of 21
Substrate Binding Regulates Redox Signaling in Human DNA
Primase
Elizabeth O’Brien
†,‡
,
Marilyn E. Holt
§
,
Lauren E. Salay
§
,
Walter J. Chazin
*,§
, and
Jacqueline
K. Barton
*,†
Division of Chemistry and Chemical Engineering, California Institute of Technology, Pasadena,
California 91125, United States
§
Departments of Biochemistry and Chemistry, Center for Structural Biology, Vanderbilt University,
Nashville, Tennessee 37240, United States
Abstract
Generation of daughter strands during DNA replication requires the action of DNA primase to
synthesize an initial short RNA primer on the single-stranded DNA template. Primase is a
heterodimeric enzyme containing two domains whose activity must be coordinated during primer
synthesis: an RNA polymerase domain in the small subunit (p48) and a [4Fe4S] cluster-containing
C-terminal domain of the large subunit (p58C). Here we examine the redox switching properties of
the [4Fe4S] cluster in the full p48/p58 heterodimer using DNA electrochemistry. Unlike with
isolated p58C, robust redox signaling in the primase heterodimer requires binding of both DNA
and NTPs; NTP binding shifts the p48/p58 cluster redox potential into the physiological range,
generating a signal near 160 mV vs NHE. Preloading of primase with NTPs enhances catalytic
activity on primed DNA, suggesting that primase configurations promoting activity are more
highly populated in the NTP-bound protein. We propose that p48/p58 binding of anionic DNA and
NTPs affects the redox properties of the [4Fe4S] cluster; this electrostatic change is likely
influenced by the alignment of primase subunits during activity because the configuration affects
the [4Fe4S] cluster environment and coupling to DNA bases for redox signaling. Thus, both
binding of polyanionic substrates and configurational dynamics appear to influence [4Fe4S] redox
signaling properties. These results suggest that these factors should be considered generally in
characterizing signaling networks of large, multisubunit DNA-processing [4Fe4S] enzymes.
Graphical Abstract
*
Corresponding Authors: jkbarton@caltech.edu, walter.j.chazin@vanderbilt.edu.
Present Address: Department of Chemistry, University of California, Berkeley, Berkeley, CA 94720
The authors declare no competing financial interest.
Supporting Information
The Supporting Information is available free of charge on the
ACS Publications website
at DOI:
10.1021/jacs.8b09914
.
Electrochemistry of electrochemically unaltered wild-type p48/p58, UV−visible spectra of wild-type and mutant p48/p58, additional
biochemistry experiments assessing the role of substrate binding order on primase initiation, an electrostatic map of the primase/p58C
DNA binding interface, and a table of DNA substrates used in the experiments (
PDF
)
HHS Public Access
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J Am Chem Soc
. Author manuscript; available in PMC 2019 April 18.
Published in final edited form as:
J Am Chem Soc
. 2018 December 12; 140(49): 17153–17162. doi:10.1021/jacs.8b09914.
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INTRODUCTION
Dynamic, coordinated DNA replication in eukaryotic cells must efficiently duplicate
genomes on the scale of 10
6
−10
9
base pairs with error rates of approximately 10
−9
mismatches per replication cycle.
1
The enzyme responsible for initiating daughter strand
synthesis in eukaryotes is a heterodimeric DNA-dependent RNA polymerase, DNA primase.
2
6
Primase is a heterodimer containing an RNA polymerase subunit, p48, and a regulatory
subunit, p58.
7
10
Primase exists in complex with the heterodimeric DNA polymerase
α
(pol-
prim), which functions to generate the RNA−DNA primer required for the bulk of genome
duplication by processive DNA polymerases
ε
and
δ
.
3
6
,
11
13
Primase binds the ssDNA
template, along with two nucleotide triphosphates (NTPs) and two catalytic metals, to
synthesize the first dinucleotide (nt). After dinucleotide synthesis, primase rapidly elongates
the primer to appropriate length (8−14 nts), before handing the primer off to polymerase
α
.
Polymerase
α
then adds 10−20 deoxynucleotide triphosphates (dNTPs) downstream of the
initial RNA primer before handing off the substrate to one of the processive DNA
polymerases to complete genome duplication.
All eukaryotic replicative polymerases, as well as the translesion synthesis B-family
polymerase
ζ
,
14
16
are reported to contain [4Fe4S] clusters, which are metabolically
expensive cofactors associated with biological redox chemistry.
17
,
18
The clusters in both
DNA polymerase
δ
19
and the [4Fe4S] domain of primase, p58C,
20
can be oxidized and
reduced when bound to DNA through DNA-mediated charge transport (DNA CT). DNA CT
is a long-range, rapid, and mismatch-sensitive biochemical process,
21
23
making it
interesting to consider as a regulatory mechanism for replication. Moreover, experiments
support the application of this chemistry in locating oxidative lesions
in vitro
and in
cells
24
26
by DNA repair enzymes containing [4Fe4S] clusters.
The p58C domain of human and yeast DNA primase has been demonstrated
electrochemically to undergo DNA-mediated redox switching between an oxidized,
[4Fe4S]
3+
state and a resting, [4Fe4S]
2+
redox state.
20
,
27
The oxidized [4Fe4S]
3+
p58C is
tightly bound to DNA, whereas the reduced [4Fe4S]
2+
p58C is only loosely bound to DNA.
