# Rrm3 and Pif1 division of labor during replication through leading and lagging strand G-quadruplex

Mor Varon, Daniel Dovrat, Jonathan Heuzé, Ioannis Tsirkas, Saurabh Singh,

Philippe Pasero, Roberto Galletto, Amir Aharoni

► To cite this version:

Mor Varon, Daniel Dovrat, Jonathan Heuzé, Ioannis Tsirkas, Saurabh Singh, et al., Rrm3 and Pif1 division of labor during replication through leading and lagging strand G-quadruplex. Nucleic Acids Research, 2024, 52 (4), pp.1753-1762. 10.1093/nar/gkad1205. hal-04795210

HAL Id: hal-04795210

https://hal.science/hal-04795210v1

Submitted on 21 Nov 2024

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# Rrm3 and Pif1 division of labor during replication through leading and lagging strand G-quadruplex

Mor Varon<fnref n="1" />, Daniel Dovrat<fnref n="1" />, Jonathan Heuzé ①,②, Ioannis Tsirkas<fnref n="1" />, Saurabh P. Singh<fnref n="3" />,
Philippe Pasero ①,②, Roberto Galletto<fnref n="3" /> and Amir Aharoni ①,*,

## Abstract

Members of the conserved Pif1 family of 5'-3' DNA helicases can unwind G4s and mitigate their negative impact on genome stability. In *Saccharomyces cerevisiae*, two Pif1 family members, Pif1 and Rrm3, contribute to the suppression of genomic instability at diverse regions including telomeres, centromeres and tRNA genes. While Pif1 can resolve lagging strand G4s *in vivo*, little is known regarding Rrm3 function at G4s and its cooperation with Pif1 for G4 replication. Here, we monitored replication through G4 sequences in real time to show that Rrm3 is essential for efficient replisome progression through G4s located on the leading strand template, but not on the lagging strand. We found that Rrm3 importance for replication through G4s is dependent on its catalytic activity and its N-terminal unstructured region. Overall, we show that Rrm3 and Pif1 exhibit a division of labor that enables robust replication fork progression through leading and lagging strand G4s, respectively.

## Graphical abstract

<figure><img src="image_2.png" /></figure>

## Introduction

G-quadruplex structures (G4s) are stable non-canonical four-stranded DNA secondary structures formed by non-Watson-Crick base pairing (1). G4s are widely spread across prokaryotic and eukaryotic genomes, and many G4 motifs are evolutionarily conserved, suggesting that they have important roles in genome regulation (2,3). Formation of G4s can perturb DNA replication by acting as an obstacle to the progression of the replisome (4). Stalled replisomes at G4 sites can lead to increased levels of double strand breaks and recombination, thereby promoting genomic instability. Thus, mutations

in DNA helicases that can unwind G4s, are linked to genomic instability and are associated with disease and cancer (5). While the relevance of G4s to genome stability is widely acknowledged, the factors and mechanisms by which cells mitigate their potential threat for genome stability remain unclear.

The Pif1 protein family, that includes highly conserved 5'-3' DNA helicases, are found in all eukaryotes and in some prokaryotes. These helicases can process G4s and mitigate their negative impacts on genome stability (4). In *Saccharomyces cerevisiae*, two helicases of the Pif1 family, Pif1 and Rrm3, contribute to multiple DNA processes including

<footnote n="1">¹Department of Life Sciences and the National Institute for Biotechnology in the Negev, Ben-Gurion University of the Negev, Be’er Sheva 84105, Israel</footnote>

<footnote n="2">²Institut de Génétique Humaine, CNRS, Université de Montpellier, Equipe Labellisée Ligue Contre le Cancer, 34396 Montpellier, France</footnote>

<footnote n="3">³Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO 63110, USA</footnote>

<footnote>*To whom correspondence should be addressed. Tel: +972 8 6472645; Email: aaharoni@bgu.ac.il</footnote>
telomere maintenance (6), centromere function (7), and
Okazaki fragment maturation during DNA replication (8,9).
Previous studies showed that Pif1 can efficiently unwind G4s
*in vitro* and can also resolve G4s *in vivo* (10), thereby mitigat-
ing G4-induced mutagenesis and genome instability (11–13).
However, Pif1 only enables replisome progression through
G4s located on the lagging strand template (10), and it re-
mains unknown whether similar mechanisms exist for repli-
cation through leading strand G4s. Rrm3 was shown to have
a broad effect on genome maintenance including the suppres-
sion of recombination at repeat sequences (14) and prevention
of replisome stalling at a variety of chromosomal loci includ-
ing centromeres, telomeres, tRNA genes and transcribed genes
(9). The ability of Rrm3 to prevent fork stalling at a variety
of genomic locations has been associated with its ability to
remove proteins bound to the DNA ahead of the replication
fork (15). Yet, while Rrm3 is associated *in vivo* with many ge-
nomic processes overcoming various obstacles, less is known
of Rrm3 activity at G4s and its importance for efficient repli-
some progression through G4 rich sequences.

