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This is to cortify that the

thesis entitled

Regulation and Subcellular Localization of
Human Thymidine Kinase Protein

presented by

Marc A. Carozza

has been accepted towards fulfillment
of the requirements for

Master of Science degeein Microbiology

[Lflizlg (,JWZ

Major professor

 

Date Sept. 28 1993

03639

MSU i: an Affirmative Action/Equal Opportunity Institution

  

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Firm

. will?! State .
University I

 

PLACE IN RETURN BOX to remove this checkout from your record.
TO AVOID FINES return on or before date due.

 

MSU Is An Affirmative Action/Equal Opportunity Institution
I: :\cIrc'\datedue. pm3-p.1

 

 

 

 

REGULATION AND SUBCELLULAR LOCALIZATION OF
HUMAN THYMIDINE KINASE PROTEIN

By

Marc A. Carozza

A DISSERTATION

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

MASTER OF SCIENCE

Department of Microbiology/Cell and Molecular Biology Program

1993

Susan E. Conrad, Major Professor

 

 

ABSTRACT

REGULATION AND SUBCELLULAR LOCALIZATION OF
HUMAN THYMIDINE KINASE

By

Marc A. Carozza

The mechanism by which the human thymidine kinase (TK) enzyme is
regulated at the post—transcriptional level was investigated in serum stimulated, Rat
3 cells stably transfected with a SV40/human TK cDNA construct. The In Vivo
subcellular localization of the TK protein was also determined. The levels of TK
mRNA, protein, and enzyme activity were measured, and while the expression of wild
type TK mRNA varied less than 2-3 fold between early G1 and S phase, there was
a 20-50 fold increase in TK enzyme activity and protein levels during the same
period. Pulse labeling experiments revealed that the rate of translation of TK
mRNA was constant during this period, while pulse-chase experiments indicated the
half life of the TK protein increased 10 fold between G1 and S. We concluded from
these results that the cell cycle regulated increase in TK protein at the Gl/S
interface is due to a change in its stability. It was also revealed through 32F labeling
experiments that TK is phosphorylated. Utilizing indirect immunofluorescence and
confocal microscopy we demonstrated that the human TK polypeptide is localized

exclusively to the cytoplasm in quiescent serum stimulated cells. In addition, this

localization did not change as the cells progressed from G1 into S phase.

 

 

 

 

To my family and my future wife

iii

 

 

ACKNOWLEDGEMENTS

I would like to express my deepest appreciation to Dr. Susan Conrad for her
time, patience, and encouragement during my time in her laboratory. I would also
like to thank the members of my guidance committee; Dr. Ron Patterson, Dr
Michele Fluck, Dr. Richard Schwartz, and Dr. Bill Smith for their helpful
suggestions. Thank you to all the people that I shared the laboratory with during the
years for making the time a little more enjoyable.

I would like to especially thank all the members of the coffee group; Perry,
Dave, Paul, and Rich for all the lively and interesting conversations we had, for the
friendship we shared, but mostly for all the laughs. Also my warmest thanks to
Mark, Larry, Sue and Maggie for their valued friendship and help through the years.

I would like to thank my mother and father for the love, guidance, and
encouragement they have given me throughout my life, I could never have made it
to where I am today without them. Lastly I would like thank my fiance Pam, for the

love and happiness she has brought into my life.

iv

 

 

 

 

TABLE OF CONTENTS

LIST OF FIGURES ........................................................................................................... vi

CHAPTER 1:

Literature Review

The Cell Cycle ........................................................................................... 1
Cyclins and Cyclin Dependent Kinases ................................................ 4
S Phase Regulated Genes ....................................................................... 9
Thymidine Kinase and its Regulation ................................................... 12
Transcriptional Regulation of Thymidine Kinase ............................... 16
Post—Transcriptional Regulation of Thymidine Kinase ...................... 18
Mechanisms of Protein Degradation ..................................................... 22
References .................................................................................................. 25
CHAPTER 2:
Regulation of Thymidine Kinase Protein Stability in Serum Stimulated
Cells .............................................................................................................................. 34
Abstract ....................................................................................................... 35
Introduction ................................................................................................ 36
Materials and Methods ............................................................................ 39
Results ......................................................................................................... 46
Discussion ................................................................................................... 69
References .................................................................................................. 73
CHAPTER 3:
Subcellular Localization of Human Thymidine Kinase in Serum
Stimulated Cells ......................................................................................................... 77
Abstract ....................................................................................................... 78
Introduction ................................................................................................ 79
Materials and Methods ............................................................................ 82
Results ......................................................................................................... 84
Discussion ................................................................................................... 91
References .................................................................................................. 92

 

 

 

 

LIST OF FIGURES

CHAPTER 1:
Figure:
1 Mammalian cell cycle with 16 hr. doubling time .......................... 3
2 Expression of mammalian cyclins and cdks during the
cell cycle .................................................................................................. 8
3 Pathways to obtain purine nucleotides and thymidylate ............. 13
4 Amino acid sequence comparison between TK from viral and
vertebrate sources ................................................................................ 15
CHAPTER 2:
Figure:
1 Mapping the 5’ ends of TK mRNA in transfected
Rat 3 cells ............................................................................................... 49
2 Patterns of TK mRNA, protein, and enzyme activity during serum
stimulation of cells containing pESVTK ......................................... 52
3 Cell cycle analysis of serum stimulated cells containing
ESVTK ................................................................................................... 54
4 Direct comparison of the translation efficiency of TK
mRNA during G1 and S phase .......................................................... 57
5 Determination of TK protein half-life during G1 and S-GZ ...... 59
6 Phosphorylation of TK protein during G1 and S phase .............. 62
7 Regulation of the truncated TK protein encoded by
SV—huASmaTK ....................................................................................... 66
8 3SS—methionine and 32Pi labeling of the truncated protein
encoded by SV-huASmaTK ................................................................. 68
CHAPTER 3:
Figure:
1 Effect of cell cycle progression on subcellular location of TR .. 88
2 Confocal microscopy ........................................................................... 90

vi

 

 

 

 

MR1

LITERATURE REVIEW

The Cell gycle

Mammalian cells exist in two varying states of reproductive activity. In the first,
termed quiescence, the cells are maintained in a non-proliferative (GO) state that is
marked by severe alterations in cellular biochemical activity (1,2). This includes; 1)
a reduction in size due to increased degradation of protein and mRNA without an
increase in their resynthesis; 2) macromolecular synthesis is one third as rapid in
quiescent cells as in proliferating ones; 3) enzyme and transmembrane transport
activities are severely reduced in G0 cells; and 4) ribosomes are monosomal rather
than polysomal (3). Cells can be induced to enter this state by various methods
including serum deprivation (2,4,5,6), contact inhibition (7,8), and drugs (9). Cells
can survive in this state for varying lengths of time or can be induced to enter the
second reproductive state, that of proliferation. The proliferative state is defined as
the increase in cell number resulting from the completion of the cell cycle, as
compared to growth, which is an increase in cell mass. Extracellular factors
determine whether a quiescent cell will reenter the cell cycle and begin to proliferate

again, or if a proliferative cell continues to divide or enters a quiescent G0 state (2).

The eukaryotic cell cycle is divided into four phases, G1, S, G2, and M (1) (Figure

 

 

 

 

1). During G1 or gap phase the cells prepare for DNA synthesis. The time interval

for G1 can vary considerably between different cell lines, and is also dependent on
the growth state of the cell. The G1 phase of the cell cycle is divided into four
subphases termed competence, entry, progression, and assembly (reviewed in 2). The
culmination of all the activities which constitute these phases is the activation and
production of all the macromolecular components needed for the replication of the
cellular genome, which represents the next phase of the cell cycle, S phase. During
S phase the entire DNA content of the nucleus is replicated completely and
precisely in a period of a few hours (2,11). In addition to DNA replication, the
complex architecture of the chromosome is also duplicated; this includes the
assembly of nucleosomes and chromosomal scaffolding (11). When the entire DNA
content is duplicated the cells enter a second gap phase termed G2, in which the
cells prepare for mitosis or M phase (1). During mitosis actual cell division takes
place through a series of sequential steps in which the segregation and division of
nuclear and cytoplasmic components occurs (12). After mitosis the cell either
continues to proliferate or enters a quiescent G0 phase, depending on extracellular
factors.

The complex events which make up the eukaryotic cell cycle are tightly
coordinated and controlled by the interaction of various regulatory systems. Genetic
experiments have demonstrated that the cell cycle is regulated at two important, and
well defined control points. The first occurs during G1 before the Gl/S transition,

and the other occurs in G2 before the G2/M transition. The progression of cells

through G1 and into S is controlled at a point in G1 designated the restriction (R)

 

 

 

 

(Division) Hows

S phase
(DNA synthesis)

 

Figure 1. Mammalian cell cycle with a 16 hr doubling time (10)

 

 

 

 

point in mammalian cells (1,2,13,14), and START in yeast (15,16).

The molecular and biochemical events that are involved in the regulation of cell
progression at (R) have only recently begun to be identified. It has been shown that
once a cell has progressed beyond the restriction point it is committed to complete
DNA synthesis and one round of cell division, even in the absence of growth factors
(13,14). Many different molecules may be involved in passage of cells through (R).
One class of molecules which have been shown to be involved at both the G1 / S and
G2/ M transitions are the cyclin dependent kinases (cdks), whose activities are

controlled by another class of molecules called cyclins.

Cyclins and gyclin Dependent Kinases

Cyclins were first identified in marine invertebrates as proteins whose levels
oscillate dramatically throughout the cell cycles (17,18,19). Since these initial studies
five classes of human cyclins have been studied and characterized; they are cyclin A,
B, C, D, and E. Cyclins have been implicated in the control of cell cycle progression
at both control points in the cell cycle. The best understood of these is the
regulation of the cells entry into mitosis, which is controlled by a mitotic protein
kinase that contains two major regulatory subunits (20). The first is a 34 kD catalytic
subunit that is the product of the cdc2 (cell division cycle) gene in
Shizosaccharomyces pombe (p34fdcz) (21,22), and its homologs in other eukaryotes.
The second, a regulatory protein subunit which is required for the protein

serine/threonine kinase activity of p34Cdcz and possibly also for targeting the complex

to different parts of the cell, is cyclin B (cdc13)(23—27). Together, these two

 

 

 

 

regulatory proteins form the Maturation Promoting Factor (MPF)(28,29), which is

responsible for regulating entry into mitosis. This complex accumulates throughout
S and G2 in a form that is kept inactive through the phosphorylation by weel of
amino acids tyrosine 15 and threonine 14 within the p34°dcz ATP binding site.
Removal of these inhibitory phosphates at the G2/ M transition by the product of the
cdc25 gene activates the p34°d°2/cyclin B complex to drive the cell into mitosis Exit
from mitosis is regulated by the degradation of cyclin B via a ubiquitin mediated

4cdc2

pathway during anaphase, generating an inactive p3 monomer (reviewed in 30-

33).

Control of the Gl/S transition in mammalian cells appears to be mediated by a
different set of cyclins and cdks than the G2/M transition. Growth factors act during
CI to regulate the accumulation of critical gene products that are required for entry
into S phase. Signals transduced from the extracellular environment impinge on the
cell cycle control machinery and determine whether a cell should proceed through
the cell cycle by progressing through (R) or enter a quiescent state (1,2). Cyclin D
appears to exert its influence on cell cycle progression by acting as a growth factor
sensor in cells which have been stimulated to reenter the cell cycle from a G0 state
(34,35,36). In serum stimulated cells cyclin D1 is localized to the nucleus and is
rapidly degraded as cells enter S phase (36,37). It has been proposed that nuclear
exclusion and/or cyclin D1 degradation are required for progression through S phase
(37). Cyclin D, in association with the cyclin dependent kinase cdk4, binds to and

phosphorylates the retinoblastoma gene product (pr) and the structurally related

protein p107 (38,39,40). Phosphorylation of pr has been shown to be a critical

 

 

 

 

 

regulator of its growth suppressing activities. During most of G1, pr can be found

in a hypophosphorylated state, and while in this state is a potent inhibitor of cell
cycle progression. Phosphorylation late in G1 inactivates the growth inhibition by
pr and allows cells to progress through the remainder of the cell cycle (41). Cyclin
D mRNA and protein synthesis is induced when murine macrophage cells are
stimulated by colony stimulating factor 1 (CSF-l) (35). The CSF-l receptor is a
protein tyrosine kinase, thus linking the regulation of the GO/Gl transition by
receptor tyrosine kinases and genes which regulate cell proliferation. Cyclin D1,
when overexpressed or deregulated, has been shown to function as an oncoprotein
in some human tumors (37). Lastly, cyclin D was shown to associate with
Proliferating Cell Nuclear Antigen (PCNA), a known DNA replication and repair
factor important during the S phase of the cell cycle (42).

