Nucleic Acids Research, Vol. 18, No. 16 4867
Further biochemical characterization of wheat DNA
primase: possible functional implication of copurification
with DNA polymerase A
Patricia Laquel, Michel Castroviejo and Simon Litvak*
Laboratoire de Biologie Moleculaire Vegetale, IBCN-CNRS, 1 rue Camille Saint Saens, 33077
Bordeaux cedex, France
Received April 10, 1990; Revised and Accepted July 27, 1990
ABSTRACT
DNA primase has been partially purified from wheat
germ. This enzyme, like DNA primases characterized
from many procaryotic and eucaryotic sources,
catalyses the synthesis of primers involved in DNA
replication. However, the wheat enzyme differs from
animal DNA primase in that it is found partially
associated with a DNA polymerase which differs greatly
from DNA polymerase a. Moreover, the only wheat DNA
polymerase able to initiate on a natural or synthetic
RNA primer is DNA polymerase A. In this report we
describe in greater detail the chromatographic
behaviour of wheat DNA primase and its copurification
with DNA polymerase A. Some biochemical properties
of wheat DNA primase such as pH optimum, Mn + 2 or
Mg + 2 optima, and temperature optimum have been
determined. The enzyme is strongly inhibited by KCI,
cordycepine triphosphate and dATP, and to a lesser
extent by cAMP and formycine triphosphate. The
primase product reaction is resistant to DNAse
digestion and sensitive to RNAse digestion. Primase
catalyses primer synthesis on M13 ssDNA as template
allowing E.coli DNA polymerase I to replicate the
primed M13 single-stranded DNA leading to doublestranded M13 DNA (RF). M13 replication experiments
were performed with wheat DNA polymerases A, B, Cl
and CM purified in our laboratory. Only DNA polymerase
A is able to recognize RNA-primed M13 ssDNA.
INTRODUCTION
Purified DNA dependent-DNA polymerases are unable to
catalyse the de novo initiation of DNA synthesis. The initiation
reaction is provided by the classical DNA-dependent RNA
polymerases or by enzymes called DNA primases that synthesize
short RNA primers; the latter can be elongated by the replicative
DNA polymerase (1,2). DNA primase activity has been purified
from a variety of eucaryotic organisms (3 — 16). In animal cells
DNA primase is strongly associated with the replicative DNA
polymerase a; considerable effort has been devoted to separating
both enzymes and characterizing the primase activity (17-19);
for a recent review see Kaguni and Lehman (20). DNA primase
differs from the usual DNA-dependent RNA polymerases in that
*To whom correspondence should be addressed
the primase has a lower molecular weight (10, 21) and a higher
resistance to certain transcription inhibitors such as rifampicin
and a-amanitin. Moreover, DNA primases are able to incorporate
dNTP to a limited extent in addition to rNTP, while nuclear RNA
polymerases are strictly specific for rNTP.
In our laboratory we have characterized several DNA
polymerases from wheat germ (22 — 31,37). DNA polymerase
CII has many of the properties of animal DNA polymerase a
(22—24), while DNA polymerase B can be considered as a
5-like DNA polymerase (Richard,M.C. et al, manuscript in
preparation) . Wheat DNA polymerase CI resembles animal 0
polymerase (26) and DNA polymerase A resembles animal DNA
polymerase y, although the wheat enzyme is not confined to the
mitochondrial compartment (27—29).
We have previously shown that partially purified preparations
from wheat embryos germinated for four hours are able to
catalyse the synthesis of short RNA primers (30). A fraction of
the wheat primase activity copurifies with a wheat germ DNA
polymerase very similar to DNA polymerase A (30,31). There
is a surprising difference with animal cells, where the primase
activity is strongly and specifically associated with DNA
polymerase a.
This report describes a further biochemical characterization
of the wheat DNA primase. Special emphasis has been given
to the chromatographic behaviour of this enzyme and to extensive
copurification with DNA polymerase A. The analysis of the
product synthesized in the presence of M13 ssDNA as template,
the effect of several inhibitory agents and the specific role that
DNA polymerase A seems to play on wheat DNA primase
synthesized RNA oligonucleotides are presented and discussed.
MATERIAL AND METHODS
Wheat embryos were prepared from the variety Marius (Brosse
Monceaux Agronomical Center, 77 130 Monceaux France).
Commercial wheat germ was a kind gift from 'Les Grands
Moulins de Paris' (usine de Bordeaux). Polynucleotides and
oligonucleotides were from Sigma. Chem. Co, BoehringerMannheim and PL. Pharmacia. Unlabeled nucleotides were
obtained from Sigma or P.L. Biochemicals. Labeled precursors
[a32P] ATP, [a 32 P] UTP, [a32P] dATP, [3H] TTP, [3H] dATP
were from Amersham and CEA Saclay. Calf thymus DNA was
4868 Nucleic Acids Research, Vol. 18, No. 16
purchased from Sigma. M13 single-stranded DNA (mp 9, strand
+) was obtained from BRL and the M13 universal primer from
Appligene-Strasbourg. Sequencing reaction kits were from
Pharmacia. Pancreatic RNase and DNase I were from Sigma and
E. coli DNA polymerase I and RNA polymerase were from
Boehringer-Mannheim. Trypsin and trypsin inhibitor were from
Sigma. Phenyl-methane-sulphonyl-fluoride (PMSF) and
proteinase K were from Boehringer-Mannheim. Dextran T40 was
from Pharmacia. Polyethylene glycol (type 400) was from Sigma
and Triton X-100 from Merck. Ficoll type 400 and polyvinylpyrrolidone (PVP-360) were from Sigma. RNasin was purchased
from Genofit. DEAE cellulose DE 52 and Phosphocellulose PI 1
were from Whatmann Inc. DNA-ceUulose (double-stranded DNA
from calf thymus) was from Sigma. Hydroxyl apatite ultrogel
was purchased from BioRad. Heparine-sepharose CL 6B was
from Pharmacia.
Enzyme purification
Wheat primase purification. The extract was prepared from 360 g
of commercial wheat germ as already described (22—26), except
for the presence of 1% Dextran T40, 1% Ficoll T400, 1%
polyvinylpyrrolidone (PVP-360) in the grinding buffer. These
molecules are known to neutralize phenols present in plant
extracts that could interfere with the enzymes. Proteins from a
wheat extract were precipitated between 20% and 70%
ammonium sulphate saturation, dialyzed against buffer A (50 mM
Tris HC1 pH 7.9; 1 mM 2-mercaptoethanol; 0.1 mM EDTA;
20% glycerol and 0.1 mM PMSF), plus 0.1 M KC1, then loaded
on a first phosphocellulose column (800ml) pre-equilibrated in
buffer A plus 0.1 M KC1. The bulk of the DNA polymerases
were retained by this resin. DNA primase activity was found
in two fractions: the first was not retained in the phosphocellulose
column and was detected in the flow through of this resin, the
second was retained together with the DNA polymerase activities
and was studied further (see diagram of purification). Proteins
were eluted from the phosphocellulose column with 0.8 M KC1
in buffer A, dialyzed against buffer B (50 mM Tris-HCl pH 7.5;
1 mM 2-mercaptoethanol; 0.1 mM EDTA; 20% glycerol and
0.1 mM PMSF), and loaded on a 200 ml DEAE cellulose column
equilibrated in buffer B. Proteins were eluted stepwise at 0.3 M
KC1, diluted with buffer A to decrease the KC1 concentration
to 0.2 M KC1, and then loaded immediately on a second
phosphocellulose column (20ml) equilibrated in buffer A plus
0.2 M KC1. Primase activity was again divided in two fractions.
The fraction of primase activity not retained on this second
phosphocellulose was directly loaded on a 10 ml DEAE-cellulose
column equilibrated in buffer B plus 0.1 M KC1. After washing,
the proteins retained were eluted by a 100 ml linear gradient of
0.1 - 0 . 6 M KC1 in buffer B. The pool of DNA primase activity
was dialyzed against buffer B and loaded on a 5 ml heparine
sepharose column equilibrated in buffer B plus 0.1 M KC1; the
primase was eluted from this column by a linear gradient of
0.1 —0.6 M KC1 prepared in buffer B. Active primase fractions
were pooled and dialyzed against buffer B plus 50% glycerol.
