 10-4 9.3 5.8 X 105


a At 1:1 ratio off complementary to noncomplementary nucleotide, 7.4 AM
UTP and CTP were used. At 10:1 ratio of complementary to
noncomplementary nucleotide 7.4 AM UTP and 0.7 pM CTP were used. All
reactions were on poly(A), 3 mM MgCl2, pH 8.









40

The error frequency was determined at 30C, 37C, 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 pM UTP,

0.7 jM 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 Pl nuclease. P1 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






















m
9*0



* 0.


0
9
e9
la

a0.
00.
o.0


m
*0.

*0.





* M.
0 .


* .

* .
D
*0.


D I

0


in 'c


*0


4
CL
&

3




S :o







3



*4






4L


i3
0


n

*0.


0
U)
*C


-4


o
-4







o
c t .
4 o


co
( 04 0






0 0





mo o,



oUo
( ca
0) 0
4- (4-4 0) 0
0 0 a (
) *0 U H

0 0
0C 0 0
H 4H 0 "0
In 0 0 3 U
4o e1

0 .a a) *1

4 C 4) -4


0 0)



E 0 0



4 0) ")

-4 -H





-4 -4 ~ 4-1


0



in in
















pG


* pA


0


ori


Figure 3-6. High-voltage ionophoretic separation of P1 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 mM 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 P1 Nuclease Digestion


Reaction Noncomplementary Complementary
Conditions Substrate Substrate Error Frequency


Poly(A) [32P]ATP [32P]UTP 2.9 2.8 X 10-3
3 mM MgC12 [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 MgC12 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 P1 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 103. 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 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









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 Tl. The protected RNA

oligonucleotide that was complementary to the DNA was isolated by gel

purification and then digested with RNase Tl. 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 Tl and U2 or RNase Tl 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 1B and IC). 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 [3P]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 Tl, 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]PO4-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













1A 1B IC 1D 2A 2B 2C 2D 3C


Figure 4-1. Isolation of RNA oligonucleotide protected from BF8 after
RNAse TI 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 TI and U2 (lanes A) or RNase TI (lanes D) or was first
hybridized with synthetic DNA oligonucleotide BF8 and then digested with
RNases TI and U2 (lanes B) or RNase TI (lanes C). Poliovirion RNA
labeled in vivo with [3P]P04, hybridized with BF8, and digested with
RNase TI is shown in lane 3C.


40^r


--^WWMZO^















1 2


a


4m


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 Tl 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 Tl digestions were done at
550C.



















+


- O




do
- 1




u- ao




- a






OP


dip
a aS


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.


D
<








1 2 3 4


a-



I
e


RNA-DNA duplex-
36-



26-


U

m


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]PO4 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 Tl resistant band relative to the

radioactivity in the two oligonucleotides that were the RNase Tl

digestion products.

Polymerase Error Frequency Determined by Using r32P]PO.-Labeled
Poliovirion RNA

The error frequency has been measured at two sites in the

poliovirus genome using the procedure described above and [32P]PO4-

labeled RNA. The two sites were at nucleotide 6883 in the 3DP~ coding

sequence and at nucleotide 5648 in the 3Cpm 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 Tl 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













1 2 3


Poly(A) { "
!=j


34 -


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]PO4 (lane 1) SP6
polymerase transcripts of pOF1205 DNA were labeled with either [32p]UMP
(lane 2) or [32P]AMP (lane 3).


.A?*
,V=,
.: -
.. '*if















1 2











































Figure 4-6. Final RNase TI 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]PO4 was
run after RNase Tl 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 Tl. 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














Table 4-1. Poliovirus Error Frequency at Specific Sites


Labeling Methodb
Protecting DNA oligonucleotidea [32P]PO4 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
3.2 X 10-3

BF11: 3971-3999 2B conserved 3.5 X 10-3

BF25: 84-109 5'NC conserved 3.8 X 10-3

BF27: 1229-1253 VP2 conserved 5.0 X 10-3

BF30: 1429-1450 VP2 variable 4.6 X 10-3

BF35: 2757-2776 VPI variable 3.2 X 10-3


aNumbers of protecting DNA oligonucleotide represent the nucleotides
protected in the poliovirus genome. 3D, 3C, 2B, VP2, and VPl 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 TI and are an average of at least 3 experiments.
