Here, we have directly examined the importance of Rrm3
for DNA replication progression through G4 sequences. We
have utilized our recently developed approach for measur-
ing the progression rate of single replication forks (16), as
they replicate through G4 sequences located on the leading
or lagging strand template, in living cells (10). Using this ap-
proach, we found that Rrm3 is important for efficient repli-
cation through G4s located on the leading strand template,
but not on the lagging strand template. We show that sta-
bilization of leading strand G4s, by increasing the number
of G4 repeats or by G4 stabilizing ligand, leads to increased
levels of fork slowdown in the absence of Rrm3. Finally, we
show that the importance of Rrm3 for replication through
leading G4s is dependent on its catalytic activity and N-
terminal unstructured region. Our results provide direct *in
vivo* evidence for a function of Rrm3 at G4s and its impor-
tance for replication through G4 sequences within the leading
strand.

**Materials and methods**

Strain generation

Strains for replication time measurement were generated on
the background of a W303 MATA<sub>a</sub> *Saccharomyces cerevisiae*
strain, expressing LacI-HaloTag and tetR-tdTomato fusion
proteins in the nucleus. LacOx256 and tetOx224 arrays
are located at chrIV:332960 and chrIV:352560 respectively,
near Autonomously Replicating Sequence (ARS) 413 with a
mid-array distance of approximately 30.6 kb (16). PIF1 was
replaced by a *kanMX* antibiotic cassette using the Lithium
Acetate (LiAc) transformation method. Rrm3-AID strain was
generated by integrating a PCR cassette containing the auxin-
inducible degron (AID) sequence followed by 6xFLAG-Tag
and an *hphMX* antibiotic cassette at the 3' end of the Rrm3
gene. The Oryza sativa *TIR1* gene was integrated at the Ade1
locus to facilitate Rrm3-AID degradation following auxin
addition. Rrm3 truncation strains were generated on the
background of Rrm3-AID strain by integrating the different
truncations followed by 6xHA-tag into the *URA3* locus using
an integrative plasmid. To generate the plasmids containing
the Rrm3 truncations, a gBlock (IDT) containing the endoge-
nous promoter of Rrm3 along with an NruI restriction site,

6xHA-tag and the CYC1 terminator was inserted into the
pRS306 integrative plasmid via Gibson assembly, following
the manufacturer's protocol (NEB). Different truncations of
the Rrm3 gene were then amplified from WT yeast genomic
DNA and inserted into the plasmid that was restricted by
NruI restriction enzyme via Gibson assembly (NEB). The G4
motifs were inserted 11.1 kb upstream of ARS413 and 8.4
kb downstream of the tetO array. The Different G4 forming
sequences were integrated into the pif1-deleted/Rrm3-AID
cells by replacing a natMX cassette, inserted at the same
genomic location, using a marker-free CRISPR/CAS9 ap-
proach as previously described (17). For the generation of a
strain containing tRNA between the lacO and tetO arrays,
the glutamate tRNA gene (YNCI0011W) was amplified from
the yeast genome along with 362 bp of upstream and 214
bp of downstream sequences to maintain proper sequence
context. The tRNA was inserted ~3 kb downstream of the
lacO array in a head-on (HO) orientation relative to the
expected direction of replisome progression in this locus, by
replacing the natMX cassette as described above. For the
generation of strains for copy number variation analysis,
G4(A+B) or G4(6) were integrated into a RRM3-AID strain
that does not contain the lacO and tetO arrays and the cog-
nate repressors. These G4s were integrated at chrIV:332960,
at a distance of 3 kb from ARS413. The G4(A+B) sequence
contains the G4A motif, separated by 96 bp of a spacer
sequence, followed by G4B (10). The full sequence is
GGGTACGGTGGGTAATAAGGAAGGTATCGGGTT
AGATCCCAGTCGAATGGATTAATCAAACAG
ATCTGTAGCCGGAGAGGCATACCCCCTGCG
ACACTTTACGAAGGCATCTGCAAAAATCATAA
CTGGGGAGGGGAAGGGGAGGGG where G4A sequence
is underlined first followed by an underline for G4B. The
G4(4) and G4(6) sequences contain the core G4 motif
(G3T3)3G3 (18), repeating 4 and 6 times, respectively, con-
taining a random linker of 40 bp between each of the G4
motifs. These repeat G4 sequences were synthesized as
synthetic genes (BioBasic). All replacements were validated
with Polymerase Chain Reaction (PCR) followed by Sanger
sequencing.

Microscopy

Yeast cells were grown overnight in synthetic complete (SC)
medium containing 4% glucose at 30°C. Yeast cultures were
diluted at OD₆₀₀ = 0.1 and SiR-HALO dye was added to a
final concentration of 400 nM (19). One hour following SiR-
HALO dye addition, 10 µg/ml of α-factor was added to ar-
rest the cells in G1 phase, and the cultures were incubated for
two additional hours to facilitate synchronization. In RRM3-
AID strains, 1 mM of indole-3 acetic acid (IAA) was added
1 h prior to the experiment to facilitate the degradation of
the endogenous Rrm3 protein. In experiments containing the
PhenDC3 compound 10 µM of PhenDC3 were added immedi-
ately after the cells were diluted. Cells were then immobilized
on microscopy slide chambers (Ibidi) coated with 2 mg/ml
concanavalin A and washed thoroughly from α-factor and
SiR-HALO dye with warm SC medium containing 4% glu-
cose prior to microscopy experiments. Live-cell imaging of the
cells was performed on an AxioObserver inverted wide-field
microscope (Zeiss) with a Colibri 7 LED light source, at 1 min
intervals for 4 h at 30°C, using a ×63 oil objective (NA = 1.4)
in 3D (8 z-sections 0.8 µm apart). LacI-Halo-SiR and TetR-
tdTomato were excited with 631 and 555 nm illumination,
respectively.