The enzyme was kept in 50% glycerol for several months without
apparent loss of activity.
Purification of animal DNA polymerases a and y. Purification
of DNA polymerases a and 7 from Xenopus laevis oocytes have
been described previously (32).
DNA polymerase assays
The incubation mixtures for the DNA polymerase assay contained
the following common reagents in a final volume of 50 /tl: 50
mM Tris-HCl pH 8.0; 5 mM MgCl2; 10 mM DTT; 1 to 5 mg
of proteins and a) 20 /tg/ml of activated DNA as template plus
100 /tM dATP, dCTP, dGTP and 10 /tM [3H] TTP (500-1500
cpm /pmol); b) 0.48 A260 /ml of the synthetic template-primer,
poly rA-oligo dT, in a ratio 5:1; 100 mM KC1, plus 10 /tM of
[3H] TTP; c) 0.48 A260/ml of the synthetic template primer poly
dT-oligo rA, in a ratio 5:1, plus 10 /tM [3H] dATP. Calf
thymus DNA was activated with pancreatic DNase as described
by Aposhian and Kornberg (33). Template primer annealing was
performed by heating and slow cooling was as already described
(22-25). Incubations were carried out for 30 minutes at 37°C.
Reactions were stopped by the addition of 1 ml of ice cold 10%
trichloracetic acid plus 0.1 M sodium pyrophosphate. The
precipitate was filtered on nitrocellulose membranes, dried and
the radioactivity counted in a PPO-POPOP-toluene scintillation
mixture.
DNA primase assay
The incubation mixture for the indirect assay of primase activity
contained in a final volume of 50 /tl: 50 mM Tris-HCl pH 8.0;
5 mM MgCl2; 10 mM DTT; 1 to 5 /tg of protein, 0.48 A 260
poly dT as template and 1 mM ATP as substrate. Incubation was
for 10 minutes at 37°C in the presence of primase. Then 0.5
units of E. coli DNA polymerase I (or 5 mg of DNA polymerase
A when indicated) plus 10 mM [3H] dATP were added in the
reaction mixture. The incubation was continued for 50 minutes
at 37°C. The reaction mixture was directly precipitated with 1
ml cold trichloracetic acid plus 0.1 M pyrophosphate, and the
radioactivity was counted as described in DNA polymerase assay.
Replicative synthesis of double-stranded (ds) M13 DNA
The incubation mixture contained the following reagents in a
100 /tl volume: 50 mM Tris-HCl pH 8.0; 2.5 mM MgCl2; 10
mM DTT; 1 mM ATP; 250 /tM of each of CTP, GTP and UTP;
1 /tg of purified primase (heparine-sepharose fraction devoid of
contaminating nuclease); 1 /tg of single-stranded (ss) M13 DNA
(mp 9, strand +). Incubation for primer synthesis was performed
at 37°C for 30 minutes. Replication of primed single-stranded
M13 DNA into double-stranded DNA was performed by addition
of the following: 250 /tM of each of dCTP, dGTP and TTP;
5 /tCi of [«32P] dATP and 0.5 unit of E. coli DNA polymerase
I. The reaction was stopped with 10 mM EDTA and extracted
twice with phenol, chloroform, and nucleic acids were
precipitated in the presence of 96% ethanol at -20°C. After
centrifugation the nucleic acid pellet was resuspended in 10 /tl
of loading buffer (0.25% bromophenol blue, 40% (w/v) sucrose
in sterile water). The reaction products were analyzed by
electrophoresis on non-denaturing 0.8% agarose gels prepared
in 40 mM Tris-acetate pH 8.0, 0.11% acetic acid and 1 mM
EDTA. Gels were run at constant voltage (100 V ). After
electrophoresis, gels were dried and autoradiographed using
KODAK X-OMAT films.
Synthesis of labeled primer RNA
The reaction was performed in a final volume of 50 /tl and
contained the following: 50 mM Tris-HCl pH 8.0; 2.5 mM
Nucleic Acids Research, Vol. 18, No. 16 4869
MgCl2; 10 mM DTT; 25 /ig/ml M13 single-stranded DNA ( mp
9, strand +); 500 units/ml of RNasin; 1 mM of each of CTP,
GTP, 5 ^Ci [a32P] UTP and ATP (800 Ci /mmol), and 6 mg
of wheat DNA primase. Incubation was performed at 37°C for
45 minutes and the reaction mixture was submitted as indicated
to: a) digestion for 15 minutes at 37°C in the presence of 2.5 /tg
of DNase I (RNase free); b) digestion for 15 minutes at 37°C
in the presence of 2.0 tig of pancreatic RNase; c) or was
immediately stopped without preliminary treatment (control) by
the addition of 0.03% SDS; 5 mM EDTA. Two fig of denatured
calf thymus DNA were added as carrier. The reaction mixture
was extracted twice with phenol and once with chloroform, the
nucleic acids were precipitated in the presence of 96% ethanol
at -20°C. After centrifugation, the nucleic acid pellet was
resuspended in 5 /*1 of loading buffer: 0.025% bromophenol blue;
0.025% xylene cyanol; 2.5% ficoll type 400. Samples ( 2 /*1)
were loaded on 7 M urea-8% polyacrylamide denaturing gel,
run at 50 mA then dried and autoradiographed.
Initiation of M13 DNA synthesis in the presence either of
E. coli RNA polymerase or wheat primase, and RNA primedM13 DNA replication assay
Seven tig of M13 ssDNA (mp 19, +) in a final volume of 500 /d
were primed in the presence of E. coli RNA polymerase in the
following conditions: 50 mM Tris-Hcl pH: 8.0; 5 mM MgCl2;
10 mM DTT, 800 units of RNasin, 250 mM of each of ATP,
CTP, GTP and UTP, and 1 unit of E. coli RNA polymerase.
RNA primer synthesis was for 30 minutes at 37°C. The RNA
primed-M13 DNA was used immediately or it was stored at
-20°C before use in the Ml3 replication assay. Eighty ^1 of
the RNA-primed M13 DNA sample were added to 250 /tM final
of each of dATP, dCTP and dGTP, 20 pCi of [3H] TTP
(500-1500 cpm/pmol) and 0 - 2 0 /il of wheat DNA polymerase
A (0-0.5 tig), B, CI or CII (0-10 tig). The final concentration
of M13 DNA was 1 /ig per assay.