Cyclins A and E have also been suggested to be involved in regulating
progression through the Gl/S transition. Both of these cyclins bind to the cyclin
dependent kinase p33“”"2, and while in this complex associate with both the
transcription factor E2F and the Rb related protein p107 (43,44). E2F has been
shown to be a critical transcription factor for many S phase inducible genes and may
be important for the transcription of genes in other phases of the cell cycle as well
(45,46,47). The Rb protein has been shown to directly bind E2F, and through this
association repress transcription of many vital genes (48,51,52). A possible model for
regulation by cyclin E would involve Cyclin E/p33€de binding to a complex of p107

and E2F during late G1, competing it away from pr and thus preventing repression

of E2F at this stage in the cell cycle. As cells traverse the Gl/S boundary the cyclin

 

 

 

E/p107/E2F association decreases, concurrent with the beginning of a cyclin

A/p107/E2F association that increases throughout S into G2 (43,53). Suggesting a
role for cyclin A in DNA synthesis. The Cyclin A/E2F and E/E2F complexes are
both dissociated by the EIA oncogene, suggesting these cyclins may be targets of
oncogenic signals (54,55). It has been hypothesized that cyclin A and its associated
kinase activity functions at both the G1 /S interface and during S phase to facilitate
progression through these two phases of the cell cycle (56). A recent report has
identified a cyclin A/p33Cdk2 binding site in the human thymidine kinase promoter,
which has been postulated to be involved in its regulation at the Gl/S border (57).
In addition to its involvement during G1 and S, Cyclin A is also thought to be
involved in the regulation of progression at the G2/M transition by its association

4cdc2

with the cyclin dependent kinases p3 (56). So we can envision two possible roles

for cyclin A in the cell cycle. During late G1 and S phase cyclin A is associated with

3cdk2 and may help regulate entry into S phase as well as help in S phase

p3
progression/maintenance. During G2 it is associated with p34“2 and is involved in
regulating entry into Mitosis. Exit from mitosis is also dependent on the degradation
of the cyclin A/p34Cdcz complex in addition to cyclin B degradation (58). In summary
the Gl/S transition has been postulated to be regulated by cyclins E and A in
association with p33fdk2, with cyclin D1/cdk4 mediating events earlier in G1 and
possibly mediating the progression through (R). Cyclin B and cyclin A in association

with p34CdC2 regulate entry into mitosis. A schematic representation of the temporal

expression of the cyclins and the cdks associated with each cyclin class is shown

illustrated in Figure 2.

 

 

 

 

 

RB
DephOSphorylation

 

. Cyclin
BIA + C002
Cyclin

D’s + CDK2

Cyclin CDK4

A + CDK2 CDK5

RB
Cyclin PhOSphoryIatton
E + CDK2

Figure 2) Expression of mammalian cyclins and cdks during the cell cycle (59)

 

 

 

 

S Phase Regulated Genes

When quiescent mammalian cells are induced to reenter the cell cycle, a series

of tightly regulated events occur in association with the activation and production of
the molecules needed for DNA synthesis, and ultimately cell division. Growth factors
and their receptors at the cell surface begin a biochemical cascade of events that
activate genes encoding transcriptional regulatory proteins. These proteins in turn
regulate the genes needed for DNA synthesis. Determining how these S phase
regulated genes are controlled and how these controls are integrated into the
biochemical signals that govern progression through G1 into S is vital for
understanding the mechanism of growth regulation. The regulation of these genes
has been found to be at multiple levels and by a variety of different mechanisms.
The best characterized representatives of these S phase inducible genes include
thymidylate synthase (TS), dihydrofolate reductase (DHFR), the replication
dependent histories, and thymidine kinase (TK).

Thymidylate synthase catalyzes the synthesis of thymidylate (TMP) via the de
novo biosynthetic pathway. The levels of its enzyme activity and mRNA increase
approximately 20 fold as cells progress from G0 through S phase. Transcription was
shown to increase only two to three fold over this same period, indicating that TS
mRNA levels are controlled primarily at the post-transcriptional level (60,61). The
human and mouse TS promoters both lack TATAA boxes but are GC rich. The
essential mouse TS promoter is located within 105 nucleotides (nt) of the authentic

AUG codon (62) and contains binding sites for Spl, PEA3 (63), and the proteins

encoded by the ets 1 and 2 proto-oncogenes (64). The 5’ flanking region also

 

 

 

10

contains potential binding sites for the transcription factors AP1, E2F, Octl, SRE,
and Yi. The human promoter elements have not been elucidated yet but contain
several regions of sequence homology to mouse TS (65). The 5’ flanking region and
sequences downstream of the authentic AUG codon are both necessary, but neither
is sufficient for growth regulated expression (66). Chu et al (67) demonstrated that
translation of TS mRNA is controlled in an autoregulatory manner by its own protein
product. This regulation was postulated to be through the proteins possible
interaction with three interconvertable secondary structures in the 5’ UTR, each
containing a stemloop structure (65).

Dihydrofolate reductase catalyzes the conversion of dihydrofolate to
tetrahydrofolate, which is required for the biosynthesis of purines and thymidylate.
DHFR is regulated at the transcriptional and post-transcriptional levels depending
on the growth state of the cell (68). At the transcriptional level, serum stimulated
cells show a 7 to 10 fold increase in the rate of transcription at the Gl/S interface
(69). The mouse and hamster DHFR genes lack a TATAA box but have GC rich
regions in their 5’ flanking regions, which were shown to be the site for
transcriptional initiation. In the hamster DHFR gene four copies of the Spl binding
site were localized to this area and were found to be important for both gene
transcription and specifying the transcriptional start sites (70). Binding sites for the
transcription factor E2F were also localized near the start site and appear to
determine the rate of transcription (45). The mouse DHFR gene contains an E2F

binding site that is recognized by the protein HIPl, which is important for

determining the site oftranscriptional initiation and for facilitating the transcriptional

 

 

 

 

burst at the G1/S interface (71).

The replication dependent histones are proteins whose expression is tightly
regulated during S phase and is coupled to DNA synthesis. They are involved in the
ordered assembly of newly replicated DNA into nucleosomes. The S phase
dependent synthesis of histones results from an increase in histone mRNA levels
which is regulated by both transcriptional and post-transcriptional mechanisms
(72,73), and can be activated rapidly and reversibly multiple times during a single S
phase (73). The approximate 3 to 5 fold increase in transcription between G1 and
S phase is regulated through promoter proximal DNA sequences and their cognate
transcription factors. This regulation is facilitated by quite limited 5’ flanking regions
which contain subtype specific consensus elements (SSCE)(74,75,76). SSCE distances
from the TATAA box are extremely consistent, usually differing by no more than 1
bp. One factor which is known to be directly involved in cell cycle control of histone
transcription is oct-l, which can specifically stimulate histone H2b as much as 20 fold
in vitro upon binding to its SSCE (77).

The rate of histone transcription varies only 3 to 5 fold between G1 and S, which
cannot account for the 20 to 35 fold increase in histone mRNA observed in many
systems, indicating that much of the regulation must be at the post-transcriptional
level. Post-transcriptional regulation of histone mRNA is mediated by two
mechanisms, the first is mRNA stability and the second is 3’ processing of pre mRNA
within the nucleus. A six to eight fold increase in the efficiency of the

endonucleolytic cleavage of the large primary histone mRNA transcript immediately

3’ to a conserved stem loop structure was found to require both the conserved stem

 

 

 

 

loop structure and a purine rich sequence (78-81). Three different trans-acting

factors were shown to be involved in this process (79,80,81), with the U7 snRNP
being the most studied. This regulation so far has only been demonstrated in G0
induced cells, but very little data has been obtained on cycling cells so they may also
be under this type of regulation. The conserved 3’ stem loop structure has also been
implicated in the very rapid decline in histone mRNA concentrations as cells
progress from S to G2. This decline was found to be due to the increase in mRNA

degradation that in part may be autoregulated by free cytoplasmic histones (82).

Thymidine Kinase and its Regulation
Thymidine kinase (TK) (ATPzThymidine 5’ phosphotransferase EC 2.7.1.21) is

a cytosolic enzyme in the pyrimidine salvage pathway which catalyzes the ATP
dependent phosphorylation of thymidine to thymidine 5’ monophosphate (dTMP),
which upon further phosphorylation to thymidine 5’ triphosphate (d'ITP) is utilized
in the synthesis of DNA during S phase (83,84) (Figure 3). Two forms of the enzyme
are present within the cell, a mitochondrial form, and a cytoplasmic form (85,86)
which differ considerably in their biochemical properties (87,88,89). Some of these
differences include isoelectric point, pH optimum, Km value, sedimentation
coefficient, and polyacrylamide gel electrophoresis mobility (87). A critical difference
that has relevance to the present thesis is that mitochondrial TK levels are constant
throughout the cell cycle while levels of the cytoplasmic form are highly regulated

(89). The study of one aspect of this cell cycle dependent regulation of cytoplasmic

TK will be the focus of the present study.

 

 

 

5Phosphoribosyl-1-pyrophosphate (PRPP)

De novo J:
synthesis of 5-PhosphoribosyI-1-amine
purine nucleotides
rated-n by Mitch!“

k-CHO from
tetrahydrofolate

Formylgiycinamide
ribotide (FGAR)

L—NH; from De novo
v synthesis of

glutamine ‘
Formylglycinamidine thymidylate

ribotide (FGAM) Uridylate (UMP)

 

 

 

/Btwed by anniularn mm by
l/‘llF-CHO from fi/lnniclfies
tetrahydrofolate .f'_cHJ from
i _ ' (IMP) - tetrahydrofolate
nosmate
- (a nucleotide) (j E
Q- [ F l . //
Guanyiate (GMP) HG?“ Adenvlate (AMP) Thymidylate (TMP)
- 9
' ' Par ADP
”Gm Inosine “Ana-my K
(hvvo-Ilnthine- . (a nucleoside) °"°3°"°"°)°W‘ ATP
Guanosine “mm b“... I IQ" _ rx
mm) Adenine Rtbose phosphate lmwnidinc
V\\ from PFIPP mm.)
Guanine K-—Ribose phosphatej Hypoxanthine Thymidine
from PRPP (a base)

Figure 3) Pathways to obtain purines and thymidylate for DNA synthesis (10)

 

 

 

 

14

The native form of human TK was determined to have a molecular weight of
96,000 consisting of four 24,000 MW monomers, suggesting a tetrameric structure
for the enzyme (90). Vertebrate TK genes have been cloned from chicken (91,92),
mouse (93), hamster (94,95) and human (97-100). Viral TK genes have been cloned
from Herpes, african swine fever, and numerous pox strains (96,101). Vertebrate TK
genes have been shown to be closely related to pox viral strains but are unrelated to
TK from herpes virus. Sequence comparisons between the vertebrate and the pox
viral genes reveal that the viral genes lack 15 amino acids (AA) from the first 17 N
terminal amino acids, and 30 out of the last 40 AA from the C terminus (Figure 4).
This observation is significant due to the fact that, unlike the vertebrate genes, pox
viral TKs are not regulated in a cell cycle dependent manner. Human TK genomic
and cDNA clones have been isolated by many groups (97-100), and the human TK
gene and its flanking regions have been sequenced in their entirety (102). The TK
gene is 12.9 kb long and contains seven exons which upon processing produce a 1,430
bp mature mRNA (98,102).