Copy number variation analysis

For copy number variation analysis, yeast cells were grown
overnight at 25°C to a cell density of 5–6 × 10⁶ cells/ml in
200 ml of YPD medium. Cells were synchronized in G1 by
the addition of 8 µg/ml of α-factor for 2:30 hours. Half an
hour following α-factor addition, IAA was added to facilitate
Rrm3 depletion only to the target cells at a final concentra-
tion of 1 mM. To slowdown replication forks, hydroxyurea
(HU) was added 20 min prior to cell release to a final con-
centration of 50 mM. To release the cells into S-phase follow-
ing α-factor synchronization, Pronase (Sigma) was added to
a final concentration of 75 µg/ml. Immediately prior to cell
release, 45 ml of culture was taken as a G1 control sample
for the extraction of genomic DNA (gDNA). Following cell
release into S-phase, 45 ml of culture samples were taken for
gDNA extraction at 35, 47 and 60 min time points. In par-
allel, 0.5 ml of cell samples were taken from asynchronous
population, G1 cells and 35, 47, 60 and 90 min following
release into S-phase for flow cytometry analysis. For gDNA
extraction, spheroplasts were prepared using Zymolyase 20T
(Nacalai) according to standard protocol followed by lysis of
the spheroplasts using G2 buffer (Qiagen) supplemented with
RNaseA and proteinase K. Standard phenol-chloroform ex-
traction was then used to separate the gDNA from the pro-
teins and lipids followed by isopropanol DNA precipitation
and the pellet was washed with cold 70% EtOH. The purity
and concentration of all gDNA samples were verified by Nan-
oDrop 1000 UV Visible Spectrophotometer (Thermo Fisher
Scientific). DNA copy number levels at the vicinity of *ARS413*
and *ARS607* were measured using real-time PCR (LightCy-
cler 480 II, Roche) in 384-well plates using a series of primers
located at the vicinity of these origins at a spacing of 1–2
kb distance (Supplementary Table S1). Primer specificity was
first examined in silico by blasting primers sequences with the
*S. cerevisiae* reference genome (*sacCer3*, April 2011 version).
Only primers that hybridize once on the reference genome
were considered as specific and were further tested for speci-
ficity and efficiency experimentally by qPCR using a range
of genomic DNA concentrations (see below). The PCR pro-
gram used for amplification included: 10 min at 95°C, then
45 cycles of: 95°C 10 s, 60°C 15 s and 72°C 15 s. Primer
specificity and efficiency were determined using ‘Tm calling’
‘Abs Quant:2nd Derivative Max’ tools from the Light Cycler
480 software (Roche). All primers were designed to generate
an amplicon of 80 bp and exhibited a single sharp peak in
the melting curve analysis showing their high specificity. The
primer efficiency calculation was performed by plotting the
correlation of quantification cycle (Cq) to log DNA template
concentrations (10, 1, 0.1 and 0.01 ng/µl) using the equation
y = Bx + A. Primers efficiency was determined using the equa-
tion: E = 10^(-1/B). All the primers used in this study had an
efficiency between 1.98 and 2.05 with a R fitting >0.999. The
qPCR reactions (Roche LightCycler FastSTART DNA master
SYBR green I) were performed according to the manufacturer
instructions. Determination of DNA concentrations was per-
formed by amplification of reference asynchronous gDNA at
four different concentrations (10, 1, 0.1 and 0.01 ng/µl). Copy
number levels at the vicinity of *ARS413* and *ARS607* were

normalized to G1 levels (copy number of 1) to determine du-
plication level of replicated regions.

Western blot analysis

For the detection of Rrm3-AID-6xFLAG by western blot, 50
ml of yeast cells were grown until late-exponential phase.
Rrm3 depletion was induced by the addition of 1 mM of
IAA for 1 h before harvest. Cells were centrifuged and pel-
lets were frozen at -20°C overnight. Next, cell pellets were
thawed and incubated in 1 ml of 0.2 M NaOH for 5 min,
followed by boiling at 95°C for 5 min in 50 µl of Laemmli
Buffer including 10% β-mercaptoethanol. Lysates were cen-
trifuged for 2 min and 15 µl of the supernatants were loaded
and run on a precast 10% SDS-PAGE gel (Bio-Rad). For the
detection of Rrm3 different truncation fused to 6xHA, 2 ml
of cells in OD600 = 1 were collected and 1 h after 1 mM IAA
addition. Then, cells were centrifuged and pellets were resus-
pended in 500 µl ddH2O and 75 µl of 1.85 M NaOH/7.5%
β-Mercaptoethanol for 15 min, followed by addition of 75 µl
of 55% TCA solution. Cells were then centrifuged and the pel-
let was boiled at 95°C in loading buffer containing 8 M urea,
5% SDS, 200 mM Tris-HCl, 1 mM EDTA and 0.1% bro-
mophenol blue. For each sample 10 µl of the supernatant was
loaded and run on a precast 10% SDS-PAGE gel (Bio-Rad).
Separated protein bands were transferred to a PVDF mem-
brane using a Trans-Blot Turbo system (BioRad) followed by
blocking with PBST (PBS buffer containing 0.1% Tween 20)
+ 5% skim milk for 1 h. The membrane was washed with
PBST buffer 3 times for 5 min, and was incubated with a pri-
mary mouse anti-FLAG antibody conjugated with HRP en-
zyme (1:1000) (Sigma-Aldrich Cat# A8592) in PBST + 1%
skim milk for 1 h. The membrane was washed three times and
detected with the EZ-ECL Chemiluminescence detection kit
(Biological Industries) according to the manufacturer's pro-
tocol. Detection of Rrm3 truncations were performed with
anti-HA antibody (1:2000, Santa Cruz Cat# sc-7392) and a
secondary goat-anti-mouse antibody (1:10 000) (Jackson Im-
munoresearch Cat# 115-035-003) conjugated with HRP. De-
tection of Pgk1 bands was performed with a primary mouse
anti-Pgk1 antibody (1:5000, Invitrogen Cat# 459250) and a
secondary goat-anti-mouse antibody (1:10 000, Jackson Im-
munoresearch Cat# 115=035-003) conjugated with HRP.