In similar conditions 1 ng of M13 ssDNA in the same buffer
with the same final concentrations of NTP and dNTP, was primed
in the presence of different amounts of wheat primase (0—2 /ig).
The RNA primed-DNA was elongated in the presence of 0.5 /tg
of wheat DNA polymerase A, 10 /ig of wheat DNA polymerases
B, CI or CII, or 0.02 /ig of E.coli DNA polymerase I. Replication
of the RNA primed M13 ssDNA was for 1 hr at 37 °C in the
presence of 0.07% polyethylene glycol PEG (type 400) and
0.05% triton X-100. The reaction was stopped as described before
and the precipitated radioactivity counted in a PPO-POPOPtoluene scintillation mixture.
Polyacrylamide gel electrophoresis
Sodium dodecyl sulphate polyacrylamide gel electrophoresis
(SDS-PAGE). SDS-PAGE was performed according to Laemmli
(34) and proteins were stained using the silver nitrate method (35).
Protein determination
Protein concentrations were determined by the method of
Bradford (36) using the bovine serum albumin as standard.
RESULTS AND DISCUSSION
Four DNA polymerases have been described from wheat germ
in our laboratory (22-26, 37). DNA polymerase B has been
described as a possible 5-like polymerase (Richard et al; submitted
for publication). DNA polymerase CI is a low molecular weight
enzyme similar to animal DNA polymerase /3 (26) whereas DNA
polymerase CII is an a-like polymerase. Wheat DNA polymerase
A has been compared to animal DNA polymerase y mainly
because of the preferred recognition of poly rA-oligo dT,
stimulation by KC1 in the presence of this template and its
resistance to aphidicolin (22,25). However, unlike the situation
with animal DNA polymerase y , wheat DNA polymerase A is
different from wheat mitochondrial DNA polymerase (27,28).
In addition, DNA polymerase A was preliminary described as
the only wheat DNA polymerase able to recognize a synthetic
RNA primer (30,31). These results and the association of this
enzyme with wheat DNA primase raises the question of the
possible involvement of this enzyme in the initiation of DNA
synthesis. Very recent work in our laboratory shows that DNA
polymerase A shares some striking similarities with retroviral
reverse transcriptase, thereby suggesting the involvement of this
enzyme in recombination events leading to the modelling of the
nuclear genome (37).
Purification of wheat primase and association with DNA
polymerase A
The first steps of the purification procedure (see Methods) were
the same as those previously described (22—25). Proteins
precipitated between 20 and 70% ammonium sulphate were
dialyzed and loaded on a first phosphocellulose column. As
illustrated in the purification diagram the wheat primase was
divided into two fractions: the first which was not retained in
the phosphocellulose support (about 50—60% of total primase
activity) had a very low amount of DNA polymerase activity;
the second retained on this first chromatographic support
contained the remaining primase activity and was eluted with the
bulk of wheat DNA polymerases at 0.8 M KC1. This fraction
of primase (40-50%) is probably under-estimated due to the
strong inhibition of the wheat primase activity in the presence
of KC1 (see figure 6). The same situation has been described in
mouse cells where 30% of the primase activity does not bind
to the phosphocellulose column, whereas all DNA polymerase
activity is retained (38). We focused our interest on this second
fraction of primase activity coeluting with the bulk of wheat DNA
polymerases. After dialysis, proteins eluted from the first
phosphocellulose column were loaded on a DEAE-cellulose
column: no primase activity was found in the flow through
fractions where DNA polymerases CI and CII eluted, while a
significant peak of primase activity was found with DNA
polymerases A and B eluted at 0.3 M KC1 (see figure 1, panel
A). This is a surprising result, since no activity has been found
to be associated with the wheat a-like DNA polymerase (CII),
while in animal and yeast cells DNA polymerase a purifies
extensively with DNA primase (4, 5, 7—9).
The situation in rice cells is different (39). These authors
showed that, unlike the animal primase, the bulk of the rice
primase is found in the flow through of the phosphocellulose
column, while the rest is retained by the resin. Both primase
fractions seem to be free of a- or 7-like DNA polymerases, a
situation different to what we have observed in the wheat system.
As mentioned in ref 40 a primase associated with a DNA
polymerase a-like activity is present in pea.
Wheat DNA primase, associated with DNA polymerases A
and B, was chromatographed on a second phosphocellulose
4870 Nucleic Acids Research, Vol. 18, No. 16
pellet
WHEAT GERM
homogenization
centrifugation
I
supernatant (cytosol)
S 100 000
20-70% (NH 4 ) 2 SO 4
centrifugation
I
pellet
supernatant
I
I
CRUDE EXTRACT
phosphocellulose I
I
Fraction Not Retained
(primase free)
Fraction Retained
(primase, 4 DNA pol)
I
DEAE-cellulose
I
Fraction Not Retained
Fraction Retained
(primase, DNA pol A,B)
(a-like DNA pol CI.CII)
I
Phosphocellulose II
I
Fraction Retained
Fraction Not Retained
(DNA pol A-primase,B)
(primase free)
I
DEAE-cellulose
(0.1-0.6 MKCI)
I
Heparine-Sepharose
(0.1-0.6 MKCI)
•0.8
-0.2
PURIFIED PRIMASE
Diagram of purification.
column and eluted with a continuous salt gradient. As in the first
phosphocellulose step, the wheat primase was separated into two
fractions: the first, which is the major form (about 75%), was
not retained in the phosphocellulose and was completely devoid
of DNA polymerase activity. The unretained fraction was purified
further (see diagram). The second fraction was eluted at 0.45
M KC1 coinciding exactly with the DNA polymerase A peak.
No primase activity was found to be associated with DNA
polymerase B (5-like) (figure 1, panel B). This highly
reproducible separation of wheat DNA primase on
phosphocellulose into two fractions reflects a significant
association between DNA polymerase A and DNA primase in
wheat germ. In yeast cells only a minor free primase peak is
detected as a shoulder of activity (5—10% of total activity) when
primase is chromatographed on phosphocellulose (8).
A DEAE-cellulose column was used to further purify the DNA
polymerase activity-free primase which was excluded from the
second phosphocellulose column. After extensive washing, the
proteins retained were eluted by a continuous KC1 gradient
( 0 . 1 - 0 . 6 M ). The DNA primase was eluted at 0.18 M KC1.
The activity peak was then loaded after dialysis on heparinesepharose and eluted with a salt gradient (0.1 - 0 . 8 M KC1). the
primase was eluted at 0.25 M KC1, dialyzed and kept at -20°C
in 50% glycerol in buffer A for several months without apparent
loss of activity.
This purified fraction of primase activity, which was devoid
of nuclease and DNA polymerase activities, was used for all the
Fig. 1. Chromatographic pattern of Wheat primase and DNA polymerase A. Five
p] of each fraction were assayed for DNA synthesis in the presence of activated
DNA ( • — • ) or poly rA-oligo dT (D—D), as described in the Methods section
(DNA polymerase assays). The same fractions were tested in the presence of
poly dT ( • — • ) , as described for the primase assay; owing to a strong inhibition
of wheat primase by KCI, 5 /il of each fraction were tested in a 100 pi reaction
volume to dilute the KCI concentration. A. DEAE-cellulose chromatogram. The
pool of enzyme activities retained and eluted at 0.8 M KCI on the first
Phosphocellulose column (see Methods and diagram of purification) was loaded
on a DEAE cellulose column and chromatographed as described in Methods.