The early development of an effective assay for TK activity has allowed its
regulation to be studied extensively. Until recently, with the advent of new molecular
biological techniques, the study of the regulation at other levels could not be
investigated. Subsequent to the successful cloning of the TK gene and its promoter
from mammalian and viral sources, study of the molecular aspects of TK regulation
were initiated. The majority of these studies utilized cells that are inherently TK

minus, transfected with recombinant TK’“ clones. Kit et al (103) first isolated TK'

murine fibroblast cell mutants (LMTK') in 1963, and since then other mouse, human,

 

 

 

15

 

 

 
 

 
  
       

nun HSCINLPYVLPGSPSXTR o OVILGPHFS STELMRRVRRFOIAQYKCLVT YAKDTRY SSSF c
HOU HSYINLPTVLPSSPSKTR o QVILGPHFS STELHRRVRRFOIAOVKCLVI vnxornv SNSF s
CHI HNCLTVPGVHPGSPGRPR o QVlFGPHF STELHRRVRRFOLAQYRCLLV YAKDTRYCTTGV 5
vv HN H QLIIGPMFS STELIRRVRRYOIAOYKCVTI YSNDNRY GTGL u
ssv HY HnHLIIGPHFA STELIRLVRRYOIAKHKCLVV vexotnv cnsv c
cpox HD v HLIIGPHF STELIRIVKRYOIAOYKCCVVIYLKDIRY cnsv Y

‘ FPOX H58 5 HVXTGPHFS TSELVRRIKRFHLSNFXCIII HCGDNRYNEDDINKVY
Asrv HNIIRKLKP T SLVLePn A TTFLIHCIYHLERLEKKVVFI srxNTRoKTIxTHsct

no
Hun THDRNTHEALPACLLRDVAQEAL GVAVIGIBEGO PDIHEFCEAH NA rv net aRK G
nou THDRNTHDALPACHLRDVTDELL GVAVIGIUEGQ PDIVDFCEHH NE rv not ORXA s
CH1 THDRNTHEARPACALCDVYGEAL GSAVIGIDEGQ POIVEFCEKH NTG‘IV oer QRKA 6
vv THDKNHFEALEATKLCDVLESIT DFSVIGIDEGO PDIVEFCERH-NE iv oer ORKP N
srv THDNHSITAVCTPSLOKIDSVAE NAEVIGIUEGO PNIATFCERH~NRG VL DGT can 5
cpox runnuuvsaisrrunvnvvnxtn NFDIIGTDEGO KDIVSFSENH'NHG ll DST ORKE N
FPOX THDLLFHEATASSNLSVLVPTLLNDGVQVIGIUEAO LDIVEPSEscgNL T No KREL G
ASFV QLRPKOCKIIESTQLSDVGSLT DIHAVVVU AH DDIIK CRT $555 It NA EQK P
2w
HUN AtLNLVPLAesgvxttAv-nEic REAAYTK'LGTEKEVEVI anox HSVIRLgYFKXASGQPAGPDNK
nou SILNLVPLAES VKLTAVIHEIF REAAYTK'LGLEKEVEVI rADK Hsv-RLIYFKKSSAQTAGSDNK
CHI SILNLVPLAES VKLNAV HE-Y REASYTK'LGAEREVEVI nADK HSVIRA
VV NILNLIPLSEH VKLTAVIHK IF KEASFSXI'LGEETEIEII HNDM OSVIRXIIYIDS
srv NISELIPLAEN TKLNAVIHYIY KNGSFSK'LGOKHEIEVI rsox st
CPOX DILXLIPLSEK TKLNAVIHE Y KDAAFSKI’ITKEKEIELI t-KEK
FPOX NVYKLLSLAET SSLTAIIVKIY ansrsx VTENKEVHDI exox
ASFV PIVRIFPYCSH Kyrcnrgnx-NQHHACFNV KNADKTLILA rSEL VTcgNNI LKNTFIKDLQPIXY
2H
HUM ENCPVPGKPGEAVAAPKLFAPQO[LOCSPAN
HOU NCLVLGQPGEALVVRKLFASOOVLOYNSAN
CHI ENVPHGVKQLDHPASRKIFAS

Figure 4) Amino acid sequence comparison between TK from viral and vertebrate
sources. Sequence designation is as follows: HUM, human cytoplasmic TK; MOU,
mouse cytoplasmic TK; Chl, chicken cytoplasmic TK; VV, Vaccinia virus; SFV,

Shope fibroma virus, CPOX, Capripox virus; FPOX, Fowlpox virus. Sequence

homology between all species are shown in black boxes. (101)

 

 

 

 

16

rat, hamster, and bacterial TK‘ cell lines have been isolated (104,105,106).
Endogenous TK+ cell lines have also been used in many studies.

The regulation of TK has been shown to be tightly coupled to the growth state
of the cell. In quiescent G0 phase cells that have been stimulated to reenter the cell
cycle by mitogenic agents, TK enzyme activity is very low in early G1 and increases
20 to 80 fold at the G1 /S transition, with mRNA and protein levels paralleling that
of activity (107-110). This increase was shown to require both protein and mRNA
synthesis due to the sensitivity of this induction to inhibitors of protein and mRNA
synthesis. In cycling HeLa cells, the same increase is seen at the G1 / S interface, but
without a substantial increase in mRNA levels throughout the same period (111).
Also when TK cDNA is expressed from a heterologous non-cell cycle promoter TK
mRNA levels are high during G1 but protein levels remain low (112,113). Thus TK

regulation appears to be operating at at multiple levels within cells.

 

Transcriptional Regulation of Thymidine Kinase

Work in our lab has shown that when quiescent G0 phase CV-1 (african green
monkey kidney) cells are serum stimulated to reenter the cell cycle, TK mRNA levels
increase 15 to 20 fold between G1 and S, with enzyme levels paralleling that of the
mRNA (108). Nuclear run on transcription assays of the same cells showed a 6 to
7 fold increase in transcription at the Gl/S interface when induced with serum, and
a 3 to 4 fold increase when mitogenically stimulated by SV-40 infection (110). In

similar studies, a 2 to 4 fold increase in transcription was detected in serum

stimulated Balbc/3T3 cells, while an 11 fold increase was observed in the

 

 

 

 

 

endogenous TK from mouse L929 cells (109,114).

Kriedberg and Kelly (115) were the first to investigate the regions within the
human TK promoter that are required for its activity. Utilizing a series of promoter
deletions linked to a human genomic clone, they measured the ability of these clones
to transform mouse L TK' cells to the TK+ phenotype. From these studies they
defined a functional promoter to within an 83 bp region upstream of the mRNA cap
site. Mutants containing fewer than 53 bp had almost no transforming ability,
locating major functional promoter elements between 53 bp and 83 bp upstream of
the mRNA cap site. They also demonstrated that the TK promoter region contains
two transcriptional start sites at +1 bp and +7 bp, a TATAA box located at -24 bp,
two inverted CCAAT boxes at -44 bp and -75 bp, and three GC rich Spl binding
sites at -115 bp, -229 bp, and -248 bp from the transcriptional start site.

Arcot et al (116) also performed a functional analysis of the TK promoter region
by determining the strengths of various TK mutants. Deletion and site directed
mutant TK promoter fragments were linked to the bacterial chloramphenicol
acetyltransferase (CAT) gene, and CAT activity was assayed in transiently transfected
mouse L TK' cells. Promoter fragments containing 251 bp and 139 bp upstream of
the cap site retained 81% and 64 % activity respectively relative to the 457 bp full
length promoter. Promoter fragments containing 88 bp and 58 bp fragments contain
28% and 19% activity respectively. To delineate the region(s) within the promoter
which convey Gl/S phase specific regulation, a number of studies were initiated

where TK promoter mutants were linked to non-cell cycle regulated genes. The

results of these studies determined that the region between -67 bp and -135 bp in the

 

 

 

 

18

TK promoter is sufficient to confer G1/ S phase regulation to these genes
(112,117,118,119). Interestingly, this area contains a pair of sequences that resemble
the Yi binding site in the mouse TK promoter. When these sites were mutated the
reporter gene used in the study was constitutively expressed, thus eliminating the S

phase specific transcriptional regulation (120).

Post-Transcriptional Regulation of Thymidine Kinase

Post-transcriptional regulation of TK expression was suspected for two reasons.
First, the 4 to 7 fold transcriptional increase at the G1 / S border does not account
for the 20 to 80 fold increase in enzyme activity during the same time interval.
Secondly, in some systems the regulation and levels of TK mRNA were demonstrated
to be independent from the regulated levels of TK protein and enzyme activity. The
mechanisms for this type of regulation were reported to be at multiple levels
including regulated nuclear processing, differences in mRNA half life, translational
regulation, and differential regulation of protein half life. In Balbc/3T3 cells, the
level of nuclear mRNA precursors and mature mRNA was shown to increase
significantly at the Gl/S border, while TK mRNA was bearly detectable in either the
nucleus or cytoplasm during G0 and most of G1 (121). This suggests a cell cycle
specific change in nuclear processing of TK mRNA. In addition the half life of
mature TK mRNA was reported to be 2 to 3 fold greater during S phase than during
G0 or G1 (109).

In addition to the post-transcriptional regulation of mRNA levels, we and others

have shown that TK enzyme activity and protein levels are regulated independently

 

 

 

 

 

19

of mRNA levels when the TK cDNA is expressed from a heterologous non-cell cycle
regulated promoter (112,113). These findings suggest that sequences within the TK
cDNA are sufficient for this type of regulation. To further investigate these findings,
our laboratory linked human TK cDNA to the SV-40 early promoter, transfected this
construct into TK' cells and measured levels of mRNA, protein and enzyme activity
throughout G1 and S phase after serum stimulation. The results of these studies
revealed a 2-5 fold increase in mRNA levels early in G1 which then remained
constant for the remainder of the cell cycle. In contrast, protein levels and enzyme
activity remained low throughout G1, paralleling that of mRNA, but were induced
20 - 50 fold at the Gl/S border (113). These results have been confirmed by
experiments outlined in chapter 2 of this thesis. This study also demonstrated that
post-transcriptional regulation in serum stimulated cells was not dependent upon the
5’ untranslated region (UTR), the authentic TK translational start site, or the first
16 amino acids of the coding sequences, suggesting that regulation does not occur at
the level of translational initiation. Regulation also does not involve the
polyadenylation site, or 430 bp of the 3’ UTR. It was concluded from these studies
that TK protein levels and enzyme activity are regulated at the translational and/or
post-translational levels in a cell cycle specific manner.

Gross and Merrill observed translational regulation of the chicken thymidine
kinase gene transfected in mouse skeletal muscle cells induced to terminally
differentiate (122,123). In this study, when mouse skeletal muscle cells containing

the chicken TK gene were induced to terminally differentiate, TK mRNA levels

remained relatively constant while pulse labeling experiments revealed a 10 fold

 

 

 

 

20

reduction in the rate of TK protein synthesis. The polysomal distribution of the TK
mRNA did not change during this period, suggesting that translational control was
not at the level of initiation. Also, the half life of the TK protein was the same in
both proliferating and terminally differentiated cells.

In many of the previously mentioned studies, various aspects of TK regulation
have been investigated in quiescent G0 phase cells that have been mitogenically
stimulated to reenter the cell cycle. In these studies, it was revealed that both TK
mRNA and enzyme activity are highly induced at the G1 /S border. To investigate
the regulation of TK in continuously cycling cells, Sherley and Kelly (111) measured
levels of TK mRNA, protein and enzyme activity in cycling HeLa cells synchronized
by centrifugal elutriation. In contrast to the results obtained in serum stimulated
cells, the increase in TK protein at the Gl/S interface was not accompanied by a
proportional increase in TK mRNA. In this system the overall pattern of protein
expression revealed low levels during G1, a sharp increase at the Gl/S border that
accumulated throughout S and G2 into M phase, followed by a rapid decline back
to G1 levels after mitosis. While the amount of TK protein increased 15-25 fold
from G1 to M, the steady state levels of TK mRNA increased less than 2-3 fold
during the same period. Two post-transcriptional mechanisms were found to be
reported to be involved in this increase in TK protein levels and enzyme activity. In
the first, pulse labeling experiments revealed a 10 fold increase in TK protein
synthesis between G1 and S phase. Secondly, the stability of the TK polypeptide

decreases abruptly after mitosis but before cytokinesis, resulting in a rapid clearance

of the enzyme from newly divided G1 cells. Protein synthesis is not required for this

 

 

 

21

M phase specific degradation of TK. In pulse chase experiments, the half life of the
TK polypeptide was in excess of 40 hours from late G1 through metaphase, while the
half life decreased to < 1 hour around the time of cell division (111). This
metaphase specific change in half life is reminiscent of the regulated degradation of
cyclins A and B, although no sequence similarities between TK and these cyclins
have yet been found.

