(Received for publication, December 26, 1996, and in revised form, May 19, 1997)
From the Department of Biological Chemistry, Johns Hopkins University School of Medicine, Baltimore, Maryland 21205
We have purified to near homogeneity a DNA
primase from a mitochondrial fraction of the trypanosomatid
Crithidia fasciculata. The enzyme is a single polypeptide
chain of 28 kDa. Using a poly(dT) template and ATP as a substrate, the
enzyme makes oligonucleotides of which the vast majority are about 10 nucleotides in size or smaller. With a single-stranded M13 DNA template
and the four rNTPs as substrates, the enzyme makes heterogeneous
oligonucleotides in the same size range. These oligonucleotides
efficiently prime the synthesis of DNA by the Klenow DNA polymerase.
Immunolocalization with antibodies against the purified enzyme confirms
that the primase is mitochondrial. Furthermore, the enzyme localizes to specific regions of the cell's single mitochondrion, above and below
the condensed kinetoplast DNA. The primase does not co-localize with
the mitochondrial topoisomerase II and DNA polymerase , both of
which are associated with two protein complexes positioned on opposite
sides of the kinetoplast disc. These localization studies have
significant implications for the mechanism of kinetoplast DNA
replication.
Trypanosomatids are protozoan parasites that cause important diseases in humans and livestock. They have an unusual mitochondrial DNA, known as kinetoplast DNA (kDNA),1 which has a remarkable structure (see Refs. 1-3 for reviews). kDNA consists of several thousand circular DNA molecules, which are all catenated into a giant network. In the trypanosomatid Crithidia fasciculata, the subject of this study, electron microscopy reveals that an isolated network is an eliptically shaped monolayer of interlocked DNA circles, about 10 µm × 15 µm in size. Each cell has one network, which is condensed into a disc-shaped structure within its single mitochondrion. The network contains two kinds of DNA circles known as minicircles and maxicircles. In C. fasciculata the network contains about 5000 minicircles, each 2.5 kilobases, and 25 maxicircles, each 38 kilobases. Maxicircles have a genetic function resembling that of mitochondrial DNAs in other eukaryotes, encoding ribosomal RNAs and proteins required for mitochondrial energy transduction. Most maxicircle transcripts undergo editing, a process by which uridine residues are inserted into or deleted from the sequence, thus forming an open reading frame. Minicircles encode small guide RNAs, which control the specificity of editing (see Refs. 4 and 5 for reviews on editing).
The network structure of kDNA must require unusual mechanisms for its
replication. For example, minicircles do not replicate while
topologically linked to the network, but instead they are released from
the network by a topoisomerase. After release they replicate as free
minicircles, using a mechanism, and the progeny minicircles are
then reattached to the network periphery. When all minicircles have
replicated, the network has doubled in size. Then it splits in two, and
the two progeny networks segregate into daughter cells during cell
division (see Refs. 3, 6, and 7 for reviews).
A current goal in our laboratory is to study proteins and enzymes that
are involved in the maintenance and replication of kDNA. These proteins
are interesting not only in their own right, but also because in at
least some cases they are assembled into mitochondrial protein
complexes, which can be easily visualized by immunofluorescence. For
example, both a topoisomerase II (8) and a DNA polymerase (9)
co-localize within two discrete protein complexes (each roughly 0.4 µm in diameter), which are situated in antipodal positions at the
edge of the kinetoplast disc (the disc is approximately 1 µm in
diameter and 0.4 µm thick). Since minicircles thought to be
replication intermediates (as visualized by fluorescence in
situ hybridization) are also present in these two complexes (9),
it is likely that these structures are involved in minicircle
replication (see Ref. 3 for review).
In this paper we report the purification of a C. fasciculata
mitochondrial DNA primase, the enzyme responsible for making short RNA
molecules that initiate the synthesis of a DNA strand. We found that
the primase is a single polypeptide of 28 kDa, and we describe some of
its enzymatic properties. Using immunofluorescence, we found
unexpectedly that the primase does not co-localize with the
topoisomerase II and the DNA polymerase in the two protein complexes that flank the kinetoplast disc. Instead it localizes above
and below the disc. This finding has significant implications for our
understanding of the mechanism of kDNA replication.
