To study the role of DDC initiation proteins in the maintenance of the cell cycle arrest, we utilized the well-characterized strain JKM179 (Lee et al., 1998), in which the site-specific HO endonuclease is expressed from a GAL1-10 promoter (GAL-HO) to induce a single DSB within the MAT locus on chromosome III (referred to as the 1-DSB strain). In this 1-DSB strain, we inserted an additional HO cleavage site 52 kb from the centromere on chromosome IV to induce another DSB (referred to as the 2-DSB strain) (Kim et al., 2007; Lee et al., 1998). At both loci, HO-mediated cleavage after galactose induction is nearly complete within 30–45 min (Lee et al., 2014). In both strains, we also deleted the HML and HMR donors to prevent repair by homologous recombination. With continuous HO expression, nonhomologous end-joining occurs in only 0.2% of these cells (Moore and Haber, 1996), thus both DSBs are essentially irreparable.

Following the induction of 2 DSBs, we monitored cell cycle arrest in four ways: (1) with an adaptation time-course assay where we micromanipulated individual G₁ cells on agar plates and scored the percentage of cells that are able to re-enter mitosis (Lee et al., 1998), (2) by monitoring the percentage of G₂/M-arrested cells in liquid culture based on cell morphology (Figure 1A), (3) by monitoring nuclear division by DAPI staining of nuclei, and (4) by assaying Rad53 phosphorylation by western blot (Pellicioli et al., 2001).

In both 1-DSB and 2-DSB strains, 4 hr after the induction of DNA damage, >90% of cells arrested at G₂/M as determined by an adaptation assay (Figure 1B and E) and DAPI staining (Figure 1D and G). In agreement, western blot analysis showed that Rad53 was hyperphosphorylated (Figure 1C and F), demonstrating that DDC was fully activated in both these strains after DNA damage. By 12–15 hr after the induction of a single persistent DSB, most 1-DSB cells adapted; that is, they escaped the G₂/M arrest and re-entered mitosis (Figure 1B). The timing of Rad53 dephosphorylation following the induction of a single irreparable DSB correlated with the timing of adaptation and escape from the G₂/M arrest (Figure 1C), as previously shown (Pellicioli et al., 2001) In contrast, in the 2-DSB strain, over 90% of cells remained permanently arrested in G₂/M with persistently hyper-phosphorylated Rad53 throughout the 24 hr time course (Figure 1E–G). We leveraged this permanent cell cycle arrest observed in the 2-DSB strain to study how the DNA damage-induced cell cycle arrest is maintained once it had been established.

We used the AID system (Nishimura et al., 2009) to conditionally deplete DDC proteins after G₂/M arrest had been established to study the maintenance of cell cycle arrest. To this end, we appended an AID tag with nine copies of the c-MYC epitope tag to the C-terminus of Ddc2, Rad9, Rad24, and Rad53, which are all components of the Mec1 signaling cascade (Memisoglu et al., 2019; Sweeney et al., 2005; de la Torre-Ruiz et al., 1998). Hereafter, all AID-tagged proteins will be designated simply as -AID, for example, Ddc2-9xMyc-AID as Ddc2-AID.

AID tagging of DDC proteins did not alter the establishment of G₂/M arrest; however, RAD9-AID, RAD24-AID, and RAD53-AID strains are hypomorphic and adapted 24 hr after DNA damage in the absence of IAA, while 2-DSB wild-type counterpart cells remained fully arrested (Figure 2—figure supplement 1A). This premature escape from the cell cycle arrest was dependent on the presence of TIR1 (Figure 2—figure supplement 1B) and is likely due to low levels of IAA as a natural intermediate in amino acid biosynthesis (Rao et al., 2010). IAA treatment prior to the induction of 2 DSBs in a control strain, which does not contain any AID-tagged proteins, did not alter the prolonged cell cycle arrest following DNA damage (Figure 2—figure supplement 2A), illustrating that IAA treatment by itself does not alter response to DNA damage. However, rapid degradation of Ddc2-AID, Rad9-AID, Rad24-AID, and Rad53-AID by IAA treatment 2 hr prior to the induction of DSBs largely prevented cell cycle arrest as well as DDC signaling, evident from the absence of detectible Rad53 hyperphosphorylation (Figure 2—figure supplement 2B–E). These results underline the importance of Ddc2, Rad9, Rad24, and Rad53 in initiating DDC signaling and cell cycle arrest in response to DNA damage, agreeing with previous reports (Pellicioli et al., 2001; Paciotti et al., 2000; Emili, 1998; de la Torre-Ruiz et al., 1998).