A similar switch is evident in the base excision repair protein, Endonuclease III, which
contains a [4Fe4S] cluster; here binding to the DNA polyanion shifts the [4Fe4S]
3+/2+
redox
potential so that the oxidized [4Fe4S]
3+
form binds DNA 500-fold more tightly than the
[4Fe4S]
2+
form.
28
,
29
This redox switch in p58C appears to regulate primase activity and
primer product distribution, yet the electrochemical behavior of the full primase enzyme,
relative to that of p58C, has yet to be reported. Since both the catalytic subunit (p48) and the
regulatory subunit containing the [4Fe4S] cluster (p58) of primase are required for binding
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of DNA and NTPs to initiate primer synthesis,
7
10
,
30
,
31
the electrochemical behavior of the
complete p48/p58 heterodimer is likely an important element of the mechanism driving
primer synthesis.
The p48 and p58C domains must be positioned near one another during priming, so that
both can contact the template DNA; however, the DNA binding interfaces of p48 and p58C
are separated by approximately 60 Å in the crystal structure of free primase.
7
These
structural data suggest that primase must undergo a significant configurational
rearrangement when binding DNA and NTPs, specifically a subunit realignment to position
p58C over the p48 active site to begin primer synthesis. Beyond configurational
rearrangement, moreover, the electrostatic environment of the [4Fe4S] cluster cofactor in
primase is likely changed dramatically by the binding of the anionic DNA and NTP
substrates, as it is for Endonuclease III. Because electrostatic environment is a major factor
in shifting the redox potential of [4Fe4S] cluster cofactors,
32
it is reasonable to consider that
binding of DNA and NTPs may change the redox behavior of the full-length p48/p58
primase enzyme. Characterization of both the electrostatic effect of DNA and NTP binding
on primase [4Fe4S] cluster redox behavior and the configurational rearrangement of primase
upon substrate binding, which positions the RNA polymerase and [4Fe4S] domains near the
DNA, close to one another, is crucial for understanding the biochemical factors coordinating
primase activity, which eventually culminates with primer termination and handoff to
polymerase
α
.
Here we use DNA electrochemistry to investigate the electrostatic effects of DNA/NTP
binding on primase redox activity. We find that full-length human primase displays a small
amount of redox signaling upon electrochemical oxidation in the presence of DNA, though
oxidation in the presence of DNA alone is a more dramatic redox switch for the isolated
p58C domain.
20
When bound to both DNA and NTPs, however, p48/p58 displays robust,
semireversible redox activity. Binding of anionic substrates changes the electrostatic
environment of the primase [4Fe4S] cluster, shifting the potential into signaling range with
other [4Fe4S] enzymes. Moreover, alignment of p58C over the active site of p48 should also
alter the electrostatic field of the cluster. Preloading p48/p58 with NTPs, additionally,
enhances catalytic activity on an exogenously primed substrate. In addition to the shift in
electrostatic environment of the cluster, preloading NTPs should promote formation of the
active primase initiation complex, with p58C and p48 proximal to the DNA template and
NTPs bound in both the 3
and 5
sites.
Thus, the redox switch driven by the primase [4Fe4S] cluster is activated upon DNA and
NTP binding, and combined with changes in configuration, regulates multistep RNA
synthesis.
MATERIALS AND METHODS
Protein Expression and Purification.
Full-length primase was expressed and purified as described previously.
33
In short, plasmid
DNA was transformed into BL21RIL (DE3) cells (Novagen), and cultured at 37 °C to an
OD
600
of 0.6, when flasks were transferred to an 18 °C incubator with shaking. After 30
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min, protein expression was induced through addition of isopropyl 1-thio-
β
-
D
-
galactopyranoside to a final concentration of 5 mM. Media was also supplemented at this
time with ferric citrate and ammonium ferrous citrate to a final concentration of 0.1 mg/mL.
The primase subunits were expressed at 18 °C for 18 h prior to harvesting and freezing at
−80 °C. After lysis, primase was first purified by nickel affinity chromatography (Amersham
Biosciences). The 6xHis tag on p48 was the cleaved with H3C protease and primase
dialyzed into a low-imidazole buffer.
33
Primase was then repassed over the nickel column to
remove the H3C protease and uncleaved protein. A heparin column was used to remove
residual contaminants before passing over a Sephadex S200 sizing column to remove
aggregates and buffer exchanged into electrochemistry storage buffer: 20 mM TRIS, pH 7.2,
150 mM NaCl, 5% glycerol.
33
Oligonucleotide Preparation.
All standard or modified phosphoramidites and DNA synthesis reagents were purchased
from Glen Research. Unmodified DNA oligonucleotides for electro-chemical experiments
were purchased from Integrated DNA Technologies, Inc. Thiol-modified DNA strands for
electrochemistry were made on an Applied Biosystems 3400 DNA synthesizer, with a C6 S
−S phosphoramidite incorporated at the 5
-terminus. Single-stranded DNA was purified
using standard procedures as described previously.
34
,
35
High pressure liquid
chromatography (HPLC) using a reverse-phase PLRP-S column (Agilent) was used, and
oligonucleo-tide mass confirmed using MALDI-TOF mass spectrometry. Thiolmodified
strands were reduced after the initial HPLC purification with 100 mM dithiothreitol (Sigma)
for 2−3 h in 50 mM Tris−HCl, pH 8.4, 50 mM NaCl. Reduced thiol-modified DNA was
purified by size exclusion chromatography (Nap5 Sephadex G-25, GE Healthcare) and
subsequent reverse-phase HPLC. Single-stranded oligonucleo-tides were then desalted using
ethanol precipitation and stored in low salt buffer (5 mM Phosphate, pH 7.0, 50 mM NaCl).