Ribo- and deoxynucleoside triphosphates were
purchased from Boehringer Mannheim, and [-32P]ATP
(3000 Ci/mmol) and [
-32P]dATP (3000 Ci/mmol) were from
NEN Life Science Products. Poly(dT), from Pharmacia Biotech Inc., had
an average length of 200-300 nucleotides.
A coupled assay was used to follow
the enzyme during purification (10). In this assay, using ATP as a
substrate, the primase synthesized non-radioactive oligo(A) primers
complementary to a poly(dT) template. The primers were then extended by
Klenow DNA polymerase and [-32P]dATP, and
acid-insoluble radioactivity was measured. The 25-µl reaction mixture
contained 50 mM Tris-HCl, pH 7.5, 50 mM NaOAc, 5 mM MgCl2, 0.1 mg/ml BSA (the 10 mg/ml stock
solution had been heated at 60 °C for 15 min), 5% (v/v) glycerol, 1 mM DTT, 0.5 µg of poly(dT), 1 mM ATP, 50 µM [
-32P]dATP (about 2400 cpm/pmol),
0-0.5 unit of primase, and 0.5 unit of Escherichia coli DNA
polymerase I large fragment (New England Biolabs). After 30 min at
30 °C, products of the reaction were precipitated by sequential
addition of 10 µl of BSA (10 mg/ml), 10 µl of 0.1 M
sodium pyrophosphate, 35 µl of H2O, and 20 µl of 50%
(w/v) trichloroacetic acid. The acid-insoluble product was collected
onto GF/C filters (Whatman) under vacuum and washed, first with 0.1 M HCl and 0.1 M sodium pyrophosphate, and then with 95% (v/v) ethanol. Radioactivity on the filters was measured by
liquid scintillation counting. One DNA primase unit is defined as the
amount required for conversion of 1 pmol of [32P]dAMP/h
into an acid-insoluble form under assay conditions. The activity is
linear with enzyme concentration in the range of 0-1 unit. The
standard buffer conditions and temperature (30 °C) were used in all
primase reactions described in this paper.
Parasites were grown in a 150-liter Fermatron fermenter (in the Department of Biochemistry, Johns Hopkins School of Hygiene and Public Health) at 26 °C in medium containing 1.8% Deltown AE80M peptone, 0.45% yeast extract, 0.45% NaCl, 0.9% glucose, and 10 mg/liter hemin (11). Alternatively, they were grown at room temperature in 6-liter flasks (each containing 4 liter of medium) with vigorous shaking (200 rpm). Cells at a density of about 4 × 107/ml were harvested by centrifugation either using a Sharples continuous flow centrifuge or a Sorvall GS3 rotor (5000 rpm, 10 min, 4 °C). They were washed in STE buffer (250 mM sucrose, 50 mM Tris-HCl, pH 7.5, 1 mM EDTA) (12). The yield of cells was 2-3 g (wet weight)/liter of culture. Two procedures, both conducted at 0-4 °C, were used for isolation of mitochondria. In Method A (modified from a previous procedure; Ref. 12), the cells (220 g, wet weight) were suspended in 1200 ml of cell disruption buffer (STE buffer supplemented with 0.15 M KCl and 5 mM MgCl2). The cell suspension (in 600-ml aliquots) was transferred to a Parr cell disruption bomb. The bomb was pressurized with nitrogen to 1800-2000 p.s.i. for 30 min, while the contents were stirred with a magnetic bar. The cells were disrupted as the suspension was released to atmospheric pressure through the discharge valve. To maintain the pressure through the discharge process, it was necessary to stop the flow, restore the nitrogen pressure to 1800-2000 p.s.i., and then resume the discharge. Microscopy revealed that more than 95% of cells were disrupted. Fluorescence microscopy, after staining with DAPI (1 µg/ml) to visualize the kDNA, showed that most of the mitochondria appeared as brightly fluorescent dots, indicating that the membranes enclosing the kDNA were still intact. To isolate a mitochondrial fraction, the suspension was centrifuged (Sorvall GS-3 rotor, 9000 rpm, 20 min, 4 °C); the pellet was resuspended in 200 ml of cell disruption buffer and centrifuged again under the same conditions. This washing procedure was repeated four times. In Method B (modified from a scheme developed by Joseph Shlomai, Hebrew University),2 the cells (450 g, wet weight) were suspended in 2000 ml of ice-cold 25 mM Tris-HCl, pH 7.5, 1 mM EDTA and then transferred to the Parr cell disruption bomb. The bomb was pressurized to 1000 p.s.i. for 30 min, while stirring with a magnetic bar. The cells were released from the discharge valve, which was connected to an 18-gauge needle. Microscopy indicated that almost all cells were broken and most mitochondria were intact. The cell homogenate was centrifuged (Sorvall GS-3 rotor, 2000 rpm, 5 min, 4 °C) to remove unbroken cells, and then the supernatant was collected and centrifuged again (Sorvall GSA rotor, 12,500 rpm, 30 min, 4 °C). The pellets from the second centrifugation, containing mitochondria, were pooled and used for the enzyme purification. Method B gives a better recovery of mitochondria, although it is probably a more crude preparation.