To test whether the DDC proteins are required for the maintenance of the cell cycle arrest following DNA damage, we employed the AID-tagged strains with 2 DSBs and depleted the AID proteins 4 hr after inducing DSBs. In the absence of IAA, DDC2-AID, RAD9-AID, or RAD24-AID strains all activated the DDC signaling 4 hr after DSB induction, with 89–99% of cells arrested in G₂/M (Figure 2A–C), demonstrating that AID tagging of these proteins did not impair their function. Within 1 hr after IAA treatment, Ddc2-AID, Rad9-AID, or Rad24-AID were all rapidly depleted, which caused a gradual Rad53 dephosphorylation, as detected by western blotting (Figure 2A–C). Moreover, agreeing with the loss of Rad53 phosphorylation, IAA treatment of DDC-AID strains triggered release from G₂/M arrest, while the untreated control cells remained fully arrested. DAPI staining of DDC2-AID cells after IAA treatment revealed the accumulation of large-budded cells with two distinct DAPI signals, indicating that cells started to progress into anaphase following Ddc2 depletion (Figure 2—figure supplement 3A). These findings illustrate that the upstream DDC factors Ddc2, Rad9, and Rad24 are essential for initiating and sustaining the cell cycle arrest in response to DNA damage.

Compared to DDC2-AID, RAD9-AID, or RAD24-AID, we found that RAD53-AID cells maintained G₂/M arrest for an additional 4 hr after complete depletion of Rad53 (Figure 2D). In contrast to Ddc2-AID depletion, cell cycle analysis by DAPI staining showed that Rad53 depletion led to a more gradual transition into late anaphase (Figure 2—figure supplement 3B). This delay after the conditional depletion of Rad53 could be due to continued signaling from downstream targets activated by Rad53 kinase, such as Dun1, or from other targets of the Mec1 kinase, downstream of Ddc2, Rad9, and Rad24.

Rad53 and Chk1 kinases both contribute to the maintenance of the cell cycle arrest after DNA damage (Dotiwala et al., 2007; Pellicioli et al., 2001). Agreeing with previously published reports (Sanchez et al., 1999), we find that Chk1 is involved in maintaining the permanent arrest following the induction of 2 DSBs. Deletion of CHK1 did not impair the induction of cell cycle arrest (Figure 3A); however, it inhibited the permanent cell cycle arrest as >95% of chk1∆ cells adapted by 24 hr (Figure 3B). We then asked whether the delay in cell cycle re-entry observed when Rad53 was degraded was due to Chk1’s independent role in maintaining arrest. To test this, we induced 2 DSBs in RAD53-AID chk1∆ cells for 4 hr and then added IAA to deplete Rad53. Compared to the depletion of Rad53-AID alone (Figure 2D), the depletion of Rad53-AID in the absence of CHK1 led to a significant decrease in the number of G₂/M-arrested cells within 1 hr of IAA treatment (Figure 3C). These results suggest that Chk1 functions in conjunction with Rad53 to sustain cell cycle arrest in response to DNA damage.

To explore further how Chk1 signaling contributes to the maintenance of DDC-dependent cell cycle arrest, we depleted DDC factors Ddc2-AID, Rad9-AID, or Rad24-AID in chk1∆ cells 4 hr after the induction of DSBs. Depletion of these upstream factors in the absence of CHK1 led to a more rapid release from the cell cycle arrest (Figure 3D–F) compared to the depletion of DDC factors alone (Figure 2A–C). Collectively, these findings demonstrate that Chk1 plays a key role in maintaining cell cycle arrest in response to DNA damage.