The mechanism for the possible translational regulation is still unknown, but a
region within TK polypeptide required for the differential protein half life has been
identified (124). Deletion of the carboxyl-terminal 40 amino acids of the TK
polypeptide stabilized the protein at mitosis and completely abolished the cell cycle
regulated degradation of the protein. The same inhibition of regulated degradation
occurred when all or part of the Escherichia coli B-galactosidase polypeptide was
fused to the carboxyl terminus of the TK protein. None of the modifications to the
TK polypeptide utilized in this study had an effect on the specific activity of the
protein (124). To investigate if this cell cycle regulation of protein half life occurs
in quiescent cells stimulated to reenter the cell cycle, the same deletion mutants and
fusion proteins were expressed from a heterologous non-cell cycle regulated promoter
in transfected murine TK' cell lines (125). The results of these studies demonstrated
that these modifications to the TK polypeptide allowed its expression during the G0
and G1 phases of the cell cycle. This is in sharp contrast to the normally low levels
of TK protein found during G0 and G1 phases in serum stimulated cells. This

suggests that the mechanisms that control TK protein half life in cycling cells are

similar to those in quiescent cells.

 

22

In the current study we set out to determine if the human thymidine kinase
polypeptide is regulated translationally, or by differences in protein stability, or both
of these mechanisms in a cell cycle specific manner in serum stimulated cells. The
results of these studies, outlined in chapter 2, indicate that the rate of translation of
TK mRNA is constant as stimulated cells progressed from G1 to S. In contrast to
this, the stability of the TK polypeptide was 10 fold lower in G1 than it was in S
phase implying that post-transcriptional regulation in our system is solely at the level
of protein stability. These results are in contrast to those reported by Sherley and
Kelly (111) in cycling HeLa cells, where TK was translated 10 times more efficiently
in S than in G1. Also, they reported that TK was degraded after mitosis but was

stable during most of G1 through S and G2.

Protein Degradation Mechanisms

The majority of intracellular proteins are in a dynamic state of synthesis and
degradation. The degradation process is highly selective, as demonstrated by the
diversity of intracellular degradation rates of individual proteins. Half lives of
proteins range from minutes to days, and understanding the molecular basis for this
heterogenicity has been a major challenge to researchers studying protein
degradation (reviewed in 126). Our knowledge of protein synthesis is far greater
than our knowledge of protein degradation, the mechanisms of which are still mostly
unknown. One reason for this discrepancy is that multiple mechanisms for protein

degradation exist within cells. Recent studies have begun to investigate the various

mechanisms by which proteolysis occurs. These include ubiquitin dependent

 

 

 

23

proteolysis, calcium dependent pathways of proteolysis, lysosomal pathways, cytosolic
pathways which are ATP dependent but ubiquitin independent, and pathways which
are independent of both ubiquitin and ATP. Multiple molecular determinants have
been found which function in the above described proteolytic mechanisms. While the
pathways of protein degradation are beginning to be characterized, very few examples
of regulated protein stability have been identified. One of the best studied is the
regulated degradation of cyclins B and A by ubiquitin. This involves a highly
conserved 9 amino acid "destruction box" in the N-terminal of the protein and critical
down stream lysine residues (127). Ubiquitin is a 76 amino acid protein which
targets abnormal and normal proteins for rapid degradation. Ubiquitin is first
activated by multiple ubiquitin specific enzymes in an ATP requiring process, the
activated ubiquitin is then conjugated to substrate proteins via isopeptide bonds
between the carboxy terminus of ubiquitin and e-amino groups of internal lysines
within the protein. Proteins are targeted for degradation when a chain of ubiquitin
molecules forms at this site by the ubiquitin molecule itself being ubiquitinated thru
a lysine residue at position 48 (128,129,130). Several molecular determinants target
proteins for ubiquitin mediated degradation including the following: denatured
proteins are more efficiently conjugated with ubiquitin than native proteins; the at-
amino terminal amino acids must be unblocked, it should not contain amino acids
with large side groups; and the N terminal amino acids fit the N-rule for selective
protein breakdown, which states that when certain amino acids are present at the N-

terminus, the protein is targeted for degradation (131). In addition to these, rapidly

degraded proteins were found to contain PEST sequences (132), and certain peptide

 

 

 

 

24

sequences (KFERQ) target proteins to lysosomes for enhanced degradation during
starvation (126).

Thymidine kinase appears to provide another model for studying the regulation
of protein stabilty. While the pattern of it’s degradation during mitosis appears to
mimic that of cyclins A and B, it differs from these two proteins by being specifically
stabilized at G1 / S. No common elements, such as the "destruction box" or critical
lysine residues, have as yet been found between TK and cyclins A and B. The study

of its regulation will therefore provide the opportunity to identify and study novel

pathways involved in the regulation of protein stability.

 

 

2)

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128) Finley, D., (1991) Annu. Rev. Cell Biol. 7: 25-69

 

 

33
129) Rechsteiner, M., (1991) Cell 66: 615-618

130) Chau, V., J .W. Tobias, A. Bachmair, D. Marriott, D.J. Ecker, D. Gonda, and A.
Varshavsky (1989) Science 243: 1576-1583

131) Bachmair, A., D. Finley, and A. Varshavsky (1986) Science 234: 179-186

132) Rogers, 8., R. Wells, and M. Rechsteiner (1986) Science 234: 364-368

 

 

CHAPTER II

REGULATION OF THYMIDINE KINASE IN SERUM STIMULATED CELLS

By

Marc A. Carozza and Susan E. Conrad

Submitted to the Journal of Biological Chemistry

34

 

 

 

 

The mechanism of post-transcriptional regulation of thymidine kinase (TK)
enzyme levels following serum stimulation of quiescent cells has been investigated
using stably transfected Rat 3 cells containing the human TK cDNA linked to a
hybrid SV40/human TK promoter. These cells expressed a wild type TK mRNA at
relatively constant levels during the first G1 and S phase after serum stimulation. In
contrast, TK enzyme activity and protein levels were low during G1 and increased
dramatically as the cells entered S phase. The specific activity of the protein did not
vary between G1 and S, and pulse labeling experiments indicated that the rate of
translation of TK mRNA was also constant throughout G1 and S. However, pulse-
chase experiments indicated that the half-life of TK protein increased approximately
10 fold between G1 and S-G2. Thus, the increase in TK protein at the G1-S
transition was the result of a change in stability of the protein. Finally, 32P-labeling
experiments indicated that the TK protein is phosphorylated. Truncation of the TK
protein by deleting 16 N-terminal amino acids resulted in a large reduction in
phOSphorylation, but the stability of the truncated protein was regulated similarly to
the wild type. These results suggest that the wild type pattern of phosphorylation is

not required for the regulation of protein turnover.

  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
   

 

 

 

 

36

INTRODUCTION

We have used the human thymidine kinase (TK) gene as a model system to
investigate the mechanisms that regulate the transition from G1 to S phase. TK
enzyme levels are tightly regulated by both the growth state and cell cycle position
of the cell; rapidly dividing cells contain high levels of enzyme activity while resting
cells contain low or undetectable levels (18). When quiescent serum-starved cells are
mitogenically stimulated, TK activity remains low throughout G1, increases sharply
as cells enter S phase, and remains elevated throughout S and G2 (12). TK enzyme
levels are also regulated in continuously cycling cells; they are low in G1, increase
at G1-S, and remain high during GZ and into M (22).

The induction of TK activity at Gl-S is the result of both transcriptional and
post-transcriptional controls. In mitogenically stimulated cells, the enzyme activity is
paralleled by the mRNA level, which is low in G0 and G1 and highly induced as
cells enter S phase (24). The increase in TK mRNA at Gl-S is largely the result of
transcriptional activation of the promoter (15,21,23,27). The sequences responsible
for regulation of the human TK promoter during serum stimulation have been
mapped to a small region that contains binding sites for the transcription factor E2F
(17). This transcription factor interacts with a number of cell cycle regulatory
molecules including pRB, p107, cyclin A, and a cdk kinase, and has been proposed

to be involved in the induction of genes at Gl-S (20). Complexes containing many

 

 

m

37

of these molecules reportedly form with both the human and mouse TK promoter
(7,17) thus, TK transcription appears to be a target for cellular molecules that
regulate the transition from G1 to S phase.

Several years ago, we and others demonstrated that TK mRNA levels do not
parallel enzyme levels during serum stimulation when the human TK cDNA is
expressed from a heterologous promoter (11,27). If the TK cDNA is expressed from
the SV40 early promoter, mRNA levels are rapidly induced 2-5 fold after serum
stimulation and are then relatively constant for the remainder of G1, S and G2. In
contrast, TK enzyme activity remains low throughout G1 and is highly induced as the
cells enter S phase. Thus, in addition to being regulated transcriptionally, TK enzyme
expression is also regulated translationally or post-translationally near the Gl-S
transition following serum stimulation. Using a series of mutant TK cDNAs
containing deletions at the 5’ 0r 3’ ends, we demonstrated that the translational/post-
translational regulation observed is independent of the entire 5’ untranslated region,
the first 48 nucleotides of protein coding sequence, and the final 430 nucleotides of
the 3’ untranslated region (11).

Translational and/ or post-translational regulation of TK expression has also
been demonstrated in two additional systems. In continuously cycling Hela cells, TK
mRNA levels fluctuate only 2-3 fold during the cell cycle, while enzyme activity and
protein levels vary 10-20 fold (22). The regulation of TK in cycling cells is the result
of both a 10 fold increase in the rate of translation of TK mRNA during S and G2

relative to G1, and a dramatic destabilization of the protein between metaphase and

cytokinesis (14,22). Regulation in cycling cells is dependent upon sequences at the

 

 

 

 

38

carboxyl terminus of the protein, since both deletion of the C-terminal 40 amino
acids and fusions of fi-galactosidase to the C-terminus lead to constitutive expression
of TK throughout the cell cycle (14). The same C-terminal mutations also disrupt
regulation of the gene in serum stimulated cells when it is expressed from a
heterologous promoter (13). Translational regulation of TK has also been reported
in differentiating myoblasts, where the rate of translation of TK mRNA decreases
approximately 10 fold as cells differentiate (8,9,19). No change in the stability of the
protein was detected in these studies.

In the current report, we investigate the molecular basis for the post-
transcriptional regulation of TK protein levels in serum stimulated cells. Our
experiments indicate that both the specific activity of the protein and the rate of
translation of the mRNA is constant throughout most of G1 and S, while the stability
of the protein increases approximately 10 fold between G1 and S. Our experiments
demonstrate that the human TK protein is phosphorylated, but because of the low
levels of TK protein in G1 we were unable to quantitate the extent of
phosphorylation as a function of cell cycle position. We have also examined the
regulation of a mutant protein lacking 16 amino acids from the amino terminus. This
region of the protein was potentially involved in regulation, since it is present in
cellular enzymes, but not in related pox-viral enzymes that are not cell-cycle or
growth regulated (1,3). Phosphorylation of the truncated protein was greatly reduced,
but its stability was regulated similarly to the wild type. Thus, the stability of TK

protein does not appear to be regulated solely by its extent of phosphorylation.

 

 

 

MATERIALS AND METHODS

Cell culture and DNA transfections

Rat 3, TK' cells (25) were grown in Dulbecco’s modified Eagle medium
(DMEM) supplemented with 10% (v/v) Hyclone calf serum. Transfection of TK
cDNA clones into Rat 3 cells was performed as described by Wigler et a1. (28). TK+
transfectants were selected by growth in DMEM plus 10% calf serum supplemented
with 1 x HAT (Hypoxanthine, Aminopterin, Thymidine) medium (Gibco). After 1-2
weeks of selection, plates containing 50-100 colonies were trypsinized, giving rise to
pooled populations of transfectants. These pools of Rat 3 (TK+) cells were
maintained in medium supplemented with 1 x HAT. Cells used in serum stimulation
experiments were passed once in medium supplemented with hypoxanthine and
thymidine (HT), and then grown in unsupplemented medium until the cells reached
confluence (4-5 days). To insure synchronous populations of cells, the confluent
monolayers were serum starved for 24 h in DMEM containing 0.1% calf serum, and
were then stimulated by replacing the media with fresh media containing 10% calf

serum.

Plasmid constructions.
The plasmids pHuTK, pSV-huAScaTK and pSV-huASmaTK have been

described previously (11). The plasmid pESVTK contains the human TK cDNA

 

 

40
linked to 67 bp of 5’ flanking sequence from the human TK gene (Fig. 1). This 67

bp fragment, which contains a TATAA element and one inverted CCAAT element,
functions as a minimal but unregulated promoter in Rat 3 cells (21). To obtain high
levels of TK expression, a Pqu to Hind III enhancer fragment from SV40 DNA was
cloned upstream of the TK gene in an orientation where the SV40 early promoter

is directed away from the TK cDNA.