Duplex DNA for electrochemistry was prepared by quantification of the complementary
single-stranded oligonucleotides by UV−visible spectroscopy, followed by annealing at
90 °C. A mixture of equimolar complementary single-stranded DNA (50
μ
M) was prepared
in low salt buffer. Thiol-modified duplex DNA substrates were then deoxygenated by
bubbling argon gas through the solution for 90−180 s. Duplex DNA was annealed on a
thermocycler (Beckman Instruments) by initial heating to 90 °C, followed by slow cooling
to 4 °C over 90 min. DNA was quantified using absorbance at 260 nm, with extinction
coefficients at 260 nm for DNA obtained using Integrated DNA Technologies online
OligoAnalyzer tool. Single-stranded DNA substrates were quantified using UV−visible
spectroscopy and stored in low salt buffer at a stock concentration for activity assays.
Multiplexed Chip Fabrication.
Multiplexed electrode platforms were prepared using standard photolithography techniques,
adapted from established protocols.
23
,
34
,
35
Nine 1 in. by 1 in. chips were patterned on 525
μ
m thick silicon wafers (SiliconQuest). A thermal oxide layer roughly 4000 Å thick was
grown on the silicon wafers using a Tytan tube furnace (Tystar). S1813 photoresist (2
μ
m
layer) was deposited onto the wafers for patterning of the chips before metal deposition.
Electron beam evaporation (CHA Industries) was then used to deposit a 3 nm titanium
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adhesion layer followed by a 100 nm gold layer, without breaking vacuum between
depositions.
Metal lift-off using Remover PG (MicroChem) was performed overnight (10−12 h) at
ambient temperature. Wafers were subsequently dried with a nitrogen gun and dehydrated at
140 °C for 10 min. A 3
μ
m layer of insulating SU-8 photoresist was deposited and patterned
onto the wafer as described previously,
23
,
34
,
35
with connective wires between contact pads
on the edges of the chips and working electrodes in the center were covered but the contact
pads and working electrodes left exposed. This ensured a fixed working electrode surface
area of 2 mm
2
. SU-8 photoresist was cured (150 °C, 15 min) and wafers cleaved into
individual chips using a Dynatex Scriber/Breaker or broken manually after scoring with a
diamond tip scriber.
DNA-Modified Electrode Assembly/Preparation.
Multiplexed chips were cleaned using sonication in acetone and isopropyl alcohol as
described previously.
34
Chips were then dried thoroughly using argon gas and ozone-cleaned
for 20 min at 20 mW using a Uvo brand ozone cleaner. Clean chips were assembled onto
polycarbonate holders with acrylic clamp and Buna-N rubber gasket according to previous
protocols, with four quadrants in the chip separated by fastened gasket and clamp.
34
Duplex
DNA substrates, with a thiol modifier at the 5
end, (25
μ
M) were deposited in a 20
μ
L
volume onto each quadrant of the multiplex chip. Substrates incubated for 18−24 h on the
gold surface to allow formation of self-assembled DNA monolayer. DNA monolayers were
washed with phosphate buffer (5 mM phosphate, pH 7.0, 50 mM NaCl, 5% glycerol) and
subsequently backfilled with 1 mM 6-mercaptohexanol (Sigma) in phosphate buffer for 45
min. Monolayers are then washed 10 times per quadrant with phosphate buffer and twice per
quadrant with TBP buffer (5 mM phosphate, pH 7.0, 50 mM NaCl, 4 mM MgCl
2
, 4 mM
spermidine) to aid in formation of a monolayer with termini accessible for p58C binding.
Assembled chips were transported into an anaerobic glovebag chamber (Coy Products) and
washed 5 times per quadrant with p58C buffer (20 mM HEPES, pH 7.2, 75 mM NaCl),
which was previously deoxygenated by argon bubbling (at least 1 s/
μ
L of solution) and
allowed to incubate at least 1−2 days in the chamber prior to the experiment.
Initial cyclic voltammetry scans of the monolayers in p58C buffer were performed to ensure
monolayer formation on each electrode. All washes were performed with 20
μ
L buffer
volumes on each quadrant. Before scanning, a 200
μ
L volume was deposited over the chip
surface, a bulk solution well for completion of a three-electrode circuit with an external
reference and counter electrode.
Sample Preparation for Electrochemistry.
Samples were stored prior to experiments in p48/p58 storage buffer (20 mM Tris, pH 7.2,
150 mM NaCl, 5% glycerol). All p48/p58 samples were transferred to HEPES
electrochemistry buffer (20 mM HEPES, pH 7.2, 150 mM NaCl, 5% glycerol) using
Amicon ultra centrifugal filters (0.5 mL, 3 kDa MWCO) (Millipore Sigma). Catalytic metals
were not included in the electrochemistry buffer so as to prevent NTP polymerization during
electrochemistry experiments. Protein was applied in a 90−140
μ
L volume to the filter and
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centrifuged for 15 min at 14000g at 4 °C. After centrifugation, 400
μ
L of HEPES
electrochemistry buffer was applied to the filter and centrifuged at 14000g for 20 min. This
procedure was repeated four times to exchange the p48/p58 protein into HEPES
electrochemistry buffer. After buffer exchange and recovery of sample by centrifugation (2
min, 1000g), the concentration of [4Fe4S] cluster-containing protein and loading of the
[4Fe4S] cluster were assessed using UV−visible spectroscopy, by absorbance of the [4Fe4S]
cluster at 410 nm (extinction coefficient = 17 000 M
−1
cm
−1
) (see Figure S2).