Quantification and statistical analysis

Time-lapse measurements were collected with ZEN 3.0 and
analyzed using a custom-made computational pipeline devel-
oped specifically for the analysis of replication rates (16). Our
MATLAB-based pipeline identifies, tracks and quantifies the
LacI-Halo-SiR and TetR-tdTomato dots in each cell. For each
strain, at least 40 cells were measured in two independent ex-
periments. Statistical analysis of replication time data was per-
formed using Monte Carlo resampling with 1 000 000 itera-
tions. Swarm plots were plotted using the Seaborn package in
Python.

Purification of Rrm3 constructs for biochemical
analysis

N-terminal deletion constructs of RRM3 (comprising residues 187-723, 213-723 and 231-723) were PCR amplified from the full-length gene (originally from the yeast strain W303a) and cloned in pET28a at the NdeI/XhoI restriction sites, yielding an N-terminal His₆-tag. The Rrm3 constructs were
over-expressed in *E. coli* Rosetta2 (DE3) pLysS by induction with 0.5 mM IPTG and growth overnight at 16°C. The cells were suspended in Buffer L (50 mM Tris-HCl pH 8.0 (4°C), 500 mM NaCl, 5 mM β-ME, 10% v/v glycerol, 1 mM PMSF) + 10 mM imidazole, lysed by sonication, followed by centrifugation at 15 000 rpm for 1 h in a Beckman JA-14 rotor. The supernatant was then loaded on a HisPur Ni-NTA superflow agarose (Thermo Scientific), washed with 20 mM imidazole in buffer L and eluted with 300 mM imidazole in Buffer L. The fractions containing Rrm3 were dialyzed in Buffer P (50 mM Tris-HCl pH 7.5 (4°C), 1 mM EDTA, 1 mM DTT, 20% v/v glycerol and 0.1 mM PMSF) + 400 mM NaCl. After extensive dialysis, the sample was diluted 2-fold with buffer P (to reduce the NaCl concentration to 200 mM), loaded on a High Q column (BioRad), the flow-through collected and loaded on a High S column (BioRad) and then eluted at 1 M NaCl with Buffer P. The eluted proteins were dialyzed in storage buffer (50 mM Tris-HCl pH 8, 600 mM NaCl, 1 mM EDTA, 1 mM DTT, 40% v/v glycerol) and stored at -80°C.

## ATPase and DNA unwinding assays

The DNA-dependent ATPase activity was determined spectrophotometrically using a β-Nicotinamide adenine dinucleotide reduced disodium salt hydrate (NADH, LADH, PK, PEP) enzyme-coupled assay as previously reported (20). ATPase activity was measured at 30°C in buffer 20 mM Hepes (pH 7.4), 20 mM NaCl, 2 mM Mg-acetate and 1 mM DTT at a constant concentration of 20 nM Rrm3, 1 mM ATP, and in the presence or absence of 200 nM ssDNA (dT<sub>60</sub>). Fluorescence experiments were performed with an L-format PC1 spectrofluorimeter (ISS, Champaign, IL). DNA unwinding experiments were performed using a DNA substrate consisting of a double-stranded region of 20 bp and a 5'ssDNA tail of 20 nt (dT20). At the blunt-end of the DNA, the 3'-end was labeled with FAM (carboxy fluorescein) while the opposite 5' end was labeled with the quencher Iowa Black FQ (IDT). Rrm3-dependent DNA unwinding of the 20 bp dsDNA region results in the de-queching of FAM fluorescence (λ<sub>ex</sub> = 490 nm, λ<sub>em</sub> = 530 nm) due to the separation of the two strands. The experiments were performed in 20 mM Hepes (pH 7.4), 20 mM NaCl, 2 mM Mg acetate, 2 mM DTT, 1mg/ml BSA, and 0.1% C12E10 at 30°C. A complex of 20 nM DNA was formed in the presence of a 4-fold excess concentration of Rrm3, and the reactions were started by the addition of 1 mM ATP and 100 nM of ssDNA trap (same as the strand carrying the quencher) to prevent re-annealing of the dsDNA region.