No primase activity was detected in the flow through with the a-like DNA
polymerases CI, CII. Primase was eluted stepwise at 0.3 M KCI with DNA
polymerases A, B. B. Phosphocellulose II chromatogram. The pool of DNA
polymerases A and B and wheat primase from the DEAE-cellulose fraction were
loaded on a second phosphocellulose column. After extensive washing, proteins
were eluted by 200 ml of a linear gradient of KCI (0.2-0.8 M). The bulk of
DNA primase activity was not retained and was further purified on DEAE-cellulose
II and heparine-sepharose (see diagram of purification). A significant primase
fraction was coeluted with DNA polymerase A (very active with poly rA-oligo
dT as a template) but not with DNA polymerase B.
experiments described in this work, except for the RNA primedDNA replication assays where the primase activity from the
heparine-sepharose fraction was too scarce; in this case, we used
the DEAE-cellulose fraction which was significantly more active.
The loss of activity in the last purification step could be due to
the rather weak protein concentration (proteins after the last step
of purification as determined by the Bradford method were in
the limit of detection and the enzyme was quite unstable in
comparison with the previous fraction i.e the DEAE-cellulose
fraction). Table I is the purification summary of the
Nucleic Acids Research, Vol. 18, No. 16 4871
Table 1. Purification of DNA primase.
Fraction
Volume
(ml)
Protein
(mg/ml)
Phosphocellulose I
retained
DEAE-cellulose I
Phosphocellulose II
not retained
DEAE-cellulose II
Heparine sepharose
90
1.22
56
30
0.52
0.30
85
132
0.11
0.025
539
116.8
7
3.6
Specific activity
(units/mg)
20.6
Fold
Purification
1
4.2
8.1
26.2
5.7
X
One unit is defined as the amount of primase enzyme in the assay giving rise
to the incorporation of 1 pmole of TMP in 1 hour at 37°C in the presence of
0.5 unit of E.coli DNA polymerase I.
1
2
CO
—
KDa
I
I
.43
.30
Fig. 2. Sodium dodecyl sulphate polyacrylamide gel electrophoresis. Samples
of 5 jig of protein, from the first and last fractions of DNA primase purification,
were subjected to SDS-PAGE (10% polyacrylamide), and run at constant voltage
of 100 V. After electrophoresis proteins were silver stained. Lane 1:
phosphocellulose I, retained fraction, lane 2: heparine-sepharose fraction.
Electrophoretic mobility of molecular weight markers was indicated on the right
of the picture.
chromatographic steps leading to the obtention of a partially
purified wheat primase devoid of all four DNA polymerases (A,
B, CI and CII) and nucleases. We checked for the possible
contamination of our purified primase fraction with a classical
nuclear RNA polymerase. Two results argue against the presence
of RNA polymerases: first, the primase acitivity of the heparinesepharose fraction was not inhibited by a-amanitin (1 mg/ml),
a known inhibitor of eukaryotic RNA polymerases II and III
(RNA polymerase I, which is not affected by this drug, is strongly
associated with the nucleolar fraction, and therefore is hardly
10
20 Ml
Fig. 3. Thermodenaturation and trypsin digestion of wheat primase. Poly dT
replication assay was performed as described in the Methods section. AThermodenaturation of primase. ( • — • ) control reaction, (A—A) 0.25 /ig of
primase previously heated for 10 minutes at 80°C, ( • — • ) primase minus ATP,
(A—A) without primase. B-Trypsin digestion of primase. Wheat primase (0.25
lig per assay) (• — • ) or 0.5 unit of DNA polymerase I (A—A), were incubated
in the presence of different concentrations of trypsin (0-500 jig/ml) for 30 minutes
at 37°C, before addition to the poly dT replication mixture. Action of trypsin
was initially abolished by preincubation of trypsin with the trypsin inhibitor, before
addition of the enzyme wheat primase (D—O) or E.coli DNA polymerase I
( • — • ) . Control reaction without any treatment (O—O).
found in the soluble fraction). Secondly, RNA polymerase
recognizes very efficiently a poly d (AT) template: no activity
was detected in the presence of poly d (AT) with purified
preparations of wheat primase, indicating that probably no RNA
polymerase was present. RNA polymerase II requires a higher
ionic strength than wheat primase for optimal activity: (for a
review on plant RNA polymerases see Becker, W.M., 41).
Moreover, RNA polymerase II, the major transcription enzyme
in wheat germ and soybean hypocotyl is strongly retained in a
phosphocellulose column, (42,43) which is not the case with
wheat DNA primase. RNA polymerase II was also purified
through a DNA-cellulose chromatographic step, while wheat
DNA primase was not retained in this support (our unpublished
results).
Our results seem to indicate that wheat primase cannot be
compared to the 'specific primase stimulating' factor of bovine
thymus described before (44). This factor is very unstable and
leads to a complete loss of activity in a few weeks, while wheat
primase can be stored for several months in 50% glycerol at
—20°C without loss of activity; moreover, the bovine factor is
4872 Nucleic Acids Research, Vol. 18, No. 16
retained on DNA cellulose in contrast to wheat primase. Nor
does wheat primase resembles the rabbit liver factor D, a poly
dT template stimulatory protein of DNA polymerases. This factor
is also very unstable: most of the activity was lost after storage
for 6 days even in the presence of 0.3 mg/ml bovin serum
albumin (45).
We analysed the protein composition of the most purified
fraction by electrophoresis on denaturing gels (SDS-PAGE) as
described in the Methods section. As shown in Figure 2 the
heparine-sepharose fraction is enriched in a doublet of about 90
Kda corresponding to the molecular weight estimated for primase
after centrifugation on urea-glycerol gradient (30). It remains to
be established which of the two polypeptides supports DNA
primase activity.
Biochemical characterization of wheat primase
Biochemical properties. Bivalent ions are required by all enzymes
involved in nucleic acid metabolism. Thus, we looked for Mg
and Mn optima in the assay for primase activity. No activity was
detected in the absence of these metals. The optimum for Mg
was between 1 -5 mM, while for Mn a sharp peak of 0.2 mM
was obtained. The best pH in the primase assay was 8.0. We
looked for the optimal temperature in the indirect assay with poly
dT and ATP as substrates for primase. The best results were
obtained at 37°C, while no activity was detected at 0, 10, 20
or 25°C. At 42°C the enzyme activity was lower than at 37°C,
and a plateau, not observed at 37°C, was noticed. We checked
the temperature sensitivity of primase by submitting the purified
primase fraction to thermodenaturation at 80 °C for 10 minutes
before adding the enzyme to the usual reaction mixture. Under
these conditions less than 25 % of the activity could be detected
(Figure 3a).