Tel1 is thought to play a minor role in response to DSBs as the establishment of DSB-induced cell cycle arrest normally depends entirely on Mec1 (Dotiwala et al., 2010). However, previous studies have shown that, in addition to Mec1, Tel1 can also target Chk1 for phosphorylation (Limbo et al., 2011; Sanchez et al., 1999). Additionally, when the initial 5’ to 3’ end resection of DSB ends is impaired, Tel1 alone can activate the DDC (Usui and Petrini, 2007). To study how Tel1 contributes to the maintenance of the permanent cell cycle arrest, we deleted TEL1 in 2-DSB strain. Unlike chk1∆ with 2 DSBs, a TEL1 deletion did not affect either the establishment of the DDC or the maintenance of checkpoint arrest up to 24 hr (Figure 3—figure supplement 1), agreeing with previously published results (Dubrana et al., 2007). Taken together, these data illustrate that DNA damage-dependent cell cycle arrest is initiated by Mec1 branch of the DDC via Ddc2, Rad9 and Rad24, and the cell cycle arrest is largely sustained by the downstream kinases Rad53 and Chk1, with minor contributions from other downstream targets of DDC.

Our findings show that a small number of cells remain arrested in the absence of CHK1 when Rad53 is depleted. We posited that Rad53 could modulate the expression of other DDC factors, which, in turn, sustain the cell cycle arrest in the absence of Rad53 and Chk1. One candidate protein is Dun1, a Rad53-activated protein kinase that regulates transcription in response to DNA damage (Chen et al., 2007; Yam et al., 2020; Zhou and Elledge, 1993). Deleting DUN1 significantly impaired checkpoint activation: compared to the wild-type control strain, only 60% of DUN1 cells arrested in G₂/M 4 hr after the DSB induction, and only 25% remained in G₂/M arrest at 7 hr (Figure 4A). Additionally, depletion of Dun1-AID 4 hr after damage induction did not cause a significant change in the proportion of G₂/M arrested in these otherwise wild-type cells, nor did it affect Rad53 phosphorylation (Figure 4B); however, deleting CHK1 triggered an exit from checkpoint arrest following the depletion of Dun1-AID 4 hr after DSB induction (Figure 4C), further demonstrating the role of Chk1 in checkpoint maintenance. These results suggest that Dun1 is required for the initiation of DDC and concomitant cell cycle arrest but is dispensable for maintenance.

Previously, we reported that Ddc2 protein abundance initially increases over time as after the induction of a single DNA break, which is followed by near-complete depletion of Ddc2 around the time that cells adapt (Memisoglu et al., 2019). Given that Ddc2 overexpression leads to permanent cell cycle arrest following DNA damage (Clerici et al., 2001), we concluded that Ddc2 abundance is intimately tied to the duration of the arrest. Here, we asked whether the presence of 2 DSBs instead of a single DSB would lead to an increase in Ddc2 protein abundance and, therefore, the permanent cell cycle arrest. We examined the levels of Ddc2 in both 1- and 2-DSB strains following DNA damage but did not detect a difference in the changes of abundance of Ddc2 protein (Figure 5—figure supplement 1), even though cells adapt to 1 DSBs and remained terminally arrested after 2 DSBs (Figure 1A–D).

If Ddc2 activity is essential to maintain the cell cycle arrest in the 2-DSB strain, then degradation of Ddc2-AID around the time wild-type cells adapt to a single DNA break should interrupt the permanent cell cycle arrest and trigger cell cycle re-entry. We find that complete depletion of Ddc2-AID 15 hr after the induction of 2 DSBs leads to diminished Rad53 hyperphosphorylation (Figure 5A). However, surprisingly, Ddc2-AID degradation did not alter the percentage of G₂/M-arrested cells even 9 hr after Ddc2 depletion (Figure 5A). Furthermore, despite the diminished Rad53 phosphorylation, Ddc2-AID cells mostly remained arrested in G₂/M after the depletion of Ddc2-AID at 15 hr as illustrated by DAPI staining (Figure 5B), in contrast to Ddc2-AID depletion soon after induction of 2 DSBs, which causes cells to rapidly resume mitosis (Figure 2A). These results hint that the maintenance of the permanent cell cycle arrest in response to 2 DSBs at later stages could be independent of the DDC signaling.