VPg-F


Polyprotein


VP4
VP2 VP3 VP1 Apl2B
T-C CVP CD
M CT O D0
-- C I *P 0)
C cv V C


3 BVPg
2C 3A 3Cpro

It
CD


Figure 4-7. Schematic diagram of the poliovirus genome with the various
sites examined by RNase TI digestion indicated. C represents constant
regions on the genome and V represents variable regions on the genome.


,p.t'-.


Poly(A)


3Dp0 -


I


r


I













4 5 6


so&


4- BF10 Hyb


BF27 Hyb


4- B F10


BF27 -.


Figure 4-8. Isolation of RNA oligonucleotides protected from BF27 and
BF10 after RNase TI digestion. DNA oligonucletides BF27 and BFO1 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 Tl.
BF27 hybridization and digestion products were run in lanes 2 and 3.
BF10 hybridization and digestion products were run in lanes 4 and 5.


2 3















1 2 3 4


Poly(A) {


FRNA --


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.

















1 2 3 4


Figure 4-10. Final RNase Tl 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 Tl 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

Tl resistant oligonucleotides were redigested with RNase Tl. The amount

of RNase Tl resistant fragment that could be redigested with Tl 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











*AAUAUAACAAAUUCCUAGCUAAAAUCAG 28mer
I T1 Digest
*AAUAUAACAAAUUCCUAG CUAAAAUCAG
18mer 10mer


Figure 4-11. Final RNase TI 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 10mer
(unlabeled). Lanes 2 5 are all separate reactions.














*AUCAUAUCCUCACUAGUUAUUAUAACUAG 29mer


T1 Digest


*AUCAUAUCCUCACUAG
16mer


UUAUUAUAACUAG
13mer


Figure 4-12. Final RNase TI 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.















*UUUUAUACUCCCUUCCCGUAACUUAG 26mer


T1 Digest


*UUUUAUACUCCCUUCCCG
18mer


UAACUUAG
8mer


Nl


Figure 4-13. Final RNase TI 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.













*CAAAAUAUG UACUACCACUACCUAG 25mer

I T1 Digest


*CAAAAUAUG
9mer


1 2


UACUACCACUACCUAG
16mer


3 4





25


Figure 4-14. Final RNase TI 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 separate
reactions.
















*UUCACUCCUGACAACAACCAG 21 mer


T1 Digest


*UUCACUCCUG
10mer



1


ACAACAACCAG
11mer


2 3


Figure 4-15. Final RNase TI digestion products of 5'-end labeled
oligonucleotide protected by BF30. Lane 1 is undigested marker. BF30
protects a 21mer and digests to a 10mer (labeled) and a llmer
(unlabeled). Lanes 2 and 3 are separate reactions.















*AUAACCCAGCUUCCACCACG 20mer


T1 Digest


*AUAACCCAG
9mer


1


CUUCCACCACG
11mer


20








9


Figure 4-16. Final RNase TI digestion products of 5'-end labeled
oligonucleotide protected by BF35. Lane 3 is undigested marker. BF35
protects a 20mer and digests to a 9mer (labeled) and a llmer
(unlabeled). Lanes 1 and 2 are separate reactions.
















+ +
0 : < 0 a <









U


U(C)
U



A
A
C -
U /
C .
A







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















+ +


















t


A
A
U
A
U
U
A
1 U
U
G
A
U

C
A

C

U


+
0 0



ar


+
<0C3<


C



U


*




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









72

nucleotides in length. Apparently 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 Tl 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 TI. 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 Tl digestion products. Unexpected high bands appeared on the

final RNase TI digestion gels which could not be accounted for. They

were resistant to DNase and additional RNase Tl 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 Tl. Therefore, it

was important to determine how efficient this digestion was. For this

reason RNase Tl resistant oligonucleotides were sequenced when possible.