36
Recovered
samples (approximately 100−150
μ
L volume) were deoxygenated for 2−3 min with argon.
Samples were then transferred into the anaerobic chamber (Coy Laboratory products).
Before deposition onto the gold electrode surface, p48/p58 samples were diluted to a molar
concentration of 5
μ
M or 7.5 μM [4Fe4S] p48/p58 with previously deoxygenated HEPES
electrochemistry buffer. Samples were deposited onto multiplex chip quadrants in 20
μ
L
volumes initially, with the remaining sample deposited in a well of bulk solution above the
chip surface, to a final volume of 200−300
μ
L.
Wild Type Human p48/p58 Electrochemistry.
All electro-chemistry was performed using a CHI620D potentiostat and 16-channel
multiplexer (CH Instruments), in an anaerobic glove chamber. Multiplex gold electrodes
were part of a three-electrode system with an external Ag/AgCl reference electrode
(Bioanalytical Systems) and platinum counter electrode. Cyclic voltammetry scans were
performed at 100 mV/s scan rates, over a potential range of +0.412 to −0.288 V vs NHE or
+512 to −188 mV vs NHE. Bulk electrolysis on DNA was performed at an applied potential
of +0.512 V vs NHE for all electrochemical oxidation reactions and −0.188 V vs NHE for
all electrochemical reduction reactions. The oxidizing potential was applied for at least 8.33
min for single oxidation reactions on a surface. The reducing potential was applied for 8.33
min in all electrochemical reduction reactions. All bulk electrolysis and cyclic voltammetry
was performed in previously deoxygenated p48/p58 storage buffer (20 mM HEPES, pH 7.2,
150 mM NaCl, 5% glycerol). Charge transfer (nC) in the cathodic peak of CV scans for
oxidized samples was assessed using the area under the current wave of the reduction signal.
Charge transfer was measured for oxidized samples using CHI software, assessing the area
under the reductive peak in CV after electrochemical oxidation. Yields for bulk electrolysis
were assessed by subtracting the total charge reported in coulombs from the product of the
electrolysis time (s) and the final current value (A). NTP-dependence of electrochemical
signals were measured by pipetting a small volume (1−3
μ
L) of 0.1 M ATP stock solution
into each quadrant of the multiplexed chip setup.
Samples were added by quadrant, as physical barriers in the setup prevent diffusion of NTPs
between electrode quadrants. After the volume of ATP stock was deposited onto the
electrode quadrant, resulting in a 3.3 mM concentration of ATP in the quadrant, CV scans
were measured (100 mV/s scan rate). Charge transfer was assessed using CHI software;
charge values were determined by calculation of the area under the reductive and oxidative
peak curves. Midpoint potentials of NTP-dependent redox signals were assessed using the
peak selection function in CHI software.
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Primase Preincubation Initiation Assays.
All primase assays were performed anaerobically, with deoxygenated buffers and reagents.
Primase was preincubated at ambient temperature, either in a stock alone, with the DNA, or
with the NTPs used in the reaction. The preincubation conditions were 30 min at ambient
temperature; only DNA or NTPs, not both substrates at once, were incubated with each
sample of primase. The
α
32
P ATP was initially dried (2.5
μ
L of 12
μ
M, eventually diluted
to 30
μ
L)
in vacuo
overnight the day preceding the reactions. The
32
P ATP-containing tubes
were then brought into the anaerobic glovebag chamber, along with concentrated stocks of
the unlabeled NTPs, CTP and UTP, and the single-stranded 50-nt initiation substrate, shown
in Table S1. Preincubation mixtures consisted of the following: protein-only preincubation
samples contained 800 nM p48/p58 in primase activity buffer (50 mM Tris, pH 8.0, 5 mM
MgCl
2
), paired with a sample of 500 nM ssDNA and 376
μ
M UTP, 224
μ
M CTP, and 2
μ
M
α
32
P ATP in activity buffer; DNA/protein preincubation samples contained 800 nM
p48/p58 and 500 nM ssDNA and were paired with a sample containing 376
μ
M UTP, 224
μ
M CTP, and 2
μ
M
α
32
P ATP in primase activity buffer; NTP and protein preincubation
tubes consisted of a sample containing 800 nM p48/p58 with 376
μ
M UTP, 224
μ
M CTP,
and 2
μ
M
α
32
P ATP and paired with a sample containing 500 nM ssDNA, all in the Tris
activity buffer. These samples were all incubated anaerobically for 30 min at ambient
temperature. The volume of reagents in each of the initial two tubes was 15
μ
L for each
reaction, making a total reaction of 30
μ
L when combined and incubated at 37 °C. The final
reaction conditions were 400 nM [4Fe4S] p48/p58, 112
μ
M CTP, 188
μ
M UTP, 1
μ
M
α
32
P
ATP, 250 nM ssDNA in 50 mM Tris, pH 8.0, 5 mM MgCl
2
. The primase reactions were
incubated for 1, 3, 5, 10, and 30 min at 37 °C in anaerobic conditions, and then quenched by
an equal volume per 5.5
μ
L reaction aliquot of 1% SDS, 25 mM EDTA quenching solution
to stop the reaction. Reactions, when quenched, were then transported out of the anaerobic
chamber and heat-denatured for 10 min at 70 °C, aerobically. Finally, to remove the excess
free
32
P-labeled nucleotide, the samples were each passed through spin columns (Mini
Quick Spin Oligo Columns, Roche) according to manufacturer’s protocols, to separate
unincorporated radioactivity from small products made during primase initiation (7−10 nt).