Trypsin digestion completely abolished wheat DNA primase
activity. After incubation, trypsin activity was stopped by the
soybean specific inhibitor. As DNA primase is usually assayed
indirectly using E. coli DNA polymerase I, it was crucial to make
controls to check the following: a) whether the bacterial DNA
polymerase was affected by trypsin, b) whether proteolysis was
efficiently arrested by the specific inhibitor and, c) whether the
soybean inhibitor had any effect itself on primer and DNA
synthesis. Results shown in Figures 3a and 3b clearly show that
the wheat primase activity (heparin-agarose fraction) was
abolished by trypsin digestion. These results, like those on the
thermal stability of the enzyme, point to the proteinic nature of
the enzyme, and argue against speculation concerning the
B
1 , 1 5 , 3 0 , 4 5 , 1 ,15,30 ( 4 5
1 2
34
56
H
1 23
Fig. 4. Electrophoretic analysis of the RNA primer. M13 replication reactions
were carried out as described in the Methods section with [a32P] ATP and [a31?]
UTP (500—1500 cpm /pmol) as labeled substrates. After 60 minutes of incubation
at 37°C, the reaction product was initially digested in the presence of 2.5 ng
DNase I (lane 2), or in the presence 2.0 ng of pancreatic RNase (DNase free),
for 15 minutes at 37°C (lane 3), before the reaction was stopped by the addition
of 0.03% SDS, 5 mM EDTA. An equal volume of loading buffer (O.O25K
bromophenol blue, 40% sucrose) was added before loading on 7 M urea-12%
polyaciyiamide gel). Lane 1 shows the control reaction product without any
treatment.
Fig. 5. Primer synthesis in the presence of single-stranded M13 DNA. A-Primasedependent replicative double-stranded M13 DNA synthesis. The purified heparine
sepharose fraction (devoid of nuclease) (0.5 ftg) was incubated for 45 minutes
in the presence of M13 ssDNA (0.5 ^g) and all four rNTP at 37°C (see Methods).
Then, 0.5 units of DNA pol I plus 250 /iM dCTP, dGTP, TTP and 0.5 jtCi
[a 32 P] dATP (3000 Ci/mmol) for 1 min (lanes 1, 5), 15 min (lanes 2, 6), 30
min (lanes 3, 7) or 45 min (lanes 4, 8). In lanes 1, 2, 3 and 4 primase was not
present in the reaction mixture ( - ) ; in lanes 5, 6, 7 and 8 primase was present
(+). B-Sequencing of the DNA polymerase I elongated product when the template
M13 ssDNA was primed with the 17 nt universal primer or by action of wheat
primase. In parallel experiments 1 /ig M13 ssDNA was either primed in the
presence of 0.3 ng of the M13 17 nt primer or in the presence of 0.5 /tg primase
(heparine-sepharose fraction) and rNTP (see Methods). The primed M13 DNA
was then elongated in the presence of dNTP plus ddNTP and E.coli DNA
polymerase I for a sequencing reaction, as described by Sanger et al. Lanes 1,
3, 5 correspond to the adenosine ladder, and lanes 2,4, 6 to the guano^ne ladder.
Lanes 1, 2 to the presence of primase in the priming reaction; lanes 3, 4 to the
absence of the primase and lanes 5, 6 to the presence of the 17 nt primer as the
priming system.
Nucleic Acids Research, Vol. 18, No. 16 4873
artifactual involvement of polyphenols in the in vitro assay of
plant DNA primase activity (46).
Analysis of the product synthesized by the primase. Preliminary
experiments reported previously showed that wheat primase was
able to recognize M13 single-stranded DNA as template (30).
In order to gain a further insight into this recognition process
the experiment described in Figure 4 was performed. As
described in the Methods section, labeled RNA primer was
synthesized in the presence of primase and M13 (mp 9, +) singlestranded DNA as template with [a32?] UTP and [a32P] ATP as
labeled precursors. After 60 minutes of incubation at 37°C the
reaction was stopped by the addition of SDS and EDTA, and
the reaction product was analysed by electrophoresis on ureapoly aerylamide gels. The control reaction product is shown in
Figure 4 (lane 1). When compared to the electrophoretic mobility
of molecular weight markers (not shown), the size of the RNA
primer is between 20 to 200 nucleotides in length
We checked the sensitivity of the primase reaction product to
RNase digestion (Figure 4, lane 3). Complete disappearance of
the labeled product after RNase digestion demonstrated that the
primase reaction product is RNA. The same product was totally
resistant to DNase I (RNase-free) digestion (Figure 4, lane 2).
RNA primer length was dependant on the presence of dNTP.
As described in the case of the calf thymus system (47), primers
of 8-15 nucleotides were synthesized in the presence of dNTP,
both with poly dT and M13 DNA. In the absence of dNTP,
primers of 20-40 nucleotides were formed. The size of the RNA
primer synthesized by the wheat primase was previously analysed
by elongation of the primase product by DNA polymerase I in
the presence of [a32P] dATP and poly dT as the template,
followed by digestion with DNase I (30). We concluded from
these experiments that a heterogenous-sized primer RNA ranging
from 2 to 15 residues was obtained. However, in the absence
of dNTP and when the product was labeled with [732P] ATP
and submitted to DNase I digestion, the size of the
oligoribonucleotide primers was much higher (30-100 nt) with
poly dT as the template. The large size of the product labeled
in the absence of dNTP prevented them from entering the gel.
a?
Fig. 6. Effect of KC1 on primase. Wheat primase (0.5 /ig) was tested with variable
concentrations of KC1 (0-200 mM) in the presence of poly dT as described in
the Methods section ( D — • ) . The E.coli DNA polymerase I (0.5 unit) was tested
in the presence of poly dT-oligo rA (• — • ) . Activities were in % as compared
to the control reaction (0 mM KG).
These results indicate the close interactions controlling primase
and DNA polymerase activities. Very recently the influence of
the primer size in the initiation of DNA synthesis was described
(48); these authors concluded that all mononucleotides and both,
oligo (1 - 2 5 nucleotides) and longer polynucleotides (100—300
nucleotides) served as primers for DNA synthesis.
Replication of single-stranded M13 DNA. We analysed the ability
of the wheat primase to allow the synthesis of the M13 doublestranded replicative form (ds RF) by DNA polymerase I. The
primase was incubated for 45 minutes in the presence of singlestranded M13 DNA (ssDNA) and all four rNTP at 37°C (see
methods). After this incubation to allow primer synthesis, RNA
primer was elongated for different time periods by DNA
polymerase I in the presence of 0.5 /tCi [a 32 P] dATP and
250 /tM of each of the three unlabeled dNTP. Figure 5 shows
that, only when primase is present, can DNA polymerase I
replicate M13 ssDNA, giving rise to the slower migrating M13
dsDNA (RF) (Figure 5a, lanes 5, 6, 7, 8). In the absence of
primase, no primer was available for DNA polymerase I and
only DNA repair activity was possible, as shown by the labeling
of the M13 ssDNA at 1, 15, 30 or 45 minutes of incubation
(Figure 5a, lanes 1, 2, 3, 4).
The specificity of the priming reaction was analysed by
sequencing the elongated product synthesized by DNA
polymerase I in the presence either of M13 ssDNA primed with
0.5 ng of the universal 17-mer M13 primer, or M13 ssDNA
preincubated with rNTP, with or without primase. The
sequencing reaction was performed according to the
dideoxynucleotide method (49). When the universal primer was
used as a primer for M13 DNA synthesis, a clear ladder for
adenosine and guanosine residues was obtained when analysed
on a 6% polyacrylamide-7 M urea sequencing gel (Figure 5b,
lanes 5, 6). In the absence of primase, no product was observed
(Figure 5b, lanes 3, 4). In the presence of primase and rNTP,
the product observed was a smear very similar in size to the
control lane (Figure 5b, lanes 1,2). The latter were more visible
on longer exposures (not shown). This result shows that the wheat
primase seems to lack specificity and initiates RNA synthesis at
random sites on the M13 genome.