RNA isolation, and Northern blotting.

Total cellular RNA was isolated and analyzed as described previously (23).

Briefly, 30 ug of RNA from each sample were electrophoresed on 1.2% agarose gels
containing 2.2 M formaldehyde. After electrophoresis, gels were photographed under
UV light to ascertain that equal amounts of RNA were loaded in each lane.
Hybridizations were performed using a 1.2 kb SmaI-BamHI fragment from the
human TK cDNA as a probe. The DNA fragment was radiolabeled using a random

prime labeling kit (Boehringer Mannheim) and a32P-dCTP.

(Moplasmic extraction, and TK enzyme assay.

Cytoplasmic extracts were prepared and TK enzyme activity was assayed as
previously described (23). Briefly, plates were washed twice with PBS and cells were
harvested in 1 ml of cold PBS. Cells were pelleted, and resuspended in 100-200
ul / plate of NP-40 lysis buffer (50 mM Tris-HCl, pH 8.0, 3.6 mM [i-mercaptoethanol,
0.5% nonidet P-40). Nuclei were pelleted by centrifugation at 12,000 x g for 1 min,

and the supernatents were removed and frozen at -70°C. To determine TK activity,

 

 

 

41

20 ul of extract, 44 ul H20, and 16 ul of 5x reaction buffer were combined on ice to
yield a final concentration of 50 mM Tris-HCl (pH 8.0), 15 mM NaF, 3.6 mM 3-
mercaptoethanol, 2.5 mM MgC12, 0.08 mM thymidine, 5 mM ATP, and 50 uCi/ml
of [3H] thymidine (40 Ci/mmole). The reaction mix was incubated for 30 min. at
37°C, then heated at 100°C for 5 min to stop the reaction. Control reactions were
performed by boiling the reaction mix (minus ATP) without prior incubation.
Reactions and controls (20 11.1) were spotted on duplicate Whatman DE81 anion-
exchange filters, air dried, then washed three times in 1 mM ammonium formate,
once in methanol, and dried. The total amount of [3H]-thymidine in the reactions
was determined by counting filters without washing. Filters were placed in
scintillation vials, and the dTMP product eluted by adding 1 ml of 0.1 M HCl, 0.2 M
KCl to each vial and agitating for 30 minutes. Liquid scintillation fluid was added,
and [3H] levels were measured using a scintillation counter. TK activity was expressed

as the percent conversion of substrate per microgram of protein in the extract.

81 nuclease analysis

A 630 bp EcoRI-Mlul fragment was excised from pHuTK, which contains the
entire TK cDNA linked to the human TK promoter (Fig. 1). The MluI site is at
position +176 in the cDNA relative to the transcriptional start site. The fragment
was labeled with T4 polynucleotide kinase (United States Biochemicals) and y32P-
ATP. The labeled DNA (20 ng) was co-precipitated with 15 pg of total RNA,
resuspended in 10 ul of hybridization buffer (80% formamide, 0.4 M NaCl, 0.04 M

Pipes pH 6.4, 1mM EDTA), heated at 90°C for 3 min, then immediately transferred

 

 

 

42

to a 60°C water bath and incubated for 12 hours. The samples were diluted to 200
pl with S1 salts (0.28 M N aCl, 0.05 M sodium acetate pH 4.6, 4 mM ZnSO4), divided
equally between 3 tubes, and digested with 50, 75, or 175 units of $1 nuclease
(Pharmacia) for 15 min at 37°C. Each sample was then extracted with
phenol/chloroform (50:50 v/v) and precipitated with ethanol. Pellets were
resuspended in sample loading buffer (95% formamide, 20 mM EDTA, 0.05%
bromophenol blue, 0.05% xylene cyanol), and electrophoresed on a 6%

polyacrylamide, 6 M urea gel in 1 x TBE (0.089 M Tris, 0.089 M boric acid, 0.002 M

EDTA).

Preparation of a-TK antiserum,

A truncated form of the human TK protein containing a deletion of the N-
terminal 16 amino acids was expressed in and isolated from E. coli as described
previously (6). To prepare antisera against the protein, approximately 1.5 mg of
partially purified TK was electrophoresed on a preparative 15% SDS-polyacrylamide
gel (SDS—PAGE). One edge of the gel was removed and stained to determine the
position of the TK band. The area corresponding to this band was removed from the
remainder of the gel, homogenized in either complete (primary injection) or
incomplete (secondary boosts) Freund’s adjuvant, and injected subcutaneously into
pathogen free rabbits. Approximately 300 ug of protein was used for the primary
injection and 100 pg for secondary boosts. Two weeks after the secondary injections,
blood was collected from the ear vein, and the a-TK antibody titer was determined.

Serum from the rabbit with the highest titer, termed 43A, was used in the studies

 

 

 

 

43

described here. This antiserum was shown to be specific to TK by the following
criteria: it recognized a 24 kd protein that was present in Rat 3 cells transfected with
the human TK cDNA, but not in untransfected Rat 3 cells. This 24 kd protein was

not recognized by pre-immune serum from the same rabbit (data not shown).

Radiolabeling, immunoprecipitation, and gel electrophoresis of TK protein.

For 35S labeling of cellular protein, confluent monolayers on 100 mm plates
were washed twice with PBS and labeled in 1 ml of methionine free DMEM
containing 10% dialyzed calf serum plus 15-20 ill of EXPRE-label (10 mCi/ml,
Dupont NEN). For 32P labeling, plates were washed twice with Tris-buffered saline
(TBS), then incubated in 1 ml of phosphate free DMEM containing 10% dialyzed
calf serum and 1 mCi 32Pi (ICN). At the end of the labeling period, the medium was
removed, plates were washed twice in cold PBS (355) or TBS (32P), and cytoplasmic
extracts were prepared as described above. For pulse-chase experiments, the labeling
medium was removed, plates were washed twice in PBS, and fresh medium
containing 300 fold excess methionine was added until the cells were harvested. The
protein concentrations of cell extracts were quantitated using the Bradford assay (2).
Total counts incorporated into protein were determined by TCA precipitation.

Labeled TK protein was immunoprecipitated by incubating 100-150 11.1 of
extract with 0.5 ul of 43A antiserum in NP-40 lysis buffer at room temperature for
1 h. At the end of the incubation, 100 pl of protein A-agarose beads (10% v/v) in
NP-40 lysis buffer were added and the incubation was continued at 4°C for 1 h with

rocking. The complexes formed were alternately washed with high salt (2M NaCl, 1%

 

 

 

 

 

44

NP-40, 0.5% deoxycholic acid, 10 mM Tris-HCL pH 7.4), and detergent (0.5% NP-40,
1% SDS in PBS) buffers. After four washes, the pellet was resuspended in 15 pl SDS
gel loading buffer, heated to 95° C for 10 min, and analyzed on 12% SDS-PAGE
minigels (16). 358 gels were soaked in Entensify solutions A and B (DuPont-NEN)
for 30 min. each, then dried under vacuum and fluorographed at -70°C. 32P gels were

dried and autoradiographed at -70°C.

Western blotting.

Total TK protein in cytosolic extracts was purified and concentrated using an
affinity resin containing 43A antibody covalently linked to protein A sepharose. This
affinity resin was prepared as previously described (10). Total cytosolic protein was
incubated with 50 ul of a 10 % (v/v) slurry of coupled 43A antisera in NP-40 lysis
buffer for 1 h at room temperature. At the end of the incubation, the sepharose
beads were pelleted, and the supernatant was removed. Samples were resuspended
in SDS gel loading buffer, heated and electrophoresed on a 12% SDS-PAGE BioRad
mini-gel. After electrophoresis, proteins were transferred to PVDF polyscreen
membrane (Dupont-NEN) at 150 mA for 1 1/2 hours in Western transfer buffer (20
mM Tris, 192 mM glycine, 20% methanol, pH 8.3) (26). The membrane was blocked
overnight at 4°C in 3% nonfat dried milk (NFDM) in PBS with rocking, then at room
temperature for 1 hour. It was then incubated in 1% NFDM containing a 1:500 (v/v)
dilution of 43A antiserum for an additional hour. The membrane was washed five
times in PBS then incubated in fresh 1% NFDM containing 1 uCi [mm-labeled

protein A at 4°C for 12 - 15 hours with rocking. The membrane was then washed in

 

 

 

45

PBS, dried, and exposed to X-ray film.

Quantitation of TK Induction.

To determine the amounts of TK mRNA and protein, an AMBIS Image
Acquisition and Analysis system (AMBIS Inc.) was used to directly detect
radioactivity on gels or membranes. To calculate induction values, the amount of
protein or mRNA at t=0 was defined as 1, and the amounts at all other time points

were expressed relative to that amount.

Fluorescence Activated Cell Sorter Analysis

At various times following serum stimulation, cells were harvested by
trypsinization and fixed by rapid injection into 10 volumes of ice cold 80% ethanol.
2 x 10° cells were removed from each sample, pelleted, washed in PBS and stained
in 1 ml of PI reagent (50 ug/ml propidium iodide, 0.1% Triton X-100, 100 uM
EDTA, 50 ug/ml RNase in PBS, pH 7.4). Stained cells were analyzed on an Ortho

cytoflurograph 50H cell sorter.

 

 

 

46

RESULTS

In previous experiments, we studied the regulation of TK activity during serum
stimulation of Rat 3 cells containing the human TK cDNA expressed from the SV40
early promoter (11). Under these conditions, TK mRNA was induced 2-5 fold within
2 hours of serum stimulation, then remained relatively constant throughout the
remainder of G1 and S. In contrast, TK protein levels were highly induced (up to 50
fold) at the Gl-S transition after serum stimulation. From this data, we concluded
that enzyme levels are regulated by translational and/or post-translational
mechanisms at the G1-S interface. Since the constructs used in our previous
experiments encode hybrid mRNAs containing insertions of SV40 sequences and
deletions of TK 5’ UTR sequences, and because these alterations might affect
translational regulation, we constructed a new hybrid gene, ESVTK (Figure 1A). This
construct contains an intact TK mRNA coding region linked to a minimal, 67 bp
human TK promoter fragment. Experiments in our lab have shown that this promoter
is not regulated by serum in Rat 3 cells (21). Because the 67 bp fragment has low
promoter activity, an SV40 enhancer fragment was included to increase the levels of
expression. To confirm that this promoter initiates transcription properly, Sl nuclease
mapping experiments were performed. Rat 3 cells were stably transfected with
pESVTK, or with the control plasmid pHuTK, and TK+ transfectants were selected

and propagated as pools of 50-100 independent transfectants. pHuTK contains the

 

 

 

 

47

TK cDNA linked to the full length (444 bp) human TK promoter (Figure 1A). Total
RNA was prepared from the transfected cells, and from untransfected Rat 3 controls,
at 14 h after serum stimulation and analyzed by $1 nuclease mapping. The results of
this experiment, shown in Figure 1B, demonstrate that the authentic TK mRNA cap
site is the major transcription initiation site used in cells containing pESVTK. They
also demonstrate that the levels of TK mRNA are higher in cells containing pESVTK
than in those containing pHuTK, presumably due to the presence of the SV40
enhancer in pESVTK.

The levels of TK mRNA, protein, and enzyme activity were examined during
serum stimulation of Rat 3 cells containing the ESVTK gene. Cells were serum
starved and stimulated, and both total cell RNA and cytoplasmic extracts were
prepared at various times after serum addition. At each time point TK mRNA was
analyzed by Northern blot (Figure 2A), TK protein was analyzed by Western blot
(Figure 2B), and TK activity was determined by enzyme assay. To quantitate the TK
mRNA and protein in each sample, the radioactivity in the TK bands on the
Northern and Western blots was measured directly using an Ambis detection and
imaging system. The total counts detected in each band are given in the legend to
Figure 2. The apparent differences between the appearance of the autoradiograms
and the numbers determined by direct scanning are due to the fact that the X-ray
film is not linear over the entire range of counts present in these experiments. To
determine the pattern of induction of TK mRNA and protein, the counts in each

band on a gel were normalized to those present in the t=0 time point. The results

 

 

 

48

Figure 1) Mapping the 5’ ends of TK mRNA in transfected Rat 3 cells. A) Maps of
TK cDNA constructs used. B) Pools of stably transfected Rat 3 cells containing either
pHuTK or pESVTK, and untransfected Rat 3 controls, were grown to confluence,
serum starved and stimulated as described in Materials and Methods. At 14 hours
after serum addition, total cell RNA was prepared and analyzed by $1 nuclease
mapping. Hybrids were digested with 50 (1), 75 (2) or 175 (3) units of $1 nuclease.