Samples were then scintillation-counted and dried overnight
in vacuo
. The samples were
then separated using 20% polyacrylamide gel electrophoresis (denaturing gel). Gels were
warmed at 1700−2000 V (90 W) for approximately 1.5 h before loading samples. Samples
were resuspended after drying in 2
μ
L of formamide loading dye, vortexed, centrifuged, and
heated at 90 °C for 1 min. They were then loaded onto the gel and run at ~2000 V (90 W)
for 3.5 h. Gels were then exposed to a phosphor screen (GE Healthcare) for 14 h and imaged
on a Typhoon 9000 Phosphorimager (GE Healthcare). Products were quantified using
ImageQuant TL software; reported numbers are mean ± SD values for
n
= 3 trials.
Preincubation Reactions: Elongation.
For primase elongation reactions, preincubation, reaction, and purification conditions were
generally similar to those of initiation assays. Reagents were prepared in essentially the
same manner as for initiation. 2.5
μ
L of 12
μ
M
α
32
P ATP was dried
in vacuo
overnight for
each elongation reaction, then transported into the anaerobic chamber. Preincubation
mixtures were prepared similarly to those used in initiation assays; two 15 μL fractions of
reagents in various combinations were prepared for each reaction and allowed to incubate in
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the anaerobic chamber for 30 min at ambient temperature before being mixed and reacted at
37 °C. The primase only preincubation samples consisted of 800 nM p48/p58 and a paired
sample of 1
μ
M dsRNA/DNA and 360
μ
M UTP, 240
μ
M CTP, and 2
μ
M
α
32
P ATP all in
primase activity buffer; DNA/protein preincubation samples consisted of 800 nM p48/p58
and 1
μ
M dsRNA/DNA and a paired sample of 360
μ
M UTP, 240
μ
M CTP, and 2
μ
M
α
32
P
ATP all in primase activity buffer; NTP/primase preincubation samples contained 800 nM
p48/p58 with 360
μ
M UTP, 240
μ
M CTP, and 2
μ
M
α
32
P ATP and a paired sample of 1
μ
M dsRNA/DNA, all in primase activity buffer. The final reaction conditions, consequently,
were 400 nM p48/p58, 120
μ
M CTP, 180
μ
M UTP, 1
μ
M
α
32
P ATP and 500 nM 2
-OMe
RNA-primed DNA in 50 mM Tris, pH 8.0, 5 mM MgCl
2
. After preincubation and mixing
for reaction, each primase assay was incubated anaerobically at 37 °C and aliquots of the
reaction chemically quenched at 1, 3, 5, 10, and 30 min of reaction time. The chemical
quencher for each 5.5
μ
L aliquot of reaction was an equal volume of 1% SDS, 25 mM
EDTA, administered anaerobically. Reactions were further aerobically heat-denatured at
70 °C for 10 min. Elongation reactions were purified initially using Mini Quick Spin Oligo
Columns (Roche) and then using
P
6
Micro Bio Spin Columns (BioRad). The Roche
columns retain all synthesized products; the initiation products 7−10 nt on the ssDNA
segment of the substrate oligonucleotide are purified and quantified with elongation
products. The BioRad spin columns have an exclusion limit of 6 kDa, or approximately 20
bases, thus eliminating short initiation products from the quantified/purified mixture; this
separation allows for comparison of truncated (30−35 nt) products and elongated products
(60 nt) in the elongation assays.
RESULTS
Human DNA Primase Redox Activity on DNA.
We first sought to investigate whether human primase participates in redox signaling when
bound to DNA. Using multiplexed DNA-modified electrodes, we electrochemically
monitored the redox activity of WT human DNA primase (p48/p58) using cyclic
voltammetry (CV) (Figure 1, Figure 2). Using a 36-mer duplex DNA substrate with a 9-nt,
5
-ssDNA overhang (Table S1) to accommodate the DNA footprint of primase,
37
we
initially observed that electrochemically unaltered WT primase, which is largely in the
[4Fe4S]
2+
redox state,
20
does not participate in DNA-mediated redox signaling (Figure 2,
S1). This behavior is similar to electrochemically unaltered human and yeast p58C
20
,
27
on
DNA, confirming that the isolated, [4Fe4S]
2+
primase enzyme is redox-inert. When we scan
the unaltered primase enzyme using square wave voltammetry (SWV), a more sensitive
electrochemical technique that minimizes background current, however, an irreversible
cathodic peak at −64 ± 7 mV vs NHE is observed. Oxidation of [4Fe4S] proteins, like
primase, from the resting [4Fe4S]
2+
state to the [4Fe4S]
3+
state can lead to further oxidation
with degradation of the oxidized cluster to the [3Fe4S]
+
degradation product.
24
We have,
moreover, observed the [3Fe4S]
+
species on the DNA electrodes for mutants of both yeast
p58C
27
and a human base excision repair enzyme MUTYH.