We then asked whether other DDC factors such as Rad9, Rad24, and Rad53 are dispensable for the prolonged arrest following the induction of 2 DSBs. However, as noted above, AID-tagged DDC activation proteins were unable to maintain this prolonged arrest in a 2-DSB strain even in the absence of IAA, which precludes their use in this analysis. To be able to study the contribution of these DDC factors in prolonged cell cycle arrest, we turned to the AID version 2 (AID2) system (Yesbolatova et al., 2020). To this end, we integrated a TIR1-F74G point mutation and used 5-phenyl-IAA (5-Ph-IAA) instead of IAA to lower the basal degradation of AID-tagged proteins. Switching to the AID2 system did not fully restore function to RAD9-AID2 and RAD24-AID2 strains as they mostly adapted after 24 hr after the exposure to DNA damage, but 87% of RAD53-AID2 cells remained in G₂/M arrest at 24 hr (Figure 2—figure supplement 1C). Using Rad53-AID2, we then asked whether Rad53 is required for extended G₂/M arrest in response to 2 DSBs. Rapid depletion of Rad53-AID2 with 5-Ph-IAA 4 hr after DSB induction led cells to escape G₂/M arrest, but as with the RAD53-AID strain, we detected a 4 hr delay in cell cycle re-entry, confirming our previous results (Figures 2D and 5C). However, degradation of Rad53-AID2 15 hr after DSB induction did not prompt cell cycle re-entry even 9 hr after complete depletion of Rad53 (Figure 5D, Table 1), akin to what we observe following the depletion of Ddc2-AID (Figure 5A and B).

Because TIR1-mediated degradation of Rad9-AID even without auxin caused most cells to adapt 24 hr after inducing DNA damage, we asked whether overexpression of RAD9-AID could overcome this effect. We added a TRP1 centromere-containing plasmid copy of RAD9-AID with its endogenous promoter (pRAD9-AID) to our 2-DSB RAD9-AID strain. Degradation of Rad9-AID by IAA 15 hr after DSB induction did not trigger release of cells from G₂/M arrest (Figure 5E, Table 1). However, unlike degradation of Ddc2-AID or Rad53-AID2 in this same situation, Rad53 remained hyperphosphorylated up to 9 hr after adding IAA (Figure 5E). Therefore, while the DDC proteins Ddc2, Rad9, and Rad53 are required for the maintenance of checkpoint arrest at early stages, surprisingly, they are dispensable for prolonged arrest following induction of 2 DSBs. These results suggest that prolonged cell cycle arrest is maintained by signaling proteins other than the Mec1 branch of the DDC.

In addition to the DDC, the SAC maintains genomic integrity by halting mitosis at the metaphase/anaphase transition in response to unattached kinetochores, to ensure accurate chromosome segregation (reviewed by McAinsh and Kops, 2023). We have previously shown that inactivation of the SAC by a MAD1, MAD2, or MAD3 deletion shortened the duration of the cell cycle arrest induced by a single DSB (Dotiwala et al., 2010). To test whether SAC is involved in enforcing and sustaining permanent cell cycle arrest in response to 2 DSBs, we deleted MAD2 in the 2-DSB strain. Adaptation assay results illustrate that nearly all mad2∆ cells arrested at 4 hr but began to adapt between 12 and 15 hr after DNA damage (Figure 6—figure supplement 1A). To explore whether deletion of MAD2 can antagonize the permanent cell cycle arrest due to hyperactive DDC signaling, we overexpressed Ddc2 in mad2∆ 2-DSB cells and assayed mitotic progression. We find that both in 1-DSB and 2-DSB strains, MAD2 deletion leads to cell cycle re-entry even when Ddc2 is overexpressed (Figure 6—figure supplement 1B). Based on these data, we concluded that mitotic inhibition is enforced by the SAC proteins as DDC factors become dispensable 12–15 hr after the induction of damage.