Small Scale Growth of C. fasciculata and Percoll Purification of MitochondriaCells were grown to about 4 × 107 cells/ml with vigorous shaking (200 rpm) at room temperature in 8 liters of 3.7% (w/v) Brain Heart Infusion supplemented with 20 µg/ml hemin using two 6-liter flasks. The cells were harvested as described above and disrupted in a Parr cell disruption bomb using Method A. After centrifugation of the lysate to sediment the mitochondria (Sorvall GSA rotor, 12,500 rpm, 30 min, 4 °C), the pellet was washed twice with 200 ml of cell disruption buffer. The final pellet was gently resuspended with 125 ml of 100% Percoll (Pharmacia) and 125 ml of cell disruption buffer by means of a Dounce homogenizer with loose pestle (type B). The Percoll suspension was evenly divided into 10 tubes and centrifuged (Beckman Ti 50.2 rotor, 26,000 rpm, 45 min, 4 °C), generating Percoll concentration gradients. In this gradient the mitochondrial fraction formed a turbid band about 1 cm thick, at about 2/3 of the distance from the top of each tube. These fractions were pooled and recentrifuged (Beckman SW27 rotor, 25,000 rpm, 90 min, 4 °C). The mitochondrial fraction formed a loose layer just above the Percoll pellet. The mitochondria were collected and washed once with STE buffer (Sorvall SS34 rotor, 15,000 rpm, 20 min, 4 °C).
Purification of DNA PrimaseAll purification procedures were conducted at 4 °C using Buffer A (50 mM Tris-HCl, pH 7.5, 50 mM KCl, 1 mM EDTA, 1 mM DTT, 10% glycerol, 1 µg/ml leupeptin), and all dialyses were performed against Buffer A.
Mitochondria (prepared by Method A from 220 g (wet weight) of cells) were lysed by gentle stirring at 4 °C for 30 min in 500 ml of mitochondrial lysis buffer (0.5 M KCl, 0.25% (v/v) Nonidet P-40, 2 mM EDTA, 10% (v/v) glycerol, 25 mM Tris-HCl, pH 7.5, 2 µg/ml leupeptin). Insoluble material was removed by centrifugation (Sorvall GSA rotor, 12,500 rpm, 30 min, 4 °C).
Crude mitochondrial lysate (Fraction 1, 500 ml) was loaded onto a 50-ml
Q-Sepharose (Pharmacia) column at a flow rate of 1 ml/min. DNA primase
activity does not bind to this column; however, this step was essential
for removal of nucleic acids in the lysate. The flow-through fraction
(Fraction 2, 450 ml) was loaded onto a 50-ml S-Sepharose (Pharmacia)
column at 2 ml/min, which was washed with 250 ml of buffer A, then with
250 ml of 0.2 M KCl in buffer A, and finally with 300 ml of
0.5 M KCl in buffer A. DNA primase activity was eluted at
the 0.5 M KCl step (Fraction 3, 300 ml). Fraction 3 was
then dialyzed and loaded onto a 30-ml double-stranded DNA-cellulose
column (Sigma) at 0.5 ml/min. The column was washed with 200 ml of
buffer A and eluted with a 240-ml gradient of 50 mM to 1 M KCl in buffer A at 0.5 ml/min. The DNA primase activity
eluted between 0.25 and 0.4 M KCl. The active fractions
were pooled (Fraction 4, 40 ml), dialyzed, and loaded onto a 1-ml Poros
HS column (PerSeptive Biosystems) at 0.5 ml/min. After washing with 20 ml of buffer A, the column was eluted with a 25-ml gradient of 50 mM to 1 M KCl in buffer A at 0.5 ml/min. The
activity was eluted at about 0.3-0.4 M KCl. Pooled active fractions (Fraction 5, 4 ml) were diluted with 4 ml of 4 M
KCl and loaded onto a phenyl-Superose column (HR5/5, Pharmacia) at 0.5 ml/min. The column was eluted with a 35-ml reverse gradient, 2 M to 0 M KCl in buffer A, at 0.5 ml/min. The
DNA primase activity eluted at about 1 M KCl. The active
fractions (Fraction 6, 1 ml) were dialyzed against buffer A containing
40% glycerol to reduce the volume to about 300 µl. The final
chromatographic procedure was gel filtration on a 30-ml Superose 12 FPLC column (Pharmacia) using buffer A at a flow rate of 0.5 ml/min.