## Results

### Rrm3 and Pif1 division of labor at G4 sequences

We recently described an approach for measuring the rate of DNA replication at a specific genomic locus in single live S. cerevisiae cells. This approach is based on monitoring the intensity of fluorescently labeled *lacO* and *tetO* arrays visualized as fluorescent dots under time-lapse microscopy (16). Tracking the intensity of each fluorescent dot in individual cells allows us to identify when each locus is being replicated and to measure progression of a single replisome through the genomic region between the arrays. We applied this approach to measure replisome progression through G4 sequences integrated between fluorescently labeled *lacO* and *tetO* arrays

located downstream of *ARS413* (10). Using this approach, we previously examined replication through two tandem G4 sequences ($G4_{(A+B)}$), derived from the yeast genome (21), located on the lagging or leading strand template (10) (Figure 1A, B).

Here, we measured replication fork progression in yeast strains containing Rrm3 fused to an auxin-inducible degron (AID), *RRM3-AID*. Rrm3 degradation in synchronized yeast populations was initiated by the addition of auxin during G1, prior to release into S phase and the initiation of DNA replication. It was previously shown, using sequencing approaches and 2D gels, that the deletion of Rrm3 leads to a significant fork stalling at tRNA genes, which are widely spread in the yeast genome (22–24). A recent study utilized a Nanopore sequencing approach for the detection of fork stalling at a head-on (HO) Glutamate tRNA gene (YNCI0011W) in *rrm3-deletion* strain (24). To examine whether our live cell microscopy system is consistent with previous measurements of fork stalling in the absence of Rrm3, we integrated this tRNA gene in a HO orientation, including upstream and downstream regions, between the *lacO* and *tetO* arrays in a *RRM3-AID* strain. Replication measurements of this strain revealed a significant replication slowdown through the tRNA upon Rrm3 depletion (Supplementary Figure S1). These results are in accordance with previous measurements using site specific or genomic approaches (22–24) highlighting the sensitivity of our approach for the detection of fork slowdown at a known roadblock due to Rrm3 depletion.

Next, we examined replication through G4s in the presence or absence of Rrm3 using the live-cell microscopy system. We found that in cells depleted of Rrm3, replication fork progression through $G4_{(A+B)}$ placed in the leading strand template is significantly slowed down, relative to cells expressing Rrm3 (Figure 1C and Supplementary Figure S2). In contrast, Rrm3 depletion does not lead to replication slowdown through $G4_{(A+B)}$ placed in the lagging strand template (Figure 1D and Supplementary Figure S2). These results highlight the importance of Rrm3 for enabling normal replication progression through leading strand G4s only.

Previously, we found that Pif1 is essential for efficient replication through lagging strand G4s, but not leading strand G4s (10). To further study the interplay between Pif1 and Rrm3 for enabling efficient replication through G4s, we examined replication in *pif1Δ* strains in the presence or absence of Rrm3. We found that *pif1*-deletion, generated on the background of *RRM3-AID* strain, leads to a mild slowdown of replication through leading strand G4s (Figure 1C) in the absence of IAA, but no slowdown was observed when *pif1*-deletion was generated on *RRM3* background (Supplementary Figure S3). Importantly, we found that in *pif1Δ* cells depleted of Rrm3, replication through leading strand $G4_{(A+B)}$ is significantly slower relative to *PIF1* WT cells depleted of Rrm3 (Figure 1C). In accordance with our previous study (10), we found that in *pif1Δ* cells replication is slowed down through lagging strand $G4_{(A+B)}$ (Figure 1D). However, further depletion of Rrm3 in the *pif1Δ* cells does not aggravate replication slowdown (Figure 1D).

To further support our findings, we utilized the copy number variation approach to examine *RRM3-AID* and *pif1*-deletion strains containing leading $G4_{(A+B)}$. Cells were synchronized and released with or without Rrm3 depletion and samples were collected for gDNA extraction. Specifically, we collected samples during G1 phase as a control and samples following 35, 47 and 60 min after release into S-phase
A

<figure><img src="image_3.png" /></figure>

B

<figure><img src="image_4.png" /></figure>

C

G4<sub>A+B</sub> leading

<figure><img src="image_5.png" /></figure>

D

G4<sub>A+B</sub> lagging

<figure><img src="image_6.png" /></figure>

**Figure 1.** Division of labor between Rrm3 and Pif1 during replication through leading and lagging strand G4s, respectively. (A, B) Schematic illustration of the experimental system, which allows monitoring the progression of single replication forks through G4s located on the leading (A) or lagging (B) strand templates using fluorescently labeled *lacO* (dark red) and *tetO* (orange) arrays. (C, D) Replication times through a region of ~30 kb (mid-array distance) containing G4<sub>(A+B)</sub> located on the leading (C) or lagging strand (D) templates for Rrm3 expressing cells, cells depleted in Rrm3, cells containing *pif1*-deletion and cells lacking both helicases. All strains contain *RRM3-AID*, and Rrm3 depletion was induced by the addition of 1 mM IAA, as described in the Materials and methods section.