38
The samples of p48/p58 were
exposed to atmospheric oxygen during purification and preparation for electrochemistry, and
this peak potential is consistent with values expected for the irreversible [3Fe4S]
+/0
reduction reaction,
38
so we assign the signal to trace amounts of [3Fe4S]
+
product formed in
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the primase protein sample (Figure 2). While we observe a residual amount of [3Fe4S]
+
protein from oxidative damage during aerobic protein preparation, we do not assign this to
an electrochemically induced effect. In the absence of oxygen, wild-type [4Fe4S] primase,
similarly to p58C, DNA polymerase
δ
, human MUTYH, and Endonuclease III,
19
,
20
,
27
,
28
,
38
is stable and can be electrochemically cycled within this mild, physiological potential
regime repeatedly during cyclic voltammetry.
We next electrochemically oxidized (
E
applied
= 512 mV vs NHE) or electrochemically
reduced (
E
applied
= −188 mV vs NHE) a sample of 7.5
μ
M [4Fe4S] DNA primase on an
electrode surface using bulk electrolysis. Strict anaerobic conditions ensured full control
over the redox state of the protein.
39
Subsequent cyclic voltammetry (CV) scans over
physiological potentials (Figure 2) show a small reductive peak on the order of ~1 nC charge
transport near −90 mV vs NHE in the oxidized sample. This peak essentially disappears
after the first scan to negative, reducing potentials. Interestingly, this small redox signal is
also observed in electrochemically reduced primase, though it is smaller than the signal in
the initial scan of oxidized primase. In contrast, the CV of the p58C domain indicates a large
reductive peak in the oxidized sample, but no measurable redox activity in the reduced
sample (Figure 2). Hence, only the primase heterodimer is capable of supporting complete
redox cycles. Because the reductive peak in the oxidized primase sample disappears after
one scan to reductive potentials and is at a different redox potential, it is not indicative of the
cluster degradation product [3Fe4S]
+
and its subsequent reduction to the [3Fe4S]
0
species.
Thus, human primase can participate in redox signaling to some degree in the presence of
DNA only, but additional factors are necessary to observe robust redox switching between
[4Fe4S]
2+
and [4Fe4S]
3+
oxidation states.
DNA-Mediated, NTP-Dependent Redox Signaling in Human Primase.
The relatively small redox signal for oxidized [4Fe4S]
3+
primase generated on a DNA
electrode (Figure 2), relative to that observed for the isolated p58C domain, indicates that
their environments are distinctly different. One important point is that the primase
heterodimer binds DNA more tightly than either p48 or p58C in both human
20
,
40
(Figure 3)
and yeast
9
primase. To verify that this is the case for our preparations, we measured DNA
binding of human primase and isolated p58C using fluorescence anisotropy, and found that
reduced full-length primase (p48/p58) binds DNA with
K
d
= 0.30 ± 0.03 μM, whereas
reduced p58C binds ~20-fold more weakly with
K
d
= 5.5 ± 0.5
μ
M.
20
We next investigated whether binding of NTPs in addition to DNA would promote redox
signaling. An NTP pool present on the electrode surface allows sampling of catalytically
relevant configurations by primase, while the absence of catalytic metals prevents
polymerization. Divalent Mg
2+
ions in millimolar concentrations moreover can coat the
DNA on the electrode surface and occlude primase binding and redox signal generation.
Upon incubating 5
μ
M p48/p58 on a DNA electrode with 3.3 mM [ATP+CTP], to promote
NTP binding at the ss/dsDNA junction of the substrate conjugated to the Au surface, we
observe that primase consistently displays a robust, semireversible redox signal. (Figure 3)
The NTP-dependent CV signal for human DNA primase is attenuated in the presence of an
abasic site in the DNA duplex (14 ± 2 nC charge transfer in the cathodic peak for well-
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matched DNA versus 9 ± 4 nC charge transfer in the cathodic peak for abasic site-containing
DNA), consistent with our previous results showing the signal is DNA-mediated.
20
(Figure
3) In our HEPES electrochemistry buffer (20 mM HEPES, pH 7.2, 150 mM NaCl, 5%
glycerol), the signal is centered near 160 mV vs NHE. This signal is within the range
expected for DNA-processing [4Fe4S] enzymes cycling between the [4Fe4S]
2+
and
[4Fe4S]
3+
states,
19
,
20
,
24
29
and is similar to the reported values for human and yeast
p58C
20
,
27
in the presence of DNA and NTPs.
The midpoint potential of primase in the presence of DNA and NTPs (160 ± 4 mV vs NHE)
is slightly higher than the midpoint potentials observed for human and yeast p58C in the
presence of DNA and NTPs, which is near 150 mV vs NHE.
20
,
27
This shift may be due to an
increased amount of insulating protein matrix surrounding full-length primase as compared
to p58C, which promotes a higher reduction potential.
32
Binding of the DNA polyanion and
negatively charged NTPs, importantly, still shifts the cluster potential of full-length primase
into the physiological range for signaling activity. This result suggests that unlike p58C, the
redox switch allows primase to cycle between the [4Fe4S]
3+
state and the [4Fe4S]
2+
state,
presumably because primase remains associated with both DNA and NTPs. In the
enzymatically competent form, primase readily participates in DNA-mediated redox
signaling.
The NTP-dependent electrochemical signal observed for p48/p58 demonstrates that primase
can readily undergo a redox switch driven by the [4Fe4S] cluster cofactor upon forming an
initiation complex with bound DNA and NTPs. Structural and biochemical evidence
7
,
40
,
41
suggest that this redox switch is accompanied by a realignment of the subunits within the
p48/p58 heterodimer. The X-ray crystal structure of free primase in the absence of substrates
shows it adopts an “open” conformation
7
with the RNA polymerase domain and the p58C
domain ~60 Å apart. However, both the p48 and p58C domains of DNA primase contribute
to binding of the DNA and two NTPs necessary to form the initiation complex.