If SAC proteins are only required at later stages of cell cycle arrest when DDC proteins are dispensable, then the depletion of SAC protein Mad2 or its binding partner Mad1 soon after the induction of DNA damage should not affect DDC or cell cycle arrest up to 12–15 hr. In agreement with this idea, after Mad1-AID or Mad2-AID depletion at 4 hr, cells remained arrested up to 15 hr following DSB induction, with persistent Rad53 hyperphosphorylation (Figure 6—figure supplement 2). Strikingly, these cells eventually re-entered the cell cycle by 24 hr (20 hr after depletion of Mad1 and Mad2). It is notable that cells resumed cell cycle progression despite persistent Rad53 hyperphosphorylation. This result reinforces our conclusion that Mad1 and Mad2 are not required for the activation and initial maintenance of arrest but are essential for prolonged cell cycle arrest in response to 2 DSBs. These data also suggest that in the absence of Mad1 or Mad2 cells become insensitive to the arrest normally imposed by DDC.

To monitor the effect of SAC proteins Mad1 and Mad2 at late stages of prolonged cell cycle arrest when DDC signaling becomes dispensable, we depleted Mad1-AID or Mad2-AID 15 hr after DSB induction. Following the depletion of Mad1-AID and Mad2-AID, we observed an immediate reduction in the percentage of G₂/M-arrested cells (Figure 6A–C). DAPI staining confirmed that Mad1-AID or Mad2-AID-deplete cells re-entered the cell cycle, evident from an increase in the G1 cell population as well as an increase in percentage of large-budded cells with 2 DAPI foci (Figure 6B and D). Similar to what we observe following Mad1-AID and Mad2-AID depletion 4 hr after damage, this cell cycle re-entry occurred despite persistent Rad53 hyperphosphorylation (Figure 6A and C).

To show that the late stages of the permanent cell cycle arrest in response to 2 DSBs is independent of DDC and dependent on SAC, we inactivated DDC by depleting Ddc2-AID together with Mad2-AID. Although the simultaneous depletion of Ddc2-AID and Mad2-AID led to Rad53 dephosphorylation as detected by western blotting, we found no statistically significant difference in the percentage of cells escaping G₂/M arrest in response to DDC2-AID MAD2-AID double depletion strain compared to Mad2-AID alone (Figure 6—figure supplement 3 and Table 1). Collectively, our findings indicate that DDC initiates and sustains the cell cycle arrest approximately for 15 hr following DNA damage, but after that DDC becomes dispensable and the permanent arrest is sustained by SAC.

To investigate the possible contribution of the MEN to the maintenance of the extended cell cycle arrest in response to 2 DSBs, we appended AID tags to upstream MEN proteins Bub2 and Bfa1. Degradation of Bub2-AID both 4 hr and 15 hr after induction of 2 DSBs suppressed G₂/M arrest (Figure 7A, Figure 7—figure supplement 1A and B), akin to what we observe following Mad1-AID or Mad2-AID depletion (Figure 6A and B). In contrast to Bub2-AID, we see that Bfa1-AID degradation did not trigger a significant release from G₂/M arrest and did not alter the phosphorylation of Rad53 (Figure 7B, Figure 7—figure supplement 1C–E). Analysis of cell cycle distribution with DAPI staining showed that neither the inactivation of Bub2 nor Bfa1 led to the accumulation of a significant number of cells with two separate DAPI-staining nuclei, which is indicative of mitotic exit defects (Figure 7B and D, Figure 7—figure supplement 1B, D, and F). These results imply that although Bub2 and Bfa1 have interdependent functions for MEN signaling, they carry out independent roles in response to DNA damage.