A large limiting factor on sequencing, however, was the small amount of

radioactivity in these RNase Tl resistant bands. When sequencing could

not be done, the RNase Tl resistant bands were redigested with RNase Tl

and quantitated to determine the percent that could be redigested. This

percentage varied from 10 50%. Sequencing showed approximately 25% of

the RNase Tl 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 VPl 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 3DP' 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 3Cpm 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 p

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 103, 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









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 interaction with the nucleotide substrates.

However, it was also clear in sequencing the TI 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.
















REFERENCES


Abbotts, J., and Loeb, L.A. (1984). On the fidelity of DNA replication:
Lack of primer position effect on the fidelity of mammalian DNA
polymerases. J. Biol. Chem. 259, 6712-6714.

Almond, J.W., Westrop, G.D., Evans, D.M., Dunn, G., Minor, P.D.,
Magrath, D., and Schild, G.C. (1987a). Studies on the attenuation of the
Sabin type 3 oral polio vaccine. J. Virol. Methods. 17, 183-189.

Almond, J.W., Westrop, G.D., Wareham, K.A., Skinner, M.A., Evans, D.M.,
Magrath, D., Schild, G.C., and Minor, P.D. (1987b). Studies on the
genetic basis of attenuation of the Sabin type 3 poliovirus vaccine.
Biochem. Soc. Symp. 53, 85-90.

Anderson-Sillman, K., Bartal, S., and Tershak, D.R. (1984).
Guanidine-resistant poliovirus mutants produce modified 37-kilodalton
proteins. J. Virol. 50, 922-9280.

Baltera, R.F., Jr., and Tershak, D.R. (1989). Guanidine-resistant
mutants of poliovirus have distinct mutations in peptide 2C. J. Virol.
63, 4441-4444.

Barrel, B.G. (1971). Fractionation and sequence analysis of radioactive
nucleotides. In Procedures in nucleic acid research, G. Contoni and D.
Davies, eds. (New York: Harper & Row Publishers, Inc.), pp. 751-828.

Batschelet, E., Domingo, E., and Weissmann, C. (1976). The proportion of
revertant and mutant phage in a growing population, as a function of
mutation and growth rate. Gene 1, 27-32.

Battula, N., and Loeb, L.A. (1974). The infidelity of avian
myeloblastosis virus deoxyribonucleic acid polymerase in polynucleotide
replication. J. Biol. Chem. 249, 4086-4093.

Beckman, Robert, A., Mildvan, Albert, S., and Loeb, Lawrence, A. (1985).
On the fidelity of DNA replication: Manganese mutagenesis in vitro.
Biochemistry 24, 5810-5817.

Bernstein, H.D., Sarnow, P., and Baltimore, D. (1986). Genetic
complementation among poliovirus mutants derived infectious cDNA clone.
J. Virol. 60, 1040-109.











Boosalis, M.S., Petruska, J., and Goodman, M.F. (1987). DNA polymerase
insertion fidelity: Gel assay for site-specific kinetics. J. Biol.
Chem. 262, 14689-14696.

Brand, C., and Palese, P. (1980). Sequential passage of influenza virus
in embryonated eggs or tissue culture: Emergence of mutants. Virology
107, 424-433.

Britlen, R.J. (1986). Rates of DNA sequence evolution differ between
taxonomic groups. Science 231, 1393-1398.

Coffin, J.M., Tsichlis, P.N., Barker, C.S., Voynow, S., and Robinson,
H.L. (1980). Variation in avian retrovirus genomes. Ann. NY. Acad. Sci.
354, 410-425.

Dasgupta, A., Zabel, P., and Baltimore, D. (1980). Dependence of the
activity of the poliovirus replicase on a host cell protein. Cell 19,
423-429.

de la Torre, J.C., Wimmer, E., and Holland, J.J. (1990). Very high
frequency of reversion to guanidine resistance in clonal pools of
guanidine-dependent type 1 poliovirus. J. Virol. 64, 664-671.

Domingo, E., Flavell, R.A., and Weissman, L. (1976). In vitro site
directed mutagenesis: Generation and properties of an infectious
extracistronic mutant of bacteriophate QB. Gene 1, 3-25.