The effect of inhibitors on wheat DNA primase activity
Effect of KCl. During the purification procedure we observed
that the primase was very sensitive to the salts used for elution.
As shown in Figure 6, DNA primase was dramatically inhibited
by KCl: at 50 mM KCl no activity was observed. As mentioned
before the indirect assay of primase in the presence of poly dT
as template depended on the use of an exogenous DNA
polymerase (E. coli DNA pol I) for primer elongation. Thus,
we checked the effect of KCl on DNA polymerase I: this enzyme
was totally resistant to 0.1 M KCl and showed the specific effect
of KCl on wheat primase under our assay conditions. A similar
inhibition of primase has been described in yeast, even if the latter
is slightly more resistant to KCl (50). A similar inhibition of DNA
primase by KCl has been described in animal cells (14).
ATP structural analogs. The best template for primase is poly
dT. Poly dC, poly dA or poly d(AT) are not recognized by the
wheat primase (30, and P.L. unpublished results). The specificity
of the ribonucleotide triphosphate substrate in the presence of
poly dT as template was analysed and the results are shown in
Table 2. Only the right combination of poly dT template and
4874 Nucleic Acids Research, Vol. 18, No. 16
ATP as precursor for primase plus dATP as the substrate for
DNA polymerase A gave rise to a significant incorporation. If
any other rNTP or dNTP was used instead of ATP or dATP (i.e
UTP and TTP) less than 10% of the original activity was
observed; this illustrates the specificity of the incorporation.
As expected the wheat primase is absolutely dependent on ATP
to synthesize an oligo rA primer in the presence of poly dT as
template (Figure 3a). We studied the effect of structural analogs
of ATP on primase activity (table 3). Cordycepine triphosphate
has a hydroxyl group in carbon 2' instead of carbon 3'. For this
reason the phosphodiester 3' —5' bond of the phosphate backbone
cannot be made. At 0.1 mM cordycepine triphosphate more than
90% of primase activity is inhibited. Under the same
concentrations, E. coli DNA polymerase I was less inhibited than
the wheat primase. These results indicate that wheat primase is
not able to discriminate between the right substrate, ATP present
at 1 mM in the assay, and this analog. The second analog,
formycine triphosphate, is a well known antibiotic isolated from
actinomycetes (Neocordia interforma) and known for its antiviral
properties (51,52). DNA polymerase I is resistant when tested
in the presence of DNA. The effect of formycine triphosphate
on wheat primase activity was less pronounced than in the case
of cordycepine triphosphate: at 1 mM formycine triphosphate
only about 15% inhibition of wheat primase activity was obtained.
We also studied the effect of ADP, cyclic AMP, dATP and
ddATP on the poly dT replication assay always in the presence
I
Table 2. Specificity of poly dT replication in the presence of primase and DNA
polymerase A.
rNTP
dNTP
dATP
+ DNA polymerase A
3%
100%
0
- DNA polymerase A
2.6%
9.5%
20
80
100
120
Time (min)
rATP
TTP
10.6%
1.1%
2.9%
1.1%
dATP
2.8%
6.1%
2.4%
2%
TTP
5%
0.7%
1.5%
1.4%
rUTP
Two jig of Wheat primase was tested in the presence of poly dT as template
and without (—), or with (+) 0.5 /ig of wheat DNA polymerase A as the replicative
DNA polymerase, at 37°C for 1 hr, as described in the Methods section. Poly
dT replication was tested either in the presence of UTP or ATP, dATP or TTP
as substrates for the wheat primase and DNA polymerase A. Activities were in%
as compared to the control reaction (100% corresponding to 17 735 cpm).
10
Table 3. Effect of ATP structural analogs on primase.
control reaction
Formycine
triphosphate
Cordycepine
triphosphate
ADP
cAMP
dATP
ddATP
0.1 mM
0.4 mM
1 mM
0.1 mM
0.4 mM
1 mM
0.1 mM
0.4 mM
1 mM
0.1 mM
0.4 mM
1 mM
5 ^M
50 yU
0.1 mM
0.5 ^M
5 fiM
10 ^M
wheat primase
DNA polymerase I
100
100
115
140
110
78
40
16
148
135
98
92
86
70
120
42
18
74
20
12
100
105
86
10
0
0
75
68
55
102
108
96
21
2
0
64
18
10
Wheat primase (0.5 /»g) was tested in the presence of poly dT template,
p
[3H] dATP, 1 mM ATP and 0.5 unit of DNA polymerase 1 plus different
amounts of ATP structural analogs. E.Coli DNA polymerase I was tested in the
same conditions with poly dT-oligo rA as the template. The activity is in % as
compared to the control reaction (without analog).
Fig.7. Recognition of natural RNA-primed DNA by wheat DNA polymerases
A, B, CI and CM. A. M13 ssDNA primed in the presence of E.coli RNA
polymerase. The enzymes were tested in the presence of 1 /ig ssDNA primed
in the presence of E.coli RNA polymerase and NTP, prepared as described in
the Methods section, and 250 fiM of each of dATP, dCTP, dGTP and
20 /iM [3H] TTP (500-1500 cpm per pmol) at 37°C for 1 hr. ( O- 3)
0 - 0 . 5 fig DNA polymerase A, (A—A) 0 - 2 ^g DNA polymerase B, (A—A)
0 - 2 /ig DNA polymerase CI, ( • — O ) 0 - 2 pg DNA polymerase CI1 tested
in the presence of RNA-primed M13 ssDNA. No activity was detected in the
absence of XTP or E.coli RNA polymerase in the priming reaction (results not
shown). ( • — • ) 0-0.5 /ig DNA polymerase A tested in the presence of singlestranded cDNA, synthesized for 1 hr at 37 °C by 2 units of the AMV reverse
transcriptase in the presence of 2 ^g globin mRNA and 500 iM of all four dNTP
and oligo dT as the primer. After RNA digestion the cDNA was precipitated
and resuspended in 20 jil sterile water and 5 /il of this cDNA solution was tested
in the presence of 0.05 A2GO °f oligo rA as primer and 20 pM [3H] TTP as
labeled substrate in a final 100 p.] reaction buffer, as described for the M13 DNA
replication assay. In the absence of the oligo rA as primer or the three unlabeled
dNTP, no activity was detected. B. Dependance on the amount of wheat primase
for the recognition of natural DNA. Denatured calf thymus DNA was primed
in the presence of different amounts of the DEAE-cellulose fraction of wheat
primase (0 to 2 /ig) and 0.02 ng E.coli DNA polymerase I ( • — • ) , 0.5 mg
DNA polymerase A (• — • ) , or 2 ng DNA polymerase B (A), CI (A), CU
(LJ) at 37°C for 1 hr as described in the Methods section.