(Supplementary Figure S4). We performed our experiments in the presence of 50 mM hydroxyurea (HU) which slow-down fork progression by ~6-fold (25) and thus increase the time window for the detection of copy number variations. To examine fork progression from *ARS413* through leading G4<sub>(A+B)</sub>, we have used qPCR with primers annealing to genetic regions located before or after the G4<sub>(A+B)</sub> sequence. This gDNA analysis revealed a decrease of copy number level downstream of the G4<sub>(A+B)</sub> sequence upon Rrm3 depletion, relative to Rrm3 expressing conditions (Supplementary Figure S5). Analysis of copy number upstream and downstream of *ARS607* in the presence or absence of Rrm3 reveals mild variation between the conditions (Supplementary Figure S5) that could results from natural G4 forming sequences located at the vicinity of this origin (21).

Overall, the live cell microscopy data and copy number variation analysis demonstrate the importance of Rrm3 for replication through leading G4s and the division of labor between Rrm3 and Pif1 for replication through leading and lagging G4s, respectively (Figure 1 and Supplementary Figure S5). In addition, our results reveal that in the absence of Rrm3 activity, Pif1 can partially suppress replication slowdown at leading G4s. However, in the absence of Pif1, Rrm3 cannot

suppress replication slowdown at lagging G4s, highlighting the higher specificity of Rrm3 for leading G4s (Figure 1C, D).

## Replication slowdown in the absence of Rrm3 is dependent on G4 stability

To examine the effect of leading strand G4 stability on replication fork progression in the presence or absence of Rrm3, we first used the G4 stabilizing ligand PhenDC3. Many G4 ligands have been thoroughly characterized *in vitro* and their effects were tested in yeast or in mammalian cells (26–28). Specifically, PhenDC3 was shown to increase genome instability and recombination at G4 regions in yeast cells (13,29). Here, we examined whether PheDC3 can affect replication fork progression through leading G4<sub>(A+B)</sub>, in WT or Rrm3 depleted cells. We found that replication fork progression through leading G4<sub>(A+B)</sub> is moderately affected by the presence of PhenDC3 in cells expressing Rrm3 (Figure 2A). However, we found that replication fork progression through the leading G4s is significantly slowed down by the presence of PhenDC3 in cells depleted of Rrm3, relative to cells expressing Rrm3 (Figure 2A). To verify that PhenDC3 leads to replication slowdown strictly due to the stabilization of G4s, we
<figure><img src="image_7.png" /></figure>

<figure><img src="image_8.png" /><figcaption>**Figure 2.** Replication fork progression through leading strand G4s in Rrm3 depleted cells is dependent on G4 stability and the number of G4 motifs. (A) Replication times through leading G4<sub>(A+B)</sub> in the presence or absence of PhenDC3 and in the presence or absence of Rrm3. Data for strains measured in the absence of PhenDC3 is adopted from Figure 1C and is shown for comparison. (B) Replication times through 4 or 6 repeats (G4(4) and G4(6), respectively) of leading strand G4 sequence (G<sub>3</sub>T<sub>3</sub>)<sub>3</sub>G<sub>3</sub>, or in the absence of G4, in Rrm3 expressing (blue) or Rrm3 depleted strains (red). Depletion of Rrm3 (red) results in significant slowdowns of replication through leading G4(4) or G4(6), relative to the respective strains expressing Rrm3 (red). All strains contain *RRM3-AID*, and Rrm3 depletion was induced by the addition of 1mM IAA, as described in the Materials and methods section.</figcaption></figure>

examined whether PhenDC3 slows replication in the absence
of G4<sub>(A+B)</sub> and found no effect (Figure 2A).

To further examine whether replication fork progression through leading G4s is affected by the nature and number of the G4 motifs, we integrated different G4 repeats into *RRM3-AID* cells. Previous bioinformatics analysis of G4 sequences in the yeast genome indicated that the average number of con-

served tandem G4 repeats is 5.1 (21). Thus, we chose to in-
tegrate 4 or 6 tandem repeats of a synthetic (G₃T₃)₃G₃ se-
quence into the leading strand template (Figure 2B, G4(4)
and G4(6), respectively), in the background of a RRM3-AID
strain. This G4 sequence was previously shown to have a mod-
erate stability of 58.8°C (18). We found that replication pro-
gression from ARS413 through genomic DNA that do not
contain the G4s is normal in the absence of Rrm3 (Figure
2B). In contrast, we found a significant replication slowdown
through G4(4) and G4(6) in cells depleted of Rrm3, rela-
tive to the Rrm3 expressing cells. To further support replica-
tion slowdown through G4(6) upon depletion of Rrm3 and
examine the effect of PhenDC3, we have utilized the copy
number variation approach described above. We constructed
a leading G4(6) strain, integrated 3 kb from ARS413, con-
taining RRM3-AID. Copy number variation experiments for
the analysis of fork progression through G4(6) in the ab-
sence of PhenDC3 revealed a mild decrease upon Rrm3 de-
pletion (Supplementary Figure S6). We found that the addi-
tion of PhenDC3 to Rrm3 depleted cells results in a larger de-
crease of copy number downstream of the G4(6), relative to
Rrm3 expressing cells treated with PhenDC3 (Supplementary
Figure S6). These results are in agreement with our live-cell
microscopy findings demonstrating that Rrm3 is required for
replication fork progression through different G4-forming se-
quences, and that the extent of slowdown is dependent on G4
stability.