Domingo, E., Martinez-Salas, E., Sobrino, F., de la Torre, J.C.,
Portela, A., Ortin, J., Lopez-Galindez, C., Perez-Brena, P., Sillanueva,
N., Najera, R., Van de Pol, S., Steinhauer, D., DePolo, N., and Holland,
J. (1985). The quasispecies (extremely heterogeneous) nature of viral
RNA genome populations: Biological relevance--a review. Gene 40, 1-8.

Domingo, E., Sabo, D., Tadastsugu, T., and Weissman, C. (1978).
Nucleotide sequence heterogeneity of an RNA phage population. Cell 13,
735-744.

Drake, J.W. (1990). Spontaneous mutation. Nature 221, 1128-1132.

Emini, E.A., Wimmer, E., Jameson, B.A., Bonin, J., and Diamond, D.
(1984a). Neutralization antigenic sites of poliovirus and peptide
induction of neutralizing antibodies. Ann. Sclavo. Collana. Monogr. 1,
139-146.

Emini, E.A., Leibowitz, J., Diamond, D.C., Bonin, J., and Wimmer, E.
(1984b). Recombinants of Mahoney and Sabin strain poliovirus type 1:
Analysis of in vitro phenotypic markers and evidence that resistance to
guanidine maps in the nonstructural proteins. Virology 137, 74-85.

Flanegan, J.B., and Baltimore, D. (1977). Poliovirus-specific
primer-dependent RNA polymerase able to copy poly(A). Proc. Natl. Acad.
Sci. U. S. A. 74, 3677-3680.











Flanegan, J.B., and Van Dyke, T.A. (1979). Isolation of a soluble and
template-dependent poliovirus RNA polymerase that copies virion RNA in
vitro. J. Virol. 32, 155-161.

Flanegan, J.B., Young, D.C., Tobin, G.J., Stokes, M.A., Murphy, C.D.,
and Oberste, S.M. (1987). Mechanism of RNA replication by the
polioivirus RNA polymerase, HeLa cell host factor, and VPg. In Positive
strand RNA viruses, UCLA symposium on molecular and cellular biology,
new series, volume 54, R. Rueckert, and M. Brinton, eds. (New York: Alan
R. Liss), pp. 273-284.

Fowler, R.G., Degnen, G.E., and Cox, E.C. (1974). Mutational specificity
of a conditional Escherichia coli mutator, mut D5. Mol. Gen. Genet. 133,
179-191.

Gojobori, T., and Yokoyama, S. (1985). Rates of evolution of the
retroviral oncogene of Maloney murine sarcoma virus and its cellular
homologues. Proc. Natl. Acad. Sci. U. S. A. 82, 4192-4206.

Goodman, M.F., Hopkins, R., and Gore, W.C. (1977). 2-aminopurine-induced
mutagenesis in T4 bacteriophage: A model relating mutation frequency to
2-aminopurine incorporation in DNA. Proc. Natl. Acad. Sci. U. S. A. 74,
4806-4810.

Gopinathan, K.P., Weymouth, L.A., Kunkel, T.A., and Loeb, L.A. (1979).
Mutagenesis in vitro by DNA polymerase from an RNA tumour virus. Nature
278, 857-859.

Hanecak, R., Semler, B.L., Anderson, C.W., and Wimmer, E. (1982).
Proteolytic processing of poliovirus polypeptides: Antibodies to
polypeptide P3-7c inhibit cleavage at glutamine-glycine pairs. Proc.
Natl. Acad. Sci. U. S. A. 79, 3973-3977.

Hogle, J.M., Chow, M., and Filman, D.J. (1985). Three-dimensional
structure of poliovirus at 2.9 A resolution. Science 229, 1358-1365.

Hogle, J.M., Chow, M., and Filman, D.J. (1987). The structure of
poliovirus. Sci. Am. 256, 42-49.

Hogle, J.M., and Filman, D.J. (1989). The antigenic structure of
poliovirus. Philos. Trans. R. Soc. Lond. [Biol] 323, 467-478.

Holland, J.J., Kohne, D., and Doyle, M.V. (1973). Analysis of virus
replication in ageing human fibroblast cultures. Nature 245, 316-318.

Holland, J.J., Spindler, K., Horodyski, F., Grabau, E., Nichol, S., and
Van de Pol, S. (1982). Rapid evolution of RNA genomes. Science 215,
1577-1585.