13
,
40
42
The
primase heterodimer must therefore undergo a configurational reorientation so that the p58C
domain is positioned over the DNA template and the p48 catalytic site in order for priming
to occur. The shift from the open configuration of DNA primase may be critical to both
proper alignment of critical domains, NTPs and DNA template, and also the change in the
electrostatic environment of the cluster in p58C.
Effects of NTP and DNA on Initiation and Elongation.
To further assess the effect of DNA template and NTP binding on initiation and elongation
activity, we preloaded p48/p58 with either DNA or NTPs under anaerobic conditions, then
measured polymerase activity during
in vitro
primer initiation and elongation. WT p48/p58
was first preincubated with template DNA (ssDNA for initiation, dsRNA/DNA for
elongation, Table S1) or NTPs for 30 min in an anaerobic chamber at ambient temperature.
Reactions were then begun by adding the remaining necessary substrates for activity to each
sample and incubating the mixtures at 37 °C. All preincubation samples contained the same
concentration of primase in the same total volume (15
μ
L). The final initiation reaction
conditions were 400 nM [4Fe4S] p48/p58, 112
μ
M CTP, 188
μ
M UTP, 1
μ
M
α
32
P ATP,
250 nM ssDNA (initiation substrate in Table S1) in 50 mM Tris, pH 8.0, 5 mM MgCl
2
, and
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the final elongation reaction conditions were 320 nM p48/p58, 500 nM primed DNA
(elongation substrate in Table S1), 180 μM [UTP], 120
μ
M [CTP], 1
μ
M
α
32
P ATP in 50
mM Tris, pH 8.0, 5 mM MgCl
2
. We measured the products synthesized after quenching
reaction mixtures at
t
= 1, 3, 5, 10, and 30 min.
We observe that preloading of primase with template DNA or NTPs does not enhance
de
novo
primer synthesis on ssDNA (Figure S3, Figure S4). Both total products and primer-
length (7−10 nt) products were quantified and normalized to primase-only incubation
conditions. Levels of primer synthesis were indistinguishable across all preincubation
conditions; loading primase with ssDNA or NTPs before the priming reaction did not confer
any advantage. These data contrast, interestingly, with the previously observed effect
reported by Sheaff et al., who observed no activity when DNA primase is incubated
aerobically on a poly(dT) substrate with ATP.
12
We suspect that the presence of atmospheric
oxygen may have nonspecifically oxidized ATP-bound p48/p58, thereby inhibiting initiation
activity that is promoted by the redox switch. As NTP binding appears electrochemically to
promote primase accessing the [4Fe4S]
3+
redox state, it is expected that NTP-bound primase
would be more susceptible to oxidation to the [4Fe4S]
3+
state, and subsequent degradation
to the [3Fe4S]
+
species in aerobic conditions.
24
,
38
Primase elongation, in contrast, is aided by preincubation with NTPs or, to a lesser extent, a
primed DNA substrate (Figure 4). Incubation with NTPs increases the number of total
products synthesized on primed DNA 1.5−3-fold compared to primase not preloaded with
substrates.
Incubation with template DNA increases catalytic activity by a modest degree;
t
= 1 min
showed no difference in products under these conditions, and
t
= 30 min showed the largest
increase, synthesizing 187% of the products formed when primase was incubated without
any substrates. These numbers increase slightly (Figure 4) when only elongation products
(32−60 nt) are quantified and compared to primase only preincubation conditions. We also
conducted these assays under aerobic conditions and saw the same general pattern with more
error, likely due to nonspecific oxidation of the primase cluster over time.
39
DISCUSSION
The dynamic interdomain movement and interactions of the heterodimeric DNA primase
enzyme (p48/p58) are distinct from the isolated catalytic domain in the p48 subunit and the
[4Fe4S] domain in the p58 subunit. Here we show that the [4Fe4S] cluster in p58C
participates in DNA-mediated redox signaling in the context of the full p48/p58 heterodimer.
Comparison of the electrochemical behavior of the primase dimer versus p58C is highly
informative because the [4Fe4S] domain acts in concert with the catalytic p48 subunit to
regulate priming. In isolation, the oxidized and reduced p58C domain exhibit very different
electrochemical properties.
20
Remarkably, electrochemically oxidized and reduced p48/p58
behave similarly in the presence of a DNA template.
We attribute this effect to the alignment of p58C in close proximity to the p48 subunit,
which is promoted by the interaction of both subunits with the polyanionic DNA.
42
We
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postulate the alignment of the two domains of the heterodimer may also affect the coupling
of the cluster to the DNA bases. Thus, the primase heterodimer readily participates in robust,
semireversible electrochemical activity only in the presence of both DNA and NTPs. NTP
binding has been demonstrated previously to enhance redox activity of [4Fe4S] enzymes on
DNA, as in the case of DNA repair helicase XPD. The [4Fe4S] cluster in XPD, an ATP-
dependent enzyme, is better coupled into the DNA bases in the presence of ATP and thus
may signal other [4Fe4S] repair enzymes on DNA when it is active.
44
The concurrent
switches to enhanced redox signaling and enzymatic activity may indicate a general linkage
between catalytic and regulatory functions of DNA-processing, [4Fe4S] enzymes. We also
observe that preloading of primase with NTPs or primed DNA enhances catalytic activity.