The size marker used was a HaeIII digest of pBR322.

 

 

A. 49

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of these calculations, together with the results of TK enzyme assays, are depicted
graphically in Figure 2C. They show a modest, 3-4 fold increase in TK mRNA by 5
h, after which mRNA levels are fairly constant. In contrast, both TK protein and TK
activity increase slightly by 5 h, concomitantly with the mRNA increase, but most
dramatically between 10 and 14 h when the mRNA levels are constant. These
experiments were repeated with a second, independent pool of transfectants with
essentially identical results (data not shown). These results are similar to those
obtained previously (11), and confirm that TK is regulated at translational and/or
post-translational levels in serum stimulated Rat 3 cells.

To determine the stage of the cell cycle at which TK protein levels increase,
the same cells used in Figure 2 were harvested at various times after serum
stimulation, stained with propidium iodide, and analyzed for DNA content in a
fluorescence activated cell sorter (FACS). The results of this experiment, shown in
Figure 3, demonstrate that the cells begin to enter S phase 10-12 hours after serum
stimulation. Thus, the dramatic increase in TK protein levels coincides with passage

through the Gl-S transition.

Translation Efficiengy of TK mRNA During G1 and S-G2. To determine the

relative translation efficiency of TK mRNA during G1 and S, pulse labeling
experiments were conducted in parallel with mRNA determinations. Cells containing
ESVTK were growth arrested, and labeled with 35S-methionine for 1 h from 4-5 (G1),
9-10 (Gl-S) and 13-14 (S-G2) h after serum stimulation. These time points were

chosen since our previous analysis (Figure 2) had shown that TK mRNA levels were

 

 

 

51

Figure 2) Patterns of TK mRNA, protein and enzyme activity during serum
stimulation of cells containing pESVTK. Rat 3 cells containing the ESVTK gene
were serum starved and stimulated as described in Materials and Methods. At
various times after serum stimulation, both total RNA and cytoplasmic extracts were
prepared. A) Northern blot of TK mRNA. 30 ug of RNA were analyzed for each
time point. The counts in each TK band were quantitated directly from the filter
using an AMBIS detection and imaging system. The total counts in each TK band
after a 48 h scan were as follows: 0 h = 2028; 2 h = 4817; 5 h = 6406; 8 h = 7053;
10 h = 9122; 12 h = 6687; 14 h = 7398; 24 h = 4917. B) Western blot of TK
protein. 370 ug of cytosolic protein from each time point were analyzed as described
in Materials and Methods. The amount of label in each TK band was quantitated
directly using the AMBIS system. The lane labeled "TK" contained recombinant TK
protein isolated from E. coli as a marker. The total counts in each TK band after a
28 h scan were as follows: 0 h = 2065; 2 h = 4200; 5 h = 9427; 8 h = 15,416; 10 h
= 13,160; 12 h = 37, 254; 14 h = 67,587; 24 h = 52,332. C) Pattern of induction of
TK mRNA, protein and enzyme activity. TK mRNA levels were determined from the
Northern blot shown in (A) and protein levels from the Western blot shown in (B).
TK enzyme activity was assayed from the same extracts used in the Western blot
shown in (B). In each case, all values were normalized to the level of expression at

t=0 to determine the pattern of induction.

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Figure 3) Cell cycle analysis of serum stimulated cells containing ESVTK. Cells were
grown to confluence, serum starved and stimulated as described in Materials and
Methods. At the indicated time (h) after serum addition, cells were fixed in ethanol,
stained with propidium iodide, and analyzed for DNA content in a fluorescence

activated cell sorter (FACS).

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constant during this interval, while protein levels increased dramatically. At the end
of the labeling period, cells were lysed, and 550 pg of protein from each time point
were immunoprecipitated and analyzed on a 12% SDS-polyacrylamide gel (Figure
4A). In the same experiment, RNA was extracted from cells at the ends of the
protein labeling periods (5, 10 and 14 h), and analyzed by Northern blotting (Figure
4A). The amounts of TK mRNA and labeled TK protein were determined directly
from the dried gel or the blot, and values at 10 and 14 h were normalized to the 5
h time point. The results of this analysis, shown in Figure 4B, indicate that both TK
mRNA levels and the rate of synthesis of TK protein are constant from 5 to 14 hours
after serum stimulation. Thus, the increase in TK protein at the G1-S interface

(Figure 2) is not the result of increased translation of the mRNA.

Determination of TK protein half-life during G1 and S phase.

Pulse-chase experiments were performed to determine the half-life of TK
protein during G1 and S phase. Quiescent cells containing ESVTK were serum-
stimulated as described above, and were then labeled with 3SS-methionine for 60 min
from either 2-3 h (G1) or 12-13 h (S) after serum addition. At the end of the labeling
period, plates were washed twice with PBS and then incubated for various times in
medium containing excess unlabeled methionine. 300 pg of protein from each time
point were immunoprecipitated and analyzed by SDS-PAGE. The results of these
experiments are shown in Fig. 5A and SB. The 3SS-labeled TK at each time point was
quantitated directly from the gel, and these data are plotted in Fig. 5C. The TK

protein half-lives calculated from this experiment were approximately 1.5 hours for

 

 

56

Figure 4: Direct comparison of the translation efficiency of TK mRNA during G1

and S phase. To determine the relative rates of translation of TK mRNA during G1

and S phase, pulse labeling experiments were done in parallel with mRNA
determinations. A) Cells were growth arrested, and labeled with 35S-methionine for

60 min from 4-5 (G1), 9-10 (Gl-S) or 13-14 (S) h after serum addition. 550 pg of
protein from each time point was immunoprecipitated and analyzed by SDS-PAGE.
The times shown above the lanes correspond to the ends of the labeling periods.
Total RNA was prepared at 5, 10 and 14 hours after serum stimulation, and 30 pg
of RNA from each time point was analyzed by Northern blotting. B) The labeled TK
protein and mRNA shown in (A) were quantitated directly from the blot or gel using
an Ambis system. The amount of mRNA/protein at 5 hours was defined as 1, and

the values for the other time points are plotted relative to that value.

Induction

 

57

Protein
mRNA

8 12

Time (hr)

0
V

 

16

 

 

 

 

58

Figure 5: Determination of TK protein half-life during G1 and S-G2. Cells were
growth arrested, serum-starved and stimulated as described in Materials and
Methods, and were labeled with 35S-methionine from 2-3 or 12-13 hours after serum
stimulation. At the end of the labeling period, the media was removed and the
incubation was continued in media containing excess unlabeled methionine for the
times indicated (min). 300 pg of protein from each time point were
immunoprecipitated and analyzed by SDS-PAGE. A) Gel of protein labeled from 2—3
h (G1) after serum addition. B) Gel of protein labeled from 12-13 h (S) after serum
addition. C) The TK bands in the gels shown in (A) and (B) were quantitated
directly using an AMBIS system, and the values are shown in graphic form. v =2-3

h labeling; 0:12-13 h labeling.

 

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the protein synthesized in G1 and 22 hours for the protein synthesized during S
phase. Most of this difference was specific to TK, since the half-lives of total cell
protein were 8.4 h for the 2-3 h labeling and 12.5 h for the 12—13 h labeling. Thus,
the increase in TK protein at the Gl-S interface is the result of an approximate 10

fold increase in TK protein stability as the cells enter S phase.

Phosphorylation of TK Protein. The results presented above indicate that TK
protein is approximately 10 fold more stable during S-G2 phase than during G1. This
difference in stability might be the result of a number of mechanisms. First, there
might be specific cellular molecules that stabilize the protein during S or proteases
that degrade it during G1. Alternatively, the protein might be differentially modified
in G1 and S, and such modifications might alter its stability. To identify modifications
of TK that might affect its stability, we assayed for phosphorylation of the protein at
various times during G1 and S. Cells containing ESVTK were serum-stimulated, and
labeled for 2 hrs with either 35’S-methionine or with 32Pi from 3—5, 6-8, 9-11 or 12-14
hrs after serum addition. The labeled proteins were then immunoprecipitated and
analyzed by SDS-PAGE (Figure 6). Phosphorylated TK was readily detected during
S phase (12-14 hrs) and at the G1—S interface (9-11 hrs), but was barely detectable
in late G1 (6-8 hrs), and not at all in early G1 (3-5 hrs). As shown previously, 358-
labeled protein was readily detected at all time points. Because of the low level of
TK protein present in early G1 cells (demonstrated by Western blot, Figure 2), we
cannot unambiguOusly determine if the low level of 32P-labeled TK at the early time

points is due to underphosphorylation of the protein relative to that seen in S phase.

 

 

61

Figure 6: Phosphorylation of TK protein during G1 and S phase. Cells were serum
starved and stimulated as described in Materials and Methods. At the times indicated
after serum addition, cells containing ESVTK were labeled for two h with either 358-
methionine (S) or 32Pi (P). Cytoplasmic extracts were prepared, and equal cell

equivalents of protein were immunoprecipitated and analyzed by SDS-PAGE.

62

5......
m. 9!.
_
m2:

HP

_

4. s... M:
p!»

mo

6 sp 3
Ph

5

 

TK’

 

 

 

63

Regulation of a TK protein lacking 16 N-terminal amino acids.

We previously examined the expression of a mutant TK protein encoded by
a cDNA construct termed SV-huASmaTK (11), which contains the SV40 early
promoter plus approximately 60 nucleotides of the SV40 5’ UTR linked to an SmaI
site within the TK protein coding sequence (Figure 1A). This construct encodes a
truncated but functional enzyme that is missing 16 N-terminal amino acids. The
pattern of expression of the truncated protein as determined by Western blot is
shown in Figure 7A. Like the wild type, it is expressed at low levels in G0 and G1,
and is highly induced at Gl-S. To determine if the truncated protein is also regulated
at the level of protein stability, we examined its half-life during G1 and S-G2. The
results of this experiment, shown in Figure 7B and 7C, demonstrate that the
truncated protein is more stable during S-G2 than during G1. Thus, the first 16
amino acids of TK coding sequence are not required for the regulation of protein
stability.

We have also examined phosphorylation of the truncated protein using Rat
3 cells containing either SV-huASmaTK or a related construct, SV-huAScaTK. SV-
huAScaTK contains the same SV40 promoter fragment as SV-huASmaTK linked to
a ScaI site in the TK 5’ UTR, and therefore encodes a full length TK protein. Pools
of stably transfected Rat 3 cells containing these constructs were labeled with 32Pi
from 1-3, 12-14, 16-18 or 21-23 h after serum addition, and analyzed by
immunoprecipitation and SDS-PAGE. These experiments, shown in Figure 8A,
demonstrate that the truncated protein, which migrates slightly faster than the wild

type protein, is phosphorylated to a much lesser extent than the wild type. The low

 

 

 

—

64

level of phosphorylation of the truncated protein is not due to a low level of
expression, since it is readily detectable in both 3SS-methionine labelings (Figure 8B)
and in Western blots (Figure 7A). In fact, in direct comparisons this protein seems
to be expressed at higher levels than the wild type. Thus, sequences within the first
16 amino acids of TK are required for the wild type level of phosphorylation to

occur.

65

Figure 7: Regulation of the truncated TK protein encoded by SV-huASmaTK. Rat3
cells containing SV-huASmaTK were serum starved and stimulated as described in
Materials and Methods. A) At the times indicated after serum addition, 370 pg of
protein was analyzed by Western blot. B) Pulse-chase analysis of protein labeled
from 2-3 h (G1) after serum stimulation. 300 ug of protein were immunoprecipitated
at each time point. C) Pulse—chase analysis of protein labeled from 12-13 h (S) after

serum stimulation. 300 ug of protein were immunoprecipitated from each time point.

66

 

 

 

67

Figure 8: 35S-methionine and 32Pi labeling of the truncated protein encoded by SV-
huASmaTK. A) Stably transfected cells containing SV-huASmaTK or SV-huAScaTK
were labeled with 32Pi for 2 h at 1, 12, 16 and 18 h following serum stimulation.
Cytoplasmic extracts were prepared, and equal cell equivalents were
immunoprecipitated and analyzed by SDS-PAGE. B) The pools of cells shown in (A)
were labeled for 20, 40 or 60 min with 35S-methionine at the indicated times after

serum stimulation. Equal cell equivalents of extracts were then immunoprecipitated

and analyzed by SDS-PAGE.