Hopfield, J.J. (1980). The energy relay: A proofreading scheme based on
dynamic cooperativity and lacking all characteristic symptoms of kinetic
proofreading in DNA replication and protein synthesis. Proc. Natl. Acad.
Sci. U. S. A. 77, 5248-5252.

Hughes, P.J., Evans, D.M., Minor, P.D., Schild, G.C., Almond, J.W., and
Stanway, G. (1986). The nucleotide sequence of a type 3 poliovirus
isolated recent outbreak of poliomyelitis in Finland. J. Gen. Virol. 67,
2093-2102.

Kew, O.M., Nottay, B.K., Hatch, M.H., Nakaro, J.H., and Obijeski, J.F.
(1981). Multiple genetic changes can occur in the oral poliovaccines
upon replication in humans. J. Gen. Virol. 56, 337-347.

Kuchta, Robert D., Benkovic, P., and Benkovic, S.J. (1988). Kinetic
mechanism whereby DNA polymerase I (Klenow) replicates DNA with high
fidelity. Biochemistry 27, 6716-6725.

Kuchta, Robert D., Mizrahi, V., Benkovic, P.A., Johnson, K.A., and
Benkovic, S.J. (1987). Kinetic mechanism of DNA polymerase I (Klenow).
Biochemistry 26, 8410-8417.

Kuge, S., Kawamura, N., and Nomoto, A. (1989). Strong inclination toward
transition mutation in nucleotide substitutions by poliovirus replicase.
J. Mol. Biol. 207, 175-182.

Kunkel, T.A., Eckstein, F., Mildvan, A.S., Koplitz, R.M., and Loeb, L.A.
(1981). Deoxynucleoside[l-thio]triphosphates prevent proofreading during
in vitro DNA synthesis. Proc. Natl. Acad. Sci. U. S. A. 78, 6734-6738.

Kunkel, T.A., and Loeb, L.A. (1981). Fidelity of mammalian DNA
polymerases. Science 213, 765-767.

Li, W.H., Wu, C.I., and Luo, C.C. (1985). A new method for estimating
synonymous and nonsynonomous rates of nucleotide substitution
considering the relative likelihood of nucleotide and codon changes.
Mol. Biol. Evol. 2, 150-174.

Loeb, L.A., and Kunkel, T.A. (1982). Fidelity of DNA synthesis. Ann.
Rev. Biochem. 52, 429-457.

Maniatis, T., Fritsch, E., and Sambrook, J. (1982). Molecular cloning: A
laboratory manual. (Cold Spring Harbor, NY: Cold Spring Harbor
Laboratory).

Merchant-Stokes, M.A. (1985). Poliovirus RNA polymerase: In vitro
enzymatic activities, fidelity of replication and characterization of a
temperature-sensitive mutant, Ph.D. Dissertation, University of Florida.











Minor, P.D., John, A., Ferguson, M., and Icenogle, J.P. (1986).
Antigenic and molecular evolution of the vaccine strain of poliovirus
during the period of excretion by a primary vaccinee. J. Gen. Virol. 67,
693-706.

Minor, P.D., Schild, G.C., Bootman, J., Evans, D.M., Ferguson, M.,
Reeve, P., Spitz, M., Stanway, G., Cann, A.J., Hauptmann, R., Clarke,
L.D., Mountfourd, R.C., and Almond, J.W. (1983). Location and primary
structure of a major antigenic site for poliovirus neutralization.
Nature 301, 674-679.

Murray, M.G., Bradley, J., Yang, X.F., Wimmer, E., Moss, E.G., and
Racaniello, V.R. (1988). Poliovirus host range is determined by a short
amino acid sequence in neutralization antigenic site 1. Science 241,
213-215.

Nottay, B.K., Kew, O.M., Hatch, M.H., Heyward, J.T., and Obijeski, J.F.
(1981). Molecular variation of type 1 vaccine-related and wild
poliovirus during replication in humans. Virology 108, 405-423.

Oberste, M.S. (1988). RNA binding and replication by the poliovirus RNA
polymerase, Ph.D. Dissertation, University of Florida.