The active fractions were pooled and dialyzed against buffer A
containing 40% glycerol (v/v) and stored at 80 °C (Fraction 7, 0.8 ml).
In a second purification, we used a mitochondrial fraction isolated by Method B. The purification steps were identical to those described in the previous paragraph with the following exceptions: 1) a 150-ml phosphocellulose column, eluted with a 1200-ml gradient from 0 to 1 M KCl in buffer A, substituted for the DNA cellulose column; 2) column sizes (except for the Poros HS column) and volumes of elution buffers were increased proportionally to the amount of protein; 3) a 10-ml phenyl-Sepharose column substituted for the phenyl-Superose, and this column was run before the Poros HS column, and 4) the final Superose 12 FPLC step was omitted because the protein was pure after Poros HS chromatography.
AntibodiesRabbit antibody to mitochondrial DNA polymerase
was a gift from Dr. Al Torri (13). Antibody to the DNA primase was
prepared by immunizing female BALB/c mice by intraperitoneal injections with purified DNA primase (Fraction 7, 2-5 µg/injection). The initial inoculations were in Freund's complete adjuvant, and four subsequent boosts were at 3-week intervals with Freund's incomplete adjuvant. The serum (at 1:1000 dilution) was screened by Western blot
for specific recognition of the 28-kDa protein. The antibody recognizes
the homogeneous primase on a Western blot (Fig. 1, lane 12;
see lane 10 for Coomassie-stained gel) and also recognizes a
polypeptide of the same size in a preparation of isolated mitochondria (Fig. 1, lane 11; see lane 9 for
Coomassie-stained gel). Preincubation of 5 µl of primase antiserum
with 5 µl of primase solution (30 min, 4 °C) resulted in 70% loss
of primase activity in the standard assay. A control experiment with
DNA polymerase
antiserum resulted in loss of only 15% of primase
activity. In an immunodepletion assay, antiserum bound to protein
A-Sepharose beads was able to deplete 74% of the primase activity. In
a control experiment with DNA polymerase
antiserum, only 10% of
the primase activity was depleted.
Immunolocalization of DNA Primase
Log phase C. fasciculata cells (2 × 107 cells/ml) were washed
with 5 mM Na2HPO4, 5 mM
NaH2PO4, 130 mM NaCl (PBS) (Sorvall
GSA rotor, 10 min, 4000 rpm, 4 °C) and resuspended in PBS at the
same cell concentration. Cell suspension (10 µl) was spotted onto
10-well slides pretreated with a 1/10 dilution of
poly-L-lysine (Sigma, 0.1%). The slides were kept in a
humidity chamber for 20 min to allow cells to adhere prior to a wash
with PBS. The cells were first fixed with 2% paraformaldehyde (2 min
at room temperature), a treatment followed by fixation in 100%
methanol (overnight at 20 °C). The fixed cells were washed with
PBS and then treated with 20 mM glycine in PBS for 10 min
to neutralize residual aldehyde groups. After washing again with PBS,
the slides were treated for 1 h at room temperature with 10%
(w/v) BSA, 0.5% (v/v) Tween 20 in PBS (blocking solution), and then
incubated in a humidity chamber for 1 h at room temperature with
mouse anti-primase serum diluted 1:250 in blocking solution. After
washing in PBS, they were incubated for 1 h at room temperature
with fluorescein isothiocyanate-conjugated goat anti-mouse IgG
(Boehringer Mannheim) diluted 1:250 in blocking solution. The slides
were washed in PBS and then in PBS with DAPI (0.1 µg/ml). After
washing with PBS for another 10 min, the slides were then mounted with
Mowiol 4-88 (Calbiochem) (14) containing p-phenylenediamine
(1 mg/ml). To double-label the cells, the primary antibody reaction (1 h, room temperature) included rabbit antibody to DNA polymerase
(1:250 dilution) and mouse antibody to DNA primase (1:250 dilution).