The p48/p58 complex may sample more configurations that promote primer synthesis when
loaded with one of the required substrates for catalysis, resulting in more complete and
efficient elongation.
It is reasonable to conclude that electrostatic interactions with the primase [4Fe4S] cluster
drive the substrate-dependent change in primase redox behavior. These clusters are tunable
cofactors, with redox potentials influenced by factors such as solvent exposure and
electrostatic environment.
32
DNA and NTPs both carry multiple negative charges, and
binding to DNA has previously been demonstrated to alter redox properties of [4Fe4S] DNA
repair proteins such as base excision repair glycoslyases MutY and Endonuclease III in
Escherichia coli
.
28
,
43
Binding of DNA shifts the redox potential of the cluster negative,
stabilizing the oxidized [4Fe4S]
3+
protein.
Electrochemically oxidized [4Fe4S]
3+
Endonuclease III was moreover directly demonstrated
to bind 500-fold more tightly to DNA than reduced [4Fe4S]
2+
Endonuclease III using
microscale thermophoresis.
28
Upon modeling the addition of negative charges from DNA
into the Endonuclease III cluster environment, the change in potential could be calculated by
summing the electrostatic interactions between negative charges on bound DNA and the
positively charged [4Fe4S] cofactor. This model predicted the potential shifts of both MutY
and Endonuclease III upon DNA binding, and it is interesting to think about expanding the
model to consider DNA-binding, NTP-binding [4Fe4S] enzymes like primase. The
electrostatic maps of the isolated p48 subunit, and the DNA-bound p58C subunit (Figure S5)
illustrate the two electrostatic surfaces we predict are aligned during substrate binding and
primase activity. We predict from these structures that the p58C [4Fe4S] cluster is
approximately 25−30 Å from the bound DNA substrate during primer synthesis. The lack of
structural data on NTP-bound primase/p58C poses a challenge for predicting the distance
between the [4Fe4S] cluster and bound NTPs during activity, but based on the changes in
primase electrochemistry upon NTP binding, as well as the observed electrostatic effects on
the [4Fe4S] cluster potentials of MutY and EndoIII,
29
,
43
we propose that NTPs are also
within at least 20−30 Å of the cluster, possibly as close as 10 Å from the cofactor. As
structural data become available on substrate-bound DNA polymerases, this electrostatic
model for predicting potential shifts will become a useful tool for predicting and analyzing
the properties of substrate-bound replication proteins that contain [4Fe4S] clusters.
Structures of substrate-bound primase will moreover elucidate important details of the
configurational realignment for p48/p58 to initiate and extend the initial RNA primer. Our
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results show that primase elongation on RNA-primed template DNA is enhanced by
preloading NTPs onto the p48/p58 heterodimer. This result illuminates an important effect,
connected with the observed configurational realignment of the primase heterodimer during
replication.
6
,
41
,
42
CONCLUSIONS
Redox switching driven by a change in [4Fe4S] oxidation state has now been demonstrated
to modulate the DNA binding affinity of several DNA-processing [4Fe4S] enzymes in a
manner that regulates activity.
19
,
20
,
25
,
26
,
43
,
44
The primase heterodimer is regulated by a
redox switch in the [4Fe4S] cluster; upon NTP binding the primase cluster can cycle readily
between the [4Fe4S]
2+
and [4Fe4S]
3+
oxidation states, the p58C domain binding more
tightly once oxidized. The electrostatic interaction of polyanionic DNA and NTPs with the
primase cluster, in concert with the configurational alignment required for catalytic activity,
may allow for efficient and regulated primer synthesis and handoff to DNA polymer-ase
α
(Figure 5). Structural analysis and modeling performed on polymerase
α
has suggested that
this enzyme also undergoes significant configurational rearrangements during priming.
11
,
45
The lagging strand polymerase, DNA polymerase
δ
, which contains a redox-active [4Fe4S]
cluster
16
,
19
also adopts different configurations bound to and dissociated from DNA, which
are related to polymerase
δ
interactions with the proliferating cell nuclear antigen (PCNA)
processivity clamp,
46
and here too activity is regulated by the oxidation state of the [4Fe4S]
cluster.
The [4Fe4S] enzymes central to DNA replication often contain many subunits and domains
with flexible tethers, which interact with one another over the trajectory of a step such as
priming and require careful coordination. Replication polymerases known to contain
[4Fe4S] clusters
14
16
bind DNA, NTPs and dNTPs, anionic substrates that alter the
electrostatic environment of the cluster. Both the charged substrates bound to the [4Fe4S]
enzyme and the configuration of the polymerase subunits during the reactions are crucial
elements determining when and how the proteins participate in redox signaling on DNA.
Understanding configurational realignment of subunits and domains, as well as the redox
properties of isolated [4Fe4S] domains versus the corresponding intact proteins, will
facilitate more accurate and thorough construction of [4Fe4S] protein redox signaling
networks in replication, and other pathways containing dynamic, multi-subunit [4Fe4S]
enzymes.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
ACKNOWLEDGMENTS
This research was supported by National Institutes of Health grants R01 GM126904 (J.K.B.), R35 GM118089
(W.J.C.), T32 GM80320 (L.E.S. and M.E.H.) and T32GM07616 (E.O.B.) with additional support from the Moore
Foundation (J.K.B.) and a Ralph M. Parsons fellowship (E.O.B.).
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