Oberste, M.S., and Flanegan, J.B. (1988). Measurement of poliovirus RNA
polymerase binding to poliovirion and nonviral RNAs using a
filter-binding assay. Nucleic Acids. Res. 16, 10339-10352.

Page, G.S., Mosser, A.G., Hogle, J.M., Filman, D.J., Rueckert, R.R., and
Chow, M. (1988). Three-dimensional structure of poliovirus serotype 1
neutralizing determinants. J. Virol. 62, 1781-1794.

Parvin, J.D., Moscona, A., Pan, W.T., Leider, J.M., and Palese, P.
(1986). Measurement of the mutation rates of animal viruses: Influenza A
virus and poliovirus type 1. J. Virol. 59, 377-383.

Pincus, S.E., Diamond, D.C., Emini, E.A., and Wimmer, E. (1986).
Guanidine-selected mutants of poliovirus: Mapping of point mutations to
polypeptide 2C. J. Virol. 57, 638-646.

Pincus, S.E., and Wimmer, E. (1986). Production of guanidine-resistant
and -dependent poliovirus mutants from cloned cDNA: Mutations in
polypeptide 2C are directly responsible for altered guanidine
sensitivity. J. Virol. 60, 793-796.

Preston, B.D., Poiesz, B.J., and Loeb, L.A. (1988). Fidelity of HIV-1
reverse transcriptase. Science 242, 1168-1171.

Richetti, M., and Buc, H. (1990). Reverse transcriptases and genomic
variability: The accuracy of DNA replication is enzyme specific and
sequence dependent. EMBO. J. 9, 1583-1593.











Roberts, J.D., Bebenek, K., and Kunkel, T.A. (1988). The accuracy of
reverse transcriptase from HIV-1. Science 242, 1171-1173.

Rose, J. (1975). Heterogeneous 5'-terminal structures occur on vesicular
stomatitis virus mRNAs. J. Biol. Chem. 250, 8098-8104.

Seal, G., Shearman, C.W., and Loeb, L.A. (1979). On the fidelity of DNA
replication: Studies with human placenta DNA polymerases. J. Biol. Chem.
254, 5229-5237.

Sedivy, J.M., Capone, J.P., RajBhandary, U.L., and Sharp, P.A. (1987).
An inducible mammalian amber suppressor: Propagation of a poliovirus
mutant. Cell 50, 379-389.

Sirover, M.A., and Loeb, L.A. (1976). Infidelity of DNA synthesis in
vitro: Screening for potential metal mutagens or carcinogens. Science
194, 1434-1436.

Sirover, M.A., and Loeb, L.A. (1977). On the fidelity of DNA
replication: Effect of metal activators during synthesis with avian
myeloblastosis virus DNA polymerase. J. Biol. Chem. 252, 3605-3610.

Spindler, K.R., Horodyski, F.M., and Holland, J.J. (1982). High
multiplicities of infection favor rapid and random evolution of
vesicular stomatitius virus. Virology 119, 96-108.

Stanway, G., Hughes, P.J., Mountford, R.C., Reeve, P., Minor, P.D.,
Schild, G.C., and Almond, J.W. (1984). Comparison of the complete
nucleotide sequences of the genomes of the neurovirulent poliovirus
P3/Leon/37 and its attenuated Sabin vaccine derivative P3/Leon/12ab.
Proc. Natl. Acad. Sci. U. S. A. 81, 1539-1543.

Steinhauer, D.A., and Holland, J.J. (1986). Direct method for
quantitation of extreme polymerase error frequencies at selected single
base sites in viral RNA. J. Virol. 57, 219-228.

Takeuchi, Y., Nagumo, T., and Hoshino, H. (1988). Low fidelity of
cell-free DNA synthesis by reverse transcriptase of human
immunodeficiency virus. J. Virol. 62, 3900-3902.

Toyoda, H., Kohara, M., Kataoka, Y., Suganuma, T., Omata, T., Imura, N.,
and Nomoto, A. (1984). Complete nucleotide sequences of all three
poliovirus serotype genomes. Implications for genetic relationship,
gene function and antigenic determinants. J. Mol. Biol. 174, 561-585.