Nucleic Acids Research, Vol. 18, No. 16 4875
of 1 mM ATP. As for ddATP no conclusion can be made about
the specific effect of this ATP analog: the same inhibition was
observed in the primase and DNA polymerase I assays. In
comparison, dATP, even at very low concentration, gave rise
to a significant inhibition of primase activity. This effect cannot
be explained merely on the basis of the isotopic dilution of this
precursor. Thus, at 5 fiM dATP (about half the [3H] dATP
concentration in the assay), 80% of primase activity was inhibited;
at 50 jiM dATP (5 times the [3H] dATP concentration) 98% of
primase was inhibited. In the same range of isotopic dilution of
the labeled precursor, DNA polymerase I was first stimulated
(at 5 fiM dATP about 120% of activity was detected), while at
50 fiM DNA polymerase I, 42% of activity was detected as
compared with the control. This illustrates that the primase is
sensitive to the dATP present in the assay at much lower
concentrations than ATP. In comparison, cyclic AMP had no
significant effect on the primase. In addition, ADP, had a weak
inhibition effect on the primase: at 1 mM ADP 55 % of primase
activity was still detected.
In vitro DNA synthesis is dependent on primase and DNA
polymerase A from wheat germ
On the basis of the copurification of primase and DNA
polymerase A, and the recognition of RNA primer by the only
DNA polymerase A when tested with poly dT as template (30),
we looked for the effect of KC1 on the recognition of poly dToligo rA by DNA polymerase A, E. coli DNA polymerase I and
DNA polymerases a and y from animal cells (a generous gift
of Dr. L. Tarrago-Litvak). DNA polymerase I was not affected
by KC1 while animal DNA polymerase g was strongly stimulated
(at 200 mM KC1 the activity was stimulated 10 fold); DNA
polymerases A and a were 90% inhibited at 200 mM KC1 (results
not shown). In this case wheat DNA polymerase A behaves
similarly as the animal replicative DNA polymerase a.
We studied the effect of ATP and cordycepine triphosphate
on the activity of DNA polymerase A, a and y when tested in
the presence of DNA. ATP had the same effect on DNA
polymerase A and a (not inhibited or slightly stimulated), while
DNA polymerase y was inhibited 50% at 15 mM ATP; the same
behavior of these enzymes was observed in the presence of poly
rA-oligo dT (results not shown). The stimulatory effect of ATP
on replicative DNA polymerase a has been described elsewhere
(53—55). This point can be also illustrated with the simian virus
SV 40 replication model, since the formation of an active initiation
complex between the T antigen and the origin of replication is
dependent on and stimulated by ATP (57). However, hydrolysis
in this case was not essential, since ATP could be replaced by
ADP, adenosine 5' (beta, gamma, imido) triphosphate, or dATP
at a concentration 30 times lower than that of ATP, while dGTP
or rGTP were inactive. This result might explain why
cordycepine triphosphate does not inhibit DNA polymerase A
when tested in the presence of poly dT-oligo rA, poly rA-oligo
dT or DNA. Owing to a possible interaction of DNA polymerase
A with the wheat primase in DNA initiation, we also checked
the effect of cordycepine triphosphate on DNA polymerase A
activity tested with poly rA-oligo dT. This enzyme was
significantly stimulated by this ATP analog (as with poly dToligo rA), even though it is not a substrate with this template
(results not shown). In these conditions the structural analog could
also play a stimulating role, as in the case of ATP in the binding
of DNA polymerase A to a natural template or to poly dT; such
is the case for the complex between replicative DNA polymerase
a and the primase in animal cells. Nevertheless, replication of
synthetic templates is rather more efficient than with natural
template, so we think that several factors may be missing in the
in vitro assay with the purified proteins. Our observations
concerning the copurification of wheat primase and DNA
polymerase A, the similar behavior of DNA polymerase A
compared to the a DNA polymerase when copying a DNA
template, and its association with the wheat primase, all support
the possible role of wheat DNA polymerase A in the initiation
of wheat DNA synthesis.
In a final experiment, the ability of wheat DNA polymerases
to replicate RNA-primed DNA was tested. Wheat DNA
polymerase A was the sole enzyme able to initiate DNA synthesis
in the presence of natural RNA-primed DNA, either in the
presence of M13 ssDNA or denatured calf thymus DNA primed
by E. coli RNA polymerase (Figure 7A). Wheat DNA
polymerases B, CI and CII tested even at higher protein
concentrations (2 ng instead of 0.5 jtg for DNA polymerase A)
did not recognize the RNA-primed M13 DNA. Incorporation was
dependant on the amount of enzyme, and increased linearly from
0 to 20 nl of protein, i.e. from 0 to 0.5 ng of DNA polymerase
A. The ability of these enzymes to recognize M13 DNA in the
presence of the 17 nucleotide universal M13 primer was also
analyzed, and only wheat DNA polymerase A was able to
elongate this primed M13 ssDNA (result not shown).
In a second assay DNA was primed by the wheat primase in
the presence of all four rNTP, and the ability of different enzymes
to elongate this template was analyzed. Only E. coli DNA
polymerase I (0.02 ng) and wheat DNA polymerase A (0.5 ^g)
could give rise to a significant incorporation of labeled TMP into
the neosynthesized DNA (figure 7B). Incorporation was
dependant on the amount of wheat primase as priming enzyme,
and increased linearly up to 1 /tg of wheat primase (10 /il of the
DEAE-cellulose fraction devoid of contaminating nucleases).The
same level of incorporation was observed in the presence of the
complex DNA polymerase A-primase (DEAE-I retained fraction)
which may be similar to the functional entity in vivo. In the
absence of primase a weak incorporation of dTMP was observed
with both enzymes probably reflecting DNA repair activity. This
result is in agreement with the chromatographic behavior of wheat
DNA primase, which copurifies extensively with DNA
polymerase A. Our results together seem to indicate that wheat
DNA polymerase A is probably involved in initiation of the
nuclear DNA synthesis in wheat. This enzyme seems different
both from the wheat mitochondrial DNA polymerase, which is
unable to recognize poly rA-oligo dT as template (28, 29), and
from the chloroplastic DNA polymerase (P.L. unpublished
results) and suggest a nuclear origin of DNA polymerase A and
primase. It is surprising that neither the wheat a-like DNA
polymerase (CII) nor the 6-like enzyme (B) were able to recognize
a natural or synthetic RNA-primed DNA template. This may be
due to the loss of some cofactors required for these enzymes.
The role of these enzymes in the wheat germ remains to be
elucidated and the development of a plant viral model,even if
the plant equivalent of animal virus SV 40 has not been described,
may throw considerable light on knowledge of DNA replication
in plants.
ACKNOWLEDGMENTS
This work was supported by the CNRS and the University of
Bordeaux n. The authors are grateful to Dr. Laura Tarrago-Litvak
for improving the manuscript.
4876 Nucleic Acids Research, Vol. 18, No. 16
REFERENCES
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
Kornberg,A. (1980) DNA replication .W; H. Freeman. San Francisco.
Kornberg.A. (1982) supplement to DNA replication.
Yagura,T., Kozu.T., and Senzo.T. (1982) J. Biol. Chem 257, 11121 -11127.
Conaway,R.C. and LehmanJ.R. (1982) Proc. Natl. Acad. Sci. USA 79,
2523-2527.
Hubscher.U. (1983) Embo J. 2, 133-136.