 

68
A Sca Sma
B Sca Sma
2O 40 60 20 40 60 20 40 60 20 40 60
. .3‘ T'”‘ ‘g . .
F! g 29:1
- :— 1.; i .34; 3 2‘.
T.” ' g "' . a- C- .— ..
2hr 10 hr
2hr 10 hr

 

 

69

DISCUSSION

The mechanism of post-transcriptional regulation of TK enzyme levels in
serum stimulated cells has been investigated in stably transfected Rat 3 cell lines. A
comparison of the patterns of enzyme activity and protein expression (Fig. 2C)
indicate that the specific activity of the protein is constant throughout G1 and S
phase. Similarly, the rate of translation of TK mRNA is constant in G1 and S phase
(from 5 to 14 hours after serum stimulation). We were unable to accurately measure
the translation efficiency in G0 and in very early G1, due to the low levels of TK
mRNA in these cells. In contrast to the rate of TK synthesis, the rate of turnover of
the protein is highly regulated, with newly synthesized TK protein being
approximately 10 times more stable during S-G2 than during G1.

These results differ from those reported in cycling cells, in which TK mRNA
was reportedly translated 10 times more efficiently during S and G2 than during Gl
(14,22). The fact that we do not detect translational regulation in our experiments
is not likely to be an artifact of the construct that we used, since ESVTK initiates
transcription at the authentic TK cap site (Figure 1). It is also unlikely to be an
artifact of studying the human gene in transfected rat cells, since the previous
experiments in cycling cells showed identical regulation of the human gene in Hela
cells and in transfected mouse L cells (14,22). It is possible that regulation is

different in serum stimulated and cycling cells, and that the newly synthesized mRNA

 

 

70

produced in serum stimulated cells is not subject to translational regulation.
Alternatively, there might be a difference between normal (Rat3) and transformed
(Hela and mouse L) cells. A second difference between our results and those
reported in cycling cells is the timing of the change in TK protein stability. In cycling
Hela cells, TK protein was unstable during mitosis, but was as stable during most of
G1 as during S and G2 (22). In contrast, our experiments indicate that TK protein
is unstable during most of G1, and that a major increase in protein stability occurs
at or about the Gl-S interface. Our results are consistent with a previous report that
regulation of TK protein levels during serum stimulation is totally abolished by
mutations which disrupt the C-terminus of the TK protein and result in its being
stable throughout the cell cycle (13). If a change in the translation rate of TK mRNA
was contributing significantly to the regulation of protein levels, one might expect
that these C-terminal mutations would only partially disrupt the pattern of expression.
The alternative explanation, that the C-terminal mutations affect both translational
and post-translational regulation, is also possible but seems less likely.

The experiments presented here also demonstrate that TK is phosphorylated,
at least during late G1 and S phase. This confirms a recent report in which
phosphorylation of TK was reported in HL—60 cells (4). Phosphoamino acid analysis
of the protein from HL-60 cells indicated that phosphorylation was exclusively on
serine residues, and tryptic peptide mapping experiments demonstrated 2 major and
several minor labeled peptides. The fact that the truncated protein encoded by SV-
huASmaTK is underphosphorylated relative to the wild type suggests that some of the

phosphorylation sites are located within the region deleted in this protein. The

 

—

71

predicted amino acid sequence of TK (3) shows that the first trypsin cleavage site is
a lysine at amino acid 16. The ASma mutant deletes up to this lysine, and then
replaces the following thr-arg with met-gly. We would therefore predict that this
mutant lacks only one tryptic peptide, which contains three serines that are potential
sites of phosphorylation. The ASma mutation may also alter phosphorylation of other
sites due to conformational changes of the protein. Although we have not rigorously
compared the specific activity of the wild type and truncated proteins, cells containing
SV-huASmaTK consistently have equal or higher levels of TK protein and activity
than those containing SV-huAScaTK or ESVTK, so neither the first 16 amino acids
of the protein nor the wild type pattern of phosphorylation is critical for activity.

The fact that TK is phosphorylated suggested the intriguing possibility that its
stability might be regulated by phosphorylation, and that it might be the target of a
Gl-S specific kinase. From our experiments, we cannot unambiguously determine
if TK is underphosphorylated during G1 relative to S-G2, since the level of protein
is very low in G1. The fact that the truncated protein encoded by SV-huASma'IK is
underphosphorylated, but is regulated at the level of protein stability, indicates that
not all of the phosphorylation sites are required for regulation. It is possible,
however, that a subset of phosphorylation sites are maintained in the truncated
protein, and that these sites are involved in regulation.

In summary, we have established that the increase in TK enzyme levels at G1-
8 in serum stimulated cells containing high levels of TK mRNA is due exclusively to
a change in the stability of the protein, and that this regulation is independent of

sequences at the amino terminus of the protein. Several possible mechanisms might

 

 

—

72

account for this change in stability. First, the protein might be differentially modified,
and this might affect its half-life. Alternatively, the change in stability might be due
to other molecules, such as proteases, whose activity changes between G1 and S. The
fact that TK protein stability changes at or near the Gl-S interface suggests that the
protein itself is a target for cellular molecules that act at this important control point
in the cell cycle. Current experiments are directed towards identifying these

molecules and analyzing their role in the regulation of protein stability.

ACKNOWLEDGEMENTS
The authors would like to thank Mr. Houman Dehghani and Ms. Kyung Eun
Kim for assistance in constructing pESVTK, and Drs. D. DeWitt, J. Wang and J.
Dodgson for helpful comments on the manuscript. This work was supported in part
by grant NIH-CA37144, by BRSG funds from the College of Veterinary Medicine,

and by State of Michigan Research Excellence Funds.

 

 

 

73

REFERENCES

1) Boyle, D., B. Coupar, A Gibbs, L. Seigman, and G. Both 1987. Fowlpox virus
thymidine kinase: Nuclear sequences and relationships to other thymidine kinases.
Virology 156: 355-365.

2) Bradford, M.M. 1976. A rapid and sensitive method for the quantitation of
microgram quantities of protein utilizing the principle of protein-dye binding. Anal.
Biochem. 72: 248-254.

3) Bradshaw, H.D., and PL. Deininger. 1984. Human thymidine kinase gene:

Molecular cloning and nucleotide sequence of a cDNA expressible in mammalian

cells. Mol. Cell. Biol. 4: 2316-2320.
4) Chang, Z.F., and D.Y. Huang. 1993. The regulation of thymidine kinase in HL—60
human promyeloleukemia cells. J. Biol. Chem. 268: 1266-1271.

5) Coppock, D., and AB. Pardee. 1987. Control of thymidine kinase mRNA during
the cell cycle. Mol. Cell. Biol. 7: 2925-2932.

6) Dagher, S.F., S.E. Conrad, E. Werner, and R.J. Patterson. 1992. Phenotypic
conversion of TK-deficient cells following electroporation of functional TK enzyme.
Exp. Cell Res. 198: 36-42.

7) Don, Q-P., P.J. Markell, and AB. Pardee. Thymidine kinase transcription is
regulated at G1 / S by a complex that contains retinoblastoma-like protein and a cdc2

kinase. Proc. Natl. Acad. Sci. USA 89: 3256-3260.

 

 

74

8) Gross, M.K., and G.F. Merrill. 1988. Regulation of thymidine kinase protein levels
during myogenic withdrawal from the cell cycle is independent of mRNA regulation.
Nucleic Acids Res. 16: 11625-11643.

9) Gross, M.K., and G.F. Merrill. 1989. Thymidine kinase synthesis is repressed in
nonreplicating muscle cells by a translational mechanism that does not affect the
polysomal distribution of thymidine kinase mRNA. Proc. Natl. Acad. Sci. USA 86:
4987-4991

10) Harlow, E., and D. Lane. 1988. Antibodies, a laboratory manual. Cold Spring
Harbor Press, Cold Spring Harbor, NY.

11) Ito, M., and SE. Conrad. 1990. Independent regulation of thymidine kinase
mRNA and enzyme levels in serum stimulated cells. J. Biol. Chem. 265: 6954-6960.
12) Johnson, L., L.G. Rao, and A. Muench. 1982. Regulation of thymidine kinase
enzyme levels in serum-stimulated mouse 3T6 fibroblasts. Exp. Cell Res., 138: 79-85.
13) Kauffman M., P. Rose, and T. Kelly. 1991. Mutations in the thymidine kinase
gene that allow expression of the enzyme in quiescent G0 cells. Oncogene 6: 1427-
1435.

14) Kauffman M., and T. Kelly. 1991. Cell cycle regulation of thymidine kinase:
Residues near the carboxyl terminus are essential for the specific degradation of
enzyme at mitosis. Mol. Cell. Biol. 11: 2538-2546.

15 ) Kim, Y.K., S. Wells, Y.F. Lau, and AS Lee. 1988. Sequences contained within the
promoter of the human thymidine kinase gene can direct cell—cycle regulation of
heterologous fusion genes. Proc. Natl. Acad. Sci. USA. 85: 5894-5898.

16) Laemmli, U.K. 1970. Cleavage of structural proteins during the assembly of the

 

 

 

 

75
head of bacteriophage T4. Nature 227: 680-685.

17) Li, L-J., G.S. Naeve, and A.S. Lee. 1993. Temporal regulation of cyclin A-p107
and p33°dk2 complexes binding to a human thymidine kinase promoter element
important for Gl-S phase transcriptional regulation. Proc. Natl. Acad. Sci, USA 90:
3554—3558.

18) Machovich, R., and O. Greengard. 1972. Thymidine kinase in rat tissues during
growth and differentiation. Biochim. Biophys. Acta 286: 375-381.

19) Merrill, G.F., S. Hauschka, and S. McKnight. TK enzyme expression in
differentiating muscle cells is regulated through an internal segment of the cellular
TK gene. Mol. Cell. Biol. 4: 1777-1784.

20) Nevins, JR. 1992. E2F: A link between the Rb tumor suppressor protein and
viral oncoproteins. Science 258: 424-429.

21) Roehl, H., and SE. Conrad. 1990. Identification of a G1/S phase inducible
region in the human thymidine kinase gene promoter. Mol. Cell. Biol. 10: 3834-3837.
22) Sherley, J.L., and T. Kelly. 1988. Regulation of human thymidine kinase during
the cell cycle. J. Biol. Chem. 263: 8350-8358.

23) Stewart, C.J., M. Ito, and SE. Conrad. 1987. Evidence for transcriptional and
post-transcriptional control of the cellular thymidine kinase gene. Mol. Cell. Biol.
7: 1156-1163.

24) Stuart, P., M. Ito, C.J. Stewart, and SE. Conrad. 1985. Induction of cellular
thymidine kinase occurs at the mRNA level. Mol. Cell Biol. 5: 1490-1497.

25) Topp, WC. 1981. Normal rat cell lines deficient in nuclear thymidine kinase.

Virology 1 13:408-411.

 

 

75
head of bacteriophage T4. Nature 227: 680-685.

17) Li, L-J., G.S. Naeve, and A.S. Lee. 1993. Temporal regulation of cyclin A-p107
and p33°dk2 complexes binding to a human thymidine kinase promoter element
important for Gl-S phase transcriptional regulation. Proc. Natl. Acad. Sci, USA 90:
3554-3558.

18) Machovich, R., and O. Greengard. 1972. Thymidine kinase in rat tissues during
growth and differentiation. Biochim. Biophys. Acta 286: 375-381.

19) Merrill, G.F., S. Hauschka, and S. McKnight. TK enzyme expression in
differentiating muscle cells is regulated through an internal segment of the cellular
TK gene. Mol. Cell. Biol. 4: 1777-1784.

20) Nevins, JR. 1992. E2F: A link between the Rb tumor suppressor protein and
viral oncoproteins. Science 258: 424-429.

21) Roehl, H., and S.E. Conrad. 1990. Identification of a Gl/S phase inducible
region in the human thymidine kinase gene promoter. Mol. Cell. Biol. 10: 3834-3837.
22) Sherley, J.L., and T. Kelly. 1988. Regulation of human thymidine kinase during
the cell cycle. J. Biol. Chem. 263: 8350-8358.

23) Stewart, C.J., M. Ito, and S.E. Conrad. 1987. Evidence for transcriptional and
post-transcriptional control of the cellular thymidine kinase gene. Mol. Cell. Biol.
7: 1156-1163.