The secondary antibody reaction (1 h, room temperature) included
rhodamine-conjugated affinity-purified goat antibody to rabbit IgG
(Boehringer Mannheim, 1:250 dilution) and fluorescein-conjugated
affinity-purified goat antibody to mouse IgG (Boehringer Mannheim,
1:250 dilution). The slides were examined by fluorescence microscopy
using a Zeiss Axioskop microscope, and the images captured on Kodak
Tmax 400 film. The negatives were scanned with a Nikon 35-mm film
scanner (LS-1000) and processed in Adobe Photoshop on a Macintosh
computer.
Table I presents a summary of the purification. Starting from the mitochondrial lysate supernatant (prepared by Method A), the DNA primase was purified 14,500-fold and the yield of activity was about 8%. Fig. 1 shows an analysis of Fractions 1-7 by SDS-PAGE and silver staining. Fraction 7 appears to contain only a single homogeneous protein of 28 kDa.
|
The final yield of primase from mitochondria prepared by Method A was
only about 10 µg of protein (Table I). We considered the possibility
that this mitochondrial isolation procedure had a low yield, due at
least in part to the extensive washing of the mitochondrial fraction in
Method A. We therefore did a second purification using a mitochondrial
fraction isolated by Method B. In the preparation shown in Table I, we
started with only 1750 mg of mitochondrial protein in a mitochondrial
lysate prepared from 220 g of cells. Using Method B in a second
purification, we started with 27,200 mg of mitochondrial protein from
450 g of cells. Using this procedure we obtained about 300 µg of
purified DNA primase in about 10% yield. Its specific activity was
comparable to that shown in Table I. Fig. 1 (lane 8) shows
an SDS-PAGE analysis of the final product. This most pure fraction, or
the Fraction 7 enzyme in Table I, was used in all experiments described
in this paper. The purified enzyme was stored at 80 °C in buffer containing 25 mM Tris-HCl, pH 7.5, 100 mM KCl,
0.5 mM EDTA, 1 mM DTT, 40% glycerol. It
maintains over 50% of its activity after 6 months of storage.
As shown in Fig. 2, the DNA primase
activity (panel A) co-eluted from the Superose 12 column
with the 28-kDa protein as indicated by the intensity of the
silver-stained band on SDS-PAGE (panel B). Primase activity
and the 28-kDa protein both peak in fraction 31. The 28-kDa protein
also co-eluted with primase activity on the phenyl-Superose and Poros
HS columns, and the ratio of activity to silver staining of the band
was comparable to that of Fraction 7. These data provide strong
evidence that the 28-kDa protein is responsible for activity. To assess
the size of the native enzyme, we also compared its behavior in gel
filtration (Superose 12 FPLC) with that of two reference single-chain
proteins, carbonic anhydrase (29 kDa) and BSA (67 kDa). The elution
position of the DNA primase was the same as that of carbonic anhydrase
(see legend of Fig. 2A), providing strong evidence that the
primase is a 28-kDa monomer.
Requirements for DNA Primase Activity
Maximal primase activity requires ATP and a divalent cation (Mg2+) (Table II). The enzyme may have a tightly bound metal ion, as 21% of maximal activity is detectable in the absence of added Mg2+. However, EDTA inhibits activity completely when added at a concentration equivalent to that of the divalent cation. Rifampicin and N-ethylmaleimide inhibit the activity only weakly. To determine whether in our standard assay (using a poly(dT) template) the incorporation of [32P]dAMP into an acid-insoluble form depends on the activity of primase, we varied the ATP concentration from 0 to 4 mM. As shown in Fig. 3A, there was no incorporation in the absence of ATP and incorporation was maximal at 1 mM (closed symbols). In a control experiment (open symbols), there was little effect of ATP on the incorporation of [32P]dAMP when primase was omitted and oligo(dA) was used as a primer. In a related experiment we used single-stranded M13 DNA as a template (Fig. 3B). Again there was dependence upon the presence of rNTPs (compare lower line to upper line), and the magnitude of [32P]dAMP incorporation depended upon the primase concentration.