Hu,S.Z., Wang.T.S.F. and Korn.D. (1984) J. Biol. Chem. 259, 2602-2609.
Shioda,M., Nelson.E.M., Bayne.M.C. and Benbow,R.M. (1983) Proc. Natl.
Acad. Sci. USA 79, 7209-7213.
Plevani,P., Badaracco.G., Augl.C. and Chang,L.M.S. (1984) J. Biol. Chem
259, 7532-7539.
Singh.H. and Dumas.L.B. (1984) J. Biol. Chem 259, 7936-7940.
Kaguni,L.S., Rossignol,J.M., Conaway,R.C. and Lehman,I.R. (1983) Proc.
Natl. Acad. Sci. USA 80, 2221-2225.
Hinckle,D.C. and Di Gate.R.J. (1983) J. Cell. Biochem. supplement 7B, 121.
Riedel.H.D., Konig.H., Stahl,H. and Knippers.R. (1982) Nucl. Acids Res.
10, 5621-5635.
Brooks,M. and Dumas.L.B. (1989) J. Biol. Chem. 264, 3602-3610.
14. Wang.T.S.F., Hu,S.Z. and Korn,D. (1984) J. Biol. Chem. 259, 1854-1865.
15. Tubo.R.A. and Berezney.R. (1987) J. Biol. Chem. 262, 6637-6642.
16. Suzuki,M., Enomoto,T., Masutani,C, Hanaoka,F., Yamada,M. and Ui,M.
(1989) J. Biol. Chem. 264, 10 065-10 071.
17. Vishwanatha,J.K. and Baril.E.F. (1986) Nucl. Acids Res.14, 8467-8487.
18. Wilson.F.E. and Sugino,A. (1985) J. Biol. Chem. 260, 8173-8181.
19. Kaiserman,H.BandBenbow,R.M. (1987) Nucl. Acids Res.15, 10 249-10
265.
20. Kaguni.L.S. and Lehman,I.R. (1988) Biochem. Biophys. Acta 950, 87-101.
21. Yagura,T., Kozu.T., and Seno,T. (1982) J. Biol. Chem. 257, 11 121-11
127.
22. Castroviejo.M., Tarrago-Litvak.L. and Litvak.S. (1975) Nucl. Acids Res.
2, 2077-2090.
23. Castroviejo.M., Tharaud,D., Tarrago-Litvak,L. and Litvak.S. (1979)
Biochem. J. 181, 183-191.
24. Castroviejo.M., Graves,P.V., Tharaud.D., Hevia-Campos.E. and Litvak.S.
(1982) Biochimie 64, 165-202.
25. Tarrago-Litvak.L., Castroviejo.M. and Litvak.S. (1975) Febs. Letters 59,
125-130.
26. Castroviejo.M., Gatius,M.T. and Litvak.S. (1990) Plant Mol. Biol. in press
27. Christophe.L., Tarrago-Litvak.L., Castroviejo.M. and Litvak.S. (1981) Plant.
Sci. Lett. 21, 181-192.
28. Ricard,B., Echeverria.M., Christophe.L. and Litvak.S. (1983) Plant Mol.
Biol. 2, 167-175.
29. Litvak.S. and Castroviejo.M. (1987) Mutation research 181, 125-130.
30. Graveline.J., Tarrago-Litvak,L., Castroviejo.M. and Litvak.S. (1984) Plant
molecular biology 3, 207-215.
31. Litvak.S., Graveline.J., Zourgui.L., Carvallo.P., Solari.A., Aoyama,H.,
Castroviejo.M. and Tarrago-Litvak.L. (1984) In proteins involved in DNA
replication. Adv. Exptl. Med. Biol. (U. Hubscher and S. Spadari eds) 179
Plenum New York, 249-262.
32. Zourgui.L., Tharaud.D., Solari,A., Litvak.S. and Tarrago-Litvak,L. (1986)
Biochem. Biophys. Acta 846, 222-232.
33. Aposhian,N.V. and Kornberg.A. (1962) J. Biol. Chem. 237, 519-525.
34. Laemmli,U.K. (1970) Nature 227, 680-685.
35. Marshall.T. (1984) Anal. Biochem. 136, 340-346.
36. Bradford.M.M. (1976) Anal. Biochem. 136, 340-346.
37. Laquel.P., Sallafranque-Andreola,M.L., Tarrago-Litvak,L., Castroviejo.M.
and Litvak.S. (1990) Biochem. Biophys. Acta. 1048, 139-148
38. Tseng.B.Y. and Ahlem.C.N. (1982) J. Biol. Chem. 257, 7280-7283.
39. Marchesi,M.L., Villagi.F., Spadari,S., Pedrali-Noy,G. and Sala,F. (1987)
Mutation Research 181, 9 3 - 1 0 1 .
40. Bryant and Dunham (1988) Oxford Survey of Plant Molecular and Cell
Biology, vol 5, pp23-55.
41. Becker.W.M. (1979) in Nucleic Acids in plants, TC;. Hall and Davies;.
JW. Eds. vol 1, 111-141. CRC. Press, Boca Raton, Florida.
42. Jendrisak,J.J. and Burgess.R.R. (1977) Biochemistry 16, 1959-1964.
43. Guilfoyle.T.J., Lin.C.Y., Chen.Y.M. and Key.J.L. (1976) Biochem.
Biophys. Acta 418, 344-357.
44. Itaya.A., Hironaka,T., Tanaka,Y., Yoshihara,K. and Kamiya.T. (1988) Eur.
J. Biochem. 174, 261-266.
45. Sharf.R., Weisman-Shomer.P. and Fry.M. (1988) Biochemistry 27,
2990-2997.
46. Roth.Y.F. (1987) Eur. J. Biochem. 165, 473-481.
47. Grosse.F. and Krauss.G. (1985) J. 3iol. Chem. 260, 1881-1888.
48. Nevinsky.G.A., Veniaminova.A.G., Levina.A.S., Podust.V.N., Lavrik.O.I.
and Holler,E. (1990) Biochemistry 29, 1200-1207.
49. Sanger,F., Nicklens.S. and Coulson.A.R. (1977) Proc. NaU. Acad. Sci. USA.
74,5463-5467.
50. Jazwinski.S.M. and Edelman.G.M. (1985) J. Biol. Chem. 260, 4995-5002.
51. Hori.M., Ito.E., Takita,T., Koyama,G., Takuchi.T. and Umezawa,H. (1964)
J. Antibiotic (Tokyo). 17A, 9 6 - 9 9 .
52. Roy-Burman,P. (1970) In Recent Results in Cancer Research. Spring verlag,
Heidelberg, New York, 7 1 - 7 5 .
53. Smith,H.C. and Berezney.R. (1982) Biochemistry 21, 6751-6761.
54. Faust.E.A. and Rankin,C.D. (1982) Nucl. Acids Res. 10, 4181-4201.
55. Lawton.K.G., Wierowski.J.V., Schester.S., Hilf.R. and Bambara.R.A.
(1984) Biochemistry 23, 4294-4300.
56 Tan,C.K., So.M.J., Downey.M. and So.A.G. (1987) Nucl. Acids Res. 15,
2269-2278.
57. Borowiec,J.A. and Hurwitz.J. (1988) Proc. Natl. Acad. Sci. USA. 85,
64-68.