The importance of Rrm3 functional domains for
replication through leading strand G4s

To examine the importance of Rrm3 helicase activity for repli-
cation through leading strand G4 sequences, we examined
replication in strains expressing the K260A Rrm3 ATPase
dead mutant (30). To enable the complementation of Rrm3
WT and mutant in Rrm3 depleted cells, we integrated a sec-
ond copy of RRM3 into the URA3 locus on the background
of the RRM3-AID strain. Using these strains, we first verified
that the endogenous Rrm3 is depleted upon auxin addition
and that the complemented Rrm3 WT and K260A mutant are
expressed to a similar level (Figure 3A). Next, we performed
live cell microscopy experiments to measure replication fork
progression through the leading strand G4<sub>(A+B)</sub> in the comple-
mented WT or K260A cells. We found that while the comple-
mentation with WT Rrm3 restores normal replication through
G4<sub>(A+B)</sub>, replication in the K260A cells is significantly slowed
down, similar to Rrm3 depletion (Figure 3B). This indicates
that the helicase activity of Rrm3 is essential for efficient repli-
cation fork progression through leading strand G4 sequences.

Previous studies revealed that Rrm3 contains a non-
conserved intrinsically disordered N-terminal domain and
a conserved helicase domain (9). To map the borders of
the Rrm3 N-terminal unstructured domain and the helicase
domain, we performed AlphaFold structural prediction of
Rrm3 (31) and IUPred sequence analysis (32). Using these
tools, we verified that the N-terminal unstructured domain
of Rrm3 spans residues 1–230, followed by a well-conserved
and folded helicase domain (9) (Supplementary Figure S7). A
previous study has identified a PCNA interacting protein (PIP)
motif in the N-terminal domain, located between residues 35–
42, and showed that deletion of the first 54 residues of Rrm3
(Δ54) completely abolishes its interaction with PCNA (33). In
addition, a recent study identified the importance of residues
<figure><img src="image_9.png" /><figcaption>**Figure 3.** Replication through leading G4s is dependent on Rrm3 catalytic activity. (A) Western blot analysis for the detection of Rrm3 depletion following the addition of auxin and the complementation of cells with WT Rrm3 or the K260A mutant expressed from the native *RRM3* promoter and located at the *URA3* locus. (B) Replication times through leading G4A+B of cells expressing the endogenous Rrm3 (WT), cells depleted of Rrm3, and cells complemented with WT Rrm3 or the K260A mutant. Data for replication through leading G4A+B in the presence or absence of endogenous Rrm3 is adapted from Figure 1C and is shown for comparison. All strains contain endogenous *RRM3*-AID, and endogenous Rrm3 depletion was induced by the addition of 1 mM IAA, as described in the Materials and methods section.</figcaption></figure>

186-212 for controlling DNA replication under hydroxyurea replication stress and as a site of interaction with Orc5 unrelated to Rrm3 helicase activity (34).

To examine the importance of the different functional regions of Rrm3 N-terminal domain for replication through leading G4s, we constructed a series of truncated versions of Rrm3 consisting of *rrm3*-Δ54, *rrm3*-Δ186, *rrm3*-Δ212 and *rrm3*-Δ230 (Figure 4A). We introduced these truncated versions of Rrm3 into the *RRM3*-AID strain and verified their expression by western blot analysis (Supplementary Figure S8). Next, we examined the effect of the different Rrm3 truncated versions on replication through leading G4(A+B) in cells depleted of the endogenous Rrm3. We found that the *rrm3*-Δ54 variant facilitates normal replication through the G4s (Figure 4A), despite lower expression of this variant (Supplementary Figure S8). These results show that the interaction of Rrm3 with PCNA is not essential for fork progression through the leading strand G4(A+B) and that low expression of Rrm3 is sufficient for efficient replication at this G4 sequence. Moreover, we found that the *rrm3*-Δ186 variant, lacking 186 aa from the N-terminus, also facilitates replication through G4(A+B) at similar rates (Figure 4B). In contrast, both the *rrm3*-Δ212 and

*rrm3*-Δ230 variants displayed a significant slowdown in leading strand G4 replication. This slowdown is similar to the slowdown observed in the Rrm3 depleted strain (Figure 4B). To verify that the slowdown in replication of the *rrm3*-Δ212 and *rrm3*-Δ230 variants is not due to reduced enzymatic activities, we performed biochemical assays to characterize the purified truncated Rrm3 variants (Figure 4C). These assays revealed that *rrm3*-Δ186, *rrm3*-Δ212 and *rrm3*-Δ230 variants are all active and have similar ATPase and DNA unwinding activities (Figure 4D,E). Overall, these results show that the N-terminal domain is essential for replication fork progression through leading strand G4s. However, while all large N-terminal truncation variants have similar enzymatic activities *in vitro*, truncations larger than Δ212 lead to a slowdown of replication through leading strand G4s *in vivo*. These observations suggest a yet uncharacterized role of the region spanning amino acids 186 to 212 in regulating Rrm3 function.