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The Products of the DNA Primase Reaction
To characterize the
products, we conducted reactions (for times up to 45 min) containing
poly(dT) template, [-32P]ATP as substrate, and DNA
primase. Analysis of the reaction products on a 20% polyacrylamide gel
revealed a ladder of fragments ranging up to about 10 nucleotides in
size (Fig. 4A). However, with
a long exposure of the autoradiogram, there were very small amounts of
larger oligonucleotides, up to about 15 nucleotides in size. We also
found in a similar experiment using M13 DNA as a template and the 4 rNTPs that the primase synthesized a very heterogeneous population of
oligonucleotides in the same size range (data not shown).
We next used a chase to determine whether the oligonucleotides
synthesized by primase can prime DNA synthesis (Fig. 4B). We first synthesized primers using [-32P]ATP as a
substrate (lane 1), and the products are the characteristic ladder of fragments up to about 10 nucleotides in size. We then added
Klenow DNA polymerase and increasing concentrations of dATP to the
reaction. As shown in lanes 2-4, the primers were elongated and the products appeared in or near the slot. The products of DNA
primase can also be elongated by the purified C. fasciculata mitochondrial DNA polymerase
(15). However, because of the low
processivity of this enzyme, the primer extensions occurred with much
lower efficiency (data not shown).
Four lines of evidence support the mitochondrial localization of this primase. First, in a small scale purification (from about 25 g of cells), we used Percoll gradient-purified mitochondria to purify the primase through the phenyl-Superose fraction. The primase obtained from the purified mitochondria had activity and chromatographic behavior similar to that obtained from the crude mitochondrial fraction.3 Second, the activity in the crude mitochondrial fraction was latent, requiring release from vesicles by treatment with a buffer containing 0.25% Nonidet P-40 and 0.5 M KCl. In a control assay in which the detergent/KCl treatment was omitted, the amount of detectable primase activity was only about 10% of that obtained with detergent/KCl.3 Third, a Western blot of proteins from a mitochondrial fraction revealed a 28-kDa polypeptide that reacted with anti-primase antibody (Fig. 1, lanes 9 and 11). Finally, immunolocalization experiments proved conclusively that the primase is localized within the single mitochondrion of this parasite.
For immunolocalization we used a polyclonal antibody against the
primase. The results indicate that the primase is indeed situated
within the mitochondrion in discrete sites near the kDNA network. Fig.
5A shows two C. fasciculata cells visualized by Nomarski optics. Fig.
5B shows the same cells stained with DAPI, which brightly
stains the kinetoplasts (the nuclei stain poorly under these conditions
and are barely visible). The kinetoplast is a disc-shaped structure,
oriented perpendicular to the flagellum, and this image shows the edges
of the discs. Fig. 5C shows the primase in the same cells,
localized above and below the kinetoplast disc. Somewhat more
fluorescence is observed on the side of the disc nearest the flagellum,
a distribution observed in many cells. We were concerned that the
images of primase and DAPI staining might have been misaligned and that
both components of the primase fluorescence may have been on the same
side of the kinetoplast disc. This possibility is highly unlikely,
given that the two cells are oriented in opposite directions and that
the distances between the DAPI stains and the primase stains in the two
cells are identical. Therefore we conclude that the primase forms a sandwich-like structure around the kinetoplast disc. DNA primase is
localized in the same positions in the majority of log phase cells
(90% or more). In another experiment, Fig. 5 (panels D-G) shows images of a single cell, which is visualized by Nomarski optics
(panel D), DAPI staining (panel E),
immunolocalization of the primase (panel F), and
immunolocalization of the mitochondrial DNA polymerase (panel
G). It is clear that the two enzymes have distinctly different
locations. The primase is situated above and below the disc
(panel F), whereas the DNA polymerase
is situated in two
protein complexes (as described previously; Ref. 9) on opposite sides
of the kinetoplast disc (panel G).
Starting with a mitochondrial fraction from C. fasciculata, we have purified a DNA primase to near homogeneity. The purified enzyme has a molecular mass of 28 kDa, and its active form is a single polypeptide chain. Assuming that the recovery of mitochondria used for purification was 100%, we can make a rough estimate that there are about 11,000 molecules of primase within the parasite's single mitochondrion. Using a poly(dT) template, the primase produces a ladder of homopolymeric products, with a maximum size of about 10 nucleotides. Using single-stranded M13 DNA as a template, the primase makes products in roughly the same size range (data not shown). However, due to sequence heterogeneity, the M13 products are not resolved into a uniform ladder by gel electrophoresis, a finding which implies that initiation on M13 occurs at multiple sites.