24) Stuart, P., M. Ito, C.J. Stewart, and S.E. Conrad. 1985. Induction of cellular
thymidine kinase occurs at the mRNA level. Mol. Cell Biol. 5: 1490-1497.

25) Topp, W.C. 1981. Normal rat cell lines deficient in nuclear thymidine kinase.

Virology 113:408-411.

 

 

 

*

76
26) Towbin, H., R. Staehelin, and J. Gordon. 1979. Electrophoretic transfer of

proteins from polyacrylamide gels to nitrocellulose sheets: Procedure and some
applications. Proc. Natl. Acad. Sci. USA 76: 4350-4354.

27) Travali S., K. Lipson, D. J askulski, E. Lauret, and R. Baserga. 1988 Role of the
promoter in the regulation of the thymidine kinase gene. Mol. Cell. Biol. 8: 1551-
1557

28) Wigler, M., A. Pellicer, S. Silverstein, and R. Axel. 1978. Biochemical transfer of

single copy eucaryotic genes using total cellular DNA as a donor. Cell 14: 725-731

 

 

 

 

Chapter III

SUBCELLULAR LOCALIZATION OF HUMAN THYMIDINE KINASE

IN SERUM STIMULATED CELLS

By

Marc A. Carozza and Susan E. Conrad

To be submitted to Experimental Cell Research

77

 

 

 

 

The experiments described in the present study were designed to determine

the subcellular localization of the human TK polypeptide in vivo, and to resolve
whether this localization changes in a cell cycle specific manner. Utilizing indirect
irnmunofluorescence and confocal microscopy we demonstrated that TK was found
exclusively in the cytoplasm, and its subcellular localization did not change between

G1 and S phase, of the cell cycle.

 

 

79

INTRODUCTION

Thymidine kinase is an enzyme in the pyrimidine salvage pathway which catalyses
the ATP dependent phosphorylation of thymidine to thymidine 5’ monophosphate,
which upon further phosphorylation is ultimately incorporated into DNA. Two forms
of the enzyme are present within cells, a mitochondrial form and a cytosolic form
(1,2). The activity of the cytosolic form is controlled in a cell cycle specific manner,
with a sharp increase in enzyme activity paralleling the onset of DNA synthesis (3,4).
Cell cycle regulation of thymidine kinase occurs at both the transcriptional and post-
transcriptional levels (5-10). In chapter 2 of this thesis we investigated the post-
transcriptional regulation of the TK polypeptide in serum stimulated cells. The
results of these studies revealed that the TK protein was regulated at the level of
protein half life, with its stability increasing 10 fold as the cells entered S phase. We
also demonstrated that TK is phosphorylated, and identified a mutant protein that
is drastically underphosphorylated.

There are conflicting views on the subcellular localization of TK during the
cell cycle. Some reports found activity in the nucleus and cytoplasm (11), while
others have found activity exclusively in the cytoplasm (10,12). The conclusions
reached in these studies were based on enzymatic activity in fractionated cells, which
may be influenced by different extraction procedures. To date the subcellular

localization of human TK has not been investigated in vivo. Kit et al (13) reported

 

 

 

___________—_—I

80

that in fractionated extracts from HeLa cells, 95% of the TK activity was found in
the cytsolic fraction, 2-3% was localized to the mitochondrial fraction, with the
remaining activity being in the microsomal and nuclear fractions. Sherley and Kelly
(10,12) were the first to purify the cytoplasmic form of TK. They reported that this
form of the enzyme represented approximately 98% of the activity recovered from
HeLa cellular extracts, and that the localization did not change during the cell cycle

A number of early studies proposed that distal precursors of DNA synthesis
were channeled into the DNA replication machinery by multienzyme complexes, thus
maintaining a higher concentration of dNTPs at the site of replication than would be
built up by freely diffusing enzymes (11,14,15). The maintenance of concentration
gradients of metabolites by the activity of multienzyme complexes is referred to as
functional compartmentation (16). Functional compartmentation has been reported
in the replication of T4 phage (17,18), Chinese hamster embryo cells (11), and in
other mammalian cells (15,19-23). These multienzyme complexes were reported to
contain thymidine kinase, DNA polymerase, thymidylate synthase, and dihydrofolate
reductase. Utilizing individual enzyme assays of fractionated cellular components
Reddy (11) reported that the subcellular localization of these enzymes shifted from
a cytoplasmic to a nuclear location as the cells progressed from G1 into S phase.
Specifically, approximately 13% of the enzyme activities were found in the nuclear
fraction of quiescent G0 phase cells, while an average of 50 % of the activities were
found in this fraction during S phase. The nuclear localization of thymidine kinase
was reported to increase from approximately 8% during G0 to 40% during S (11).

It is not clear why these studies obtained contradictory results. One possible

 

 

a

81

variable that should be considered are the methods by which the location of these
enzymes, including thymidine kinase was determined. Specifically, all of the
publications based their findings on enzymatic assays of the various components of
fractionated cells to determine the location of enzymes within cells. The results
obtained by this method of analysis can be influenced by the extraction and
separation procedures used to fractionate cells. In addition, they may not allow the
localization of an enzyme or enzyme complex to a certain region within a
compartment. In the present study we determined the in vivo localization of human
cytoplasmic thymidine kinase during serum stimulation of cells utilizing indirect
immunofluorescence. Our experiments indicate that as the cell progresses from G1
to S, TK is found exclusively in the cytoplasm. Its subcellular localization does not
change during this period, and the major phosphorylation sites within the protein do

not appear to influence localization.

 

 

 

82

MATERIALS AND METHODS

Cell Culture:

Rat 3 TK‘ cells were grown in Dulbecco’s modified Eagle medium (DMEM)
supplemented with 10% Hyclone calf serum. Rat 3 cells stably transfected with the
recombinant plasmids ESVTK and SV-huASma TK (described in chapter 2) were
grown in the same medium supplemented with 1 x HAT (Hypoxanthine,
Aminopterin, Thymidine) medium (Gibco). Cells utilized in indirect
immunofluorescence experiments were grown on glass coverslips until they were
approximately 70% confluent. Synchronous populations of cells were obtained by
serum starving monolayers for 30 hours in DMEM containing .1% calf serum, and
then serum stimulating by replacing the media with DMEM containing 10 % calf
serum. At various times post serum stimulation, coverslips were removed and

processed for indirect immunofluorescence.

Indirect Immunofluorescence:

Coverslips were removed from growth media, washed twice in PBS, then fixed
for 15 minutes in PBS containing 2% formaldehyde at room temperature. Fixed
coverslips were washed once in PBS and then in PBS containing 10% calf serum,
and were then incubated inverted for 1 hour in 100 pl of the primary antisera 43A
(described previously) diluted 1:40 in PBS containing 10% calf serum and .2%

saponin at room temperature on parafilm. Coverslips were then washed twice in

 

 

_——.l

83

PBS / 10% calf serum and incubated for an additional hour in rhodamine conjugated
goat antirabbit antiserum diluted 1:1000 in PBS/10% calf serum/ .2 % saponin.
Coverslips were then washed once each in PBS/10% calf serum, PBS, and then
distilled water. For microscopic observation, processed cover slips were mounted
inverted on glass slides in a drop of water. Indirect immunofluorescence and
photomicroscopy were performed using a Zeiss Axiophot microscope. Confocal

microscopy was performed using a Zeiss laser scanning confocal microscope.

 

84

RESULTS

In chapter 2 we demonstrated that as serum stimulated cells progress from G0
through the G1 /S interface, TK mRNA levels and the rate of TK protein synthesis
remained constant while the half life of the TK polypeptide increased 10 fold. We
concluded from these studies that the observed increase in TK protein levels at the
G1 / S interface was due to an increase in the stability of the protein and not a result
of an increase in translation. One possible mechanism to explain this observation is
that the subcellular localization of TK changes in a cell cycle specific manner, and
that this change stabilizes the TK protein.

To determine the subcellular localization of human TK in vivo, and to resolve
whether this localization changes between G1 and S phase, we utilized indirect
immunofluorescence to visualize the TK polypeptide in synchronized serum
stimulated cells. Previous studies had determined that these cells enter S phase at 10-
12 hours after serum stimulation (figure 3, chapter 2). We therefore chose time
points to span from the middle of G1 to early S phase. The results of these studies
are shown in figure 1. To verify the fluorescence observed was not due to
nonspecific binding of the antibodies used in the study, Rat 3 TK' cells were
processed along with the ESVTK cells and the 14 hour (S phase) time point is shown
in figure 1F. As expected the level of TK was very low during G1 and as the cells

progressed from G1 into S phase, the amount of fluorescence increased dramatically,

 

85
and the localization of the TK protein did not appear to change (figure 1A-E).

Based on the data shown in figure 1, TK appears to be localized around the
nucleus, but is not found within the nucleus (figure 1D,1E). Due to the physical
structure of adherent cells however, the increased intensity observed around the
nucleus could be caused by the increase in cytoplasmic volume in this area. The
image that we see would therefore be due to an additive affect of the light passing
through this region. Also we could not unequivocally determine if TK was associated
with any membranous structures within the cell, including the nuclear envelope. To
resolve these questions, we utilized laser scanning confocal microscopy to better
visualize the localization of TK immunofluorescence. Utilizing a computer aided
optical display, we observed a confocal image of the immunofluorescence from both
the top of the cell (figure 2A,2B), and from a cross section of the labeled TK cells
(figures 2C,2D). The results of this study revealed that the apparent perinuclear
localization observed in the previous experiment was in fact due to the increase in
cytoplasmic volume around the nucleus. Note that in top view of the cell the
intensity of the fluorescence does not change from the nuclear envelope out to the
periphery of the cell (figure 2A,2B), this observation was reinforced by the cross
sectional view where the same distribution was observed. Also by scanning from the
top of the nucleus to the attached basal membrane it was shown that TK is not

contained within or associated with the nucleus (data not shown).

 

 

 

86

Figure 1) Effect of cell cycle progression on subcellular location of TK. At various
times post-serum stimulation, coverslips were harvested and processed for
immunofluorescence analysis as described in materials and methods. The times
shown are; (A) 5hr (G1), (B) 8hr (G1), (C) 10hr (Gl/S), (D) 12hr (S), and (E) 14hr
(S). A control slide containing Rat 3 cells was harvested at 14 hrs post serum

stimulation and processed along with TK containing cells (1F)

87

 

 

 

 

 

88

Figure 2) Confocal microscopy of labeled cells containing ESVTK 14 hrs post-serum
stimulation. A and B), Top view of cells imaged using confocal microscopy to focus

near the base of the cells. C and D), Cross sectional view of cells imaged in A and

B.

89

 

 

90

DISCUSSION

The subcellular localization of the human thymidine kinase protein has been
investigated to determine the location of the protein in vivo. We also determined if
this localization changed as the cells progressed from G1 to S phase in the cell cycle.
The results of these studies revealed that the TK protein is found exclusively in the
cytoplasm, and is evenly distributed throughout the cell (figure 2). It was also
demonstrated that the localization of the TK polypeptide does not change during the
cell cycle (figure 1A-E). These findings confirm studies in which all the TK activity
was associated with the cytoplasmic fractions (10,12). It disproves the studies that
found significant amounts of activity in the nucleus during S phase (11).

Also due to the specificity of our TK antibody, this procedure would be a

useful tool in measuring TK expression in proliferating cells

ACKNOWLEDGEMENTS
The authors wish to thank Marty Regier for the gift of her
immunofluorescence labeling procedure, Mark Shieh for his technical assistance on

the Ziess Axiophot, and Dr. J oAnn Whallon for conducting the confocal microscopy.

 

91

REFERENCES

1) Berk, A.J., and D. Clayton (1973) J. Biol. Chem. 248: 2722-2729

2) Kit, S., W.C. Ieung, and LA. Kaplan (1973) Eur. J. Biochem. 39: 43-48

3) Johnson, L., L.G. Rao, and A. Muench (1982) Exp. Cell Res., 138: 79-85

4) Stuart, P., M. Ito, C.J. Stewart, and S.E. Conrad (1985) Mol. Cell Biol. 5: 1490-
1497

5) Travali S., K. Lipson, D. Jaskulski, E. Lauret, and R. Baserga (1988) Mol. Cell.
Biol. 8: 1551-1557

6) Stewart, C.J., M. Ito, and S.E. Conrad (1987) Mol. Cell. Biol. 7: 1156-1163

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