## Discussion

Here, we discovered that Rrm3 is essential for proper repli-
some progression through different G4 sequences located on
<figure><img src="image_10.png" /><figcaption>**Figure 4.** Fast replication through leading strand G4s requires Rrm3 N-terminal domain. (A) Schematic illustration of the different truncated Rrm3 variants examined including *rrm3*-Δ54 for the deletion of the PIP region in Rrm3, *rrm3*-Δ186, *rrm3*-Δ212 and *rrm3*-Δ230 for the deletion of the whole N-terminal domain. (B) Replication times through leading G4A+B of WT cells, cells depleted in Rrm3, cells complemented with WT or different N-terminal truncated Rrm3 shown in (A). Truncation of the first 212 or 230 residues leads to a dramatic slowdown in replication relative to full length Rrm3, highlighting the importance of these residues for replication through leading G4s. Data for replication through leading G4A+B in the presence or absence of endogenous Rrm3 is adapted from Figure 1A and is shown for comparison. (C) Coomassie stained SDS-PAGE showing the recombinantly purified *rrm3*-Δ186, *rrm3*-Δ212 and *rrm3*-Δ230 variants. (D, E) Helicase (D) and ATPase (E) activity of the different truncated Rrm3 versions in (C) showing no significant difference in activity between the different truncations.</figcaption></figure>

the leading strand template (Figures 1 and 2). The importance of Rrm3 catalytic activity for leading G4 replication (Figure 3) suggests that either Rrm3 helicase activity is needed to unwind G4 structures or its sweepase activity (15,35) is used to remove proteins bound to them. In either case, enzymatically active Rrm3 is required for preventing fork stalling during leading strand replication at G4 forming sequences. Previous studies revealed the importance of Rrm3 in preventing replisome stalling at more than a thousand genomic sites, including centromeres, telomeres, tRNA genes and ribosomal DNA (15). Fork progression through these sites was shown to be dependent on Rrm3 catalytic activity as well (6). Since these pausing sites overlap with sites of stable protein-DNA interactions, it was suggested that Rrm3 could remove protein-DNA complexes ahead of the replication fork (15). However, many of these sites, including gene promoters and telomeres, also contain G4 motifs, raising the possibility that Rrm3 activity at G4s can contribute to replisome progression at these regions. Our findings that Rrm3 facilitates fork progression through leading strand G4s support this possibility, expanding the different functions of Rrm3 during genome replication.

Our current findings that Rrm3 is essential for normal replication only through leading strand G4s (Figure 1) complement our results showing that Pif1 is essential for normal replication through lagging strand G4s (10). These findings highlight a clear division of labor between Rrm3 and Pif1 in enabling efficient replisome progression through leading and lagging strand G4 sequences, respectively. Previous studies character-

ized the number, distribution, and conservation of G4 DNA motifs in yeast (2,21). Based on sequence pattern criteria and conservation, more than 650 G4 sequences were identified in the yeast *Saccharomyces cerevisiae* distributed over all 16 yeast chromosomes (21). Since the yeast genome contain hundreds of G4s and origins of replication (21,36,37), the specialized activities of Rrm3 and Pif1 may enable high flexibility of origin firing for efficient fork progression through G4 sequences located on the leading or lagging strand templates. Such flexibility in G4 replication directionality can be particularly useful when origin firing program is altered due to replication stress and checkpoint activation (38). While we observed a division of labor between Rrm3 and Pif1, our findings that Pif1 can reduce the level of replication slowdown through leading G4s in the absence of Rrm3, reveal some level of cooperation between the two helicases (Figure 1). Interestingly, Rrm3 cannot act on lagging G4s in the absence of Pif1, highlighting its higher specificity in promoting leading strand G4s replication. Cooperation between Rrm3 and Pif1 has been previously observed for replication fork progression through different genomic regions, including ribosomal DNA, tRNA genes and centromeres (7,24,30).

At the molecular level, the ability of Rrm3 to facilitate fork progression through leading strand G4s can be explained by previous findings showing that Rrm3 physically interacts with Pol2, the catalytic subunit of the leading strand-specific DNA polymerase $\varepsilon$ in yeast (39). This interaction suggests that Rrm3 activity at leading strand G4s is tightly coupled to
polymerase progression on the leading strand template. Our results showing the importance of Rrm3 N-terminal region for replication through leading G4s, specifically residues 186–212 (Figure 4), suggest that this region may be important for Rrm3-Pol2 interaction; however, additional experiments are needed to test this possibility. Interestingly, while Rrm3 contains, within the first 54 amino acids, a PIP box that enables its interaction with PCNA (33), deletion of this N-terminal region did not affect fork progression through leading strand G4 sequences (Figure 4). In agreement with these findings, previous studies showed that deletion of Rrm3 PIP region had no effect on Rrm3 control of DNA synthesis during replication stress (34). In contrast, it was shown that mutation or deletion of two PIP regions in Pif1 reduces fork progression through lagging strand G4s (10) and decreases the efficiency of break induced replication (40). While PCNA is located on both leading and lagging strand replication machineries (41), the high functional importance of Pif1-PCNA interaction for replication through lagging G4s is correlated with the importance of PCNA interaction with other lagging strand proteins including Cdc9 ligase, Fen1 and Polymerase δ for lagging strand DNA synthesis (42–44).

## Data availability

All data and code for analysis are available upon request.

## Supplementary data

Supplementary Data are available at NAR Online.

## Funding

Israeli Science foundation (ISF) [707/21]; Binational Science Foundation (BSF-NSF) [2019617, 2021737]; work by R.G. was funded by the National Institutes of Health [1R35GM139508]. Funding for open access charge: Grant and University support.

## Conflict of interest statement

None declared.

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