The primase products can be efficiently elongated by Klenow DNA
polymerase in the presence of the appropriate dNTPs. However, when
poly(dT) is used as a template, the only primers that are efficiently
extended by Klenow polymerase are the 10-mers and, to a lesser extent,
the 9-mers (see Fig. 4B, lanes 3 and
4). Shorter oligonucleotides are not elongated efficiently
by the DNA polymerase. A similar effect has been observed with
mammalian primases (16). A chase experiment, in which non-radioactive
ATP was added after synthesis of primers with
[-32P]ATP, did not result in the shorter
oligonucleotides being extended (data not shown). It is possible that
many of the shorter oligonucleotides had dissociated from the dT
template and therefore could not serve as intermediates in primer
synthesis. In another experiment, we found that the primers were poorly
extended by the C. fasciculata mitochondrial DNA polymerase
, a result not surprising given that that enzyme may not be the
major replicative enzyme in this organelle (see below).
Intracellular enzyme localization studies have been valuable in
clarifying our understanding of kDNA replication. The C. fasciculata kinetoplast system is ideal for these studies because
the non-replicating cell has only one kDNA network, which resides
within its single mitochondrion. The network in vivo is
condensed into a characteristic disc-shaped structure, about 1 µm in
diameter and 0.4 µm thick, well within the resolution of fluorescence
microscopy. Previous immunolocalization studies had demonstrated that a
mitochondrial topoisomerase II (8) and a mitochondrial DNA polymerase
(9) co-localize within two protein complexes, which are situated in antipodal positions adjacent to the kinetoplast disc (see localization of DNA polymerase
in Fig. 5G). Based on the presence of
two enzymes involved in replication, and other evidence (9), we had
hypothesized that these two protein complexes are involved in
minicircle replication. Another topoisomerase II (7), as well as some
histone-like DNA binding proteins (17), localize within the kinetoplast
disc. An hsp70 heat shock protein surrounds but does not appear to
penetrate the kinetoplast disc (18, 19). We now report that the primase
has a distinct localization, being situated both above and below the
kinetoplast disc. Little if any appears to flank the edge of the disc
or is associated with the two complexes containing DNA polymerase
and topoisomerase II. We have not yet determined whether the primase
covers the entire upper and lower surfaces of the kinetoplast disc or
if it covers only part of these surfaces. We also do not know if the
enzyme is completely excluded from the kinetoplast disc or if it
penetrates the upper and lower regions of the disc. Studies at higher
resolution, involving confocal fluorescence microscopy or
immunoelectron microscopy, will be needed to address these issues.
What is the significance of the primase localization to the mechanism
of minicircle replication? A current view of this mechanism is shown in
Fig. 6. The diagram shows a section
through the kinetoplast disc, with interlocked minicircles forming a
monolayer (see Ref. 3 for further information about this arrangement
and about the replication scheme). Covalently closed minicircles are
released from the central region of the network by a topoisomerase II, which is situated within the disc. Ultimately these free minicircles migrate to one of the two protein complexes (containing topoisomerase II and DNA polymerase ) that flank the kinetoplast disc. The newly
replicated progeny minicircles, containing gaps, are attached to the
network periphery adjacent to these protein complexes, in another
topoisomerase reaction (20). The localization of the primase above and
below the disc raises the possibility that replication may not actually
occur within the antipodal protein complexes. Instead, minicircle
replication could occur in the region above or below the kinetoplast
disc. A replicative DNA polymerase in this location (not yet discovered
but possibly related to DNA polymerase
) could have the major
responsibility for DNA synthesis. The minicircle progeny could then
migrate to one of the two protein complexes where many of the gaps in
the discontinuously synthesized strand could be repaired immediately
prior to network attachment (21). The DNA polymerase
is ideally
situated to carry out this reaction, and it is known to have a
preference for gap filling. The progeny minicircles could then be
attached to the network periphery by topoisomerase II. In an
alternative scheme, the covalently closed minicircle, immediately after
release from the network, could associate with primase and other
proteins to form a replication initiation complex. It could then
migrate to one of the two antipodal protein complexes to complete its replication. Further studies, including the localization of additional replication enzymes, will be needed to distinguish between these possibilities.
We thank members of our laboratory for many enlightening discussions, consistent support, and comments on the manuscript.