Toyoda, H., Nicklin, M.J., Murray, M.G., Anderson, C.W., Dunn, J.J.,
Studier, F.W., and Wimmer, E. (1986). A second virus-encoded proteinase
involved in proteolytic processing of poliovirus polyprotein. Cell 45,
761-770.











Tuschall, D.M., Hiebert, E., and Flanegan, J.B. (1982). Poliovirus
RNA-dependent RNA polymerase synthesizes full- length copies of
poliovirion RNA, cellular mRNA, and several plant virus RNAs in vitro.
J. Virol. 44, 209-216.

Van Dyke, T.A., and Flanegan, J.B. (1980). Identification of poliovirus
polypeptide P63 as a soluble RNA-dependent RNA polymerase. J. Virol. 35,
732-740.

Van Dyke, T.A., Rickles, R.J., and Flanegan, J.B. (1982). Genome-length
copies of poliovirion RNA are synthesized in vitro by the poliovirus
RNA-dependent RNA polymerase. J. Biol. Chem. 257, 4610-4617.

Villa-Komaroff, L., McDowell, M., Baltimore, D., and Lodish, H.F.
(1974). Translation of reovirus mRNA, poliovirus RNA and bacteriophage
QP RNA in cell-free extracts of mammalian cells. Methods. Enzymol. 30,
709-723.

Watanabe, S.M., and Goodman, M.F. (1982). Kinetic measurement of
2-aminopurine cytosine and 2-aminopurine thymine base pairs as a test of
DNA polymerase fidelity mechanisms. Proc. Natl. Acad. Sci. U. S. A. 79,
6429-6433.

Westrop, G.D., Wareham, K.A., Evans, D.M., Dunn, G., Minor, P.D.,
Magrath, D.I., Taffs, F., Marsden, S., Skinner, M.A., Schild, G.C.
(1989). Genetic basis of attenuation of the Sabin type 3 oral poliovirus
vaccine. J. Virol. 63, 1338-1344.

Yin, F.H., and Lomax, N.B. (1983). Host range mutants of human
rhinovirus in which nonstructural proteins are altered. J. Virol. 48,
410-418.

Young, D.C., Dunn, B.M., Tobin, G.J., and Flanegan, J.B. (1986).
Anti-VPg antibody precipitation of product RNA synthesized in vitro by
the poliovirus polymerase and host factor is mediated by VPg on the
poliovirion RNA template. J. Virol. 58, 715-723.

Young, D.C., Tuschall, D.M., and Flanegan, J.B. (1985). Poliovirus
RNA-dependent RNA polymerase and host cell protein synthesize product
RNA twice the size of poliovirion RNA in vitro. J. Virol. 54, 256-264.

Ypma-Wong, M.F., Filman, D.J., Hogle, J.M., and Semler, B.L. (1988).
Structural domains of the poliovirus polyprotein are major determinants
for proteolytic cleavage at Gln-Gly pairs. J. Biol. Chem. 263,
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.
















I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.


es B. Flanegan, air
Professor of Immunology and
Medical Microbiology

I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.


Ernest Hiebert
Professor of Plant Pathology

I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.


Lindsey Hut -Fletcher
Professor of Pathology and
Laboratory Medicine

I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.


Richard Moyer
Professor of Immun 1 gy and
Medical Microbio y

I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is fully
adequate, in scope and quality, as a dissertation for the degree of
Doctor of Philosophy.


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


Dean, College o Medicine

^>) ^.--Cy^ ^Y-^^^Z6! ~~


Dean, Graduate School









































UNIVERSITY OF FLORIDA
3 1111111111111111111111111111111262 08554 493911111111111111 I I
3 1262 08554 4939




Full Text
xml version 1.0 encoding UTF-8
REPORT xmlns http:www.fcla.edudlsmddaitss xmlns:xsi http:www.w3.org2001XMLSchema-instance xsi:schemaLocation http:www.fcla.edudlsmddaitssdaitssReport.xsd
INGEST IEID EOV9KERVW_75GQ01 INGEST_TIME 2012-02-20T21:29:41Z PACKAGE AA00009078_00001
AGREEMENT_INFO ACCOUNT UF PROJECT UFDC
FILES