In contrast to the permanent cell cycle arrest we observe in response to 2 DSBs, a recent study using two HO-mediated persistent DSBs showed that cells in fact adapt (Sadeghi et al., 2022). One difference between these two 2-DSB systems is the relative position of the DSBs, which might affect how SAC components become engaged, and thus might determine the extent of mitotic arrest. Supporting this, we previously showed that deleting CEN3 in a strain with a DSB at MAT on chromosome III eliminated the Mad2-dependent delay in adaptation, but deleting CEN3 when the DSB was on chromosome VI had no effect (Dotiwala et al., 2010). In our adaptation-defective 2-DSB strain, the DSBs are located at MAT (86 kb from CEN3) and near FAB1 (42 kb from CEN6). Sadeghi et al. employed two strains, both of which escape prolonged G₂/M arrest, with at least one DSB site far from its centromere; at URA3 (36 kb from its centromere) and ADH1 (170 kb) or at MIC2 (32 kb) and DLD2 (316 kb).

To investigate whether the distance between the second DSB site and the centromere would affect whether cells will remain permanently arrested, we created several strains that contain a second cut site at various distances from the centromere, in addition to the cut site at MAT on chromosome III (Figure 8). Strain YSL53, which has a second DSB at chromosome V, 86 kb away from CEN5 (Lee et al., 1998), and strain DW417, which has a second DSB 52 kb away from CEN6 (Lee et al., 2014), mostly remained in G₂/M arrest 24 hr after the induction of DNA breaks. However, in GEM188, which had a second cut site 230 kb away from the CEN2, only 37% of cells remained in G₂/M arrest by 24 hr. Thus, the increased distance of the second DSB site to the centromere in GEM188 appears to have led to a less robust triggering of the SAC compared with YSL53 and DW417.

Our previous data have suggested that the involvement of the SAC in prolonging DSB-induced arrest was dependent on centromere sequences on the broken chromosome and involved post-translational modification of chromatin by the Mec1- and Tel1-dependent phosphorylation of the histone H2A (Dotiwala et al., 2010). In budding yeast, histone H2B is also targeted by DDC kinases upon DNA damage (Lee et al., 2014). To test whether the presence of these chromatin modifications around centromeres would be sufficient to elicit a SAC response, we examined cell cycle progression in strains in which both histone H2A and/or histone H2B genes were mutated to their putative phosphomimetic forms (H2A-S129E and H2B-T129E). We note that although histone H2A-S129E is recognized by an antibody specific for the phosphorylation of histone H2A-S129 (Eapen et al., 2012), the mutation to S129E may not be fully phosphomimetic. As shown in Figure 9, there was no effect on the growth rate of either the single or the double mutants, suggesting that cells did not experience a SAC-dependent delay in entering mitosis because of these modifications.

All AID-tagged mutant strains were derived from a modified version of strains JKM179. To create the strain with two HO cleavage sites (DW417), an HO-cut site, designated HOcse6, with an adjacent HPH marker was integrated into chromosome VI, 52 kb from the centromere. To create AID strains, we first integrated osTIR1 at URA3 after digesting plasmid pNHK53 (Nishimura et al., 2009) with StuI. To integrate osTIR1-F74G at URA3, the plasmid pMK420 (Yesbolatova et al., 2020) was digested with StuI. For degron-tagging of DDC proteins, AID-9xMyc (AID) PCR products were generated with mixed oligos with homology to the C-terminal end of the corresponding open-reading frames by using plasmids pKan–AID–9xMyc (pJH2892) or pNat–AID–9xMyc (pJH2899) as templates (Morawska and Ulrich, 2013). Deletion of ORFs and insertion of AID tags were introduced with the one-step PCR homology cassette amplification and the standard yeast transformation method (Wach et al., 1994). Cas9 editing was done by inserting a gRNA into plasmid bRA90 (Anand et al., 2017) and co-transformed into our strain of interest with a donor sequence. Transformants were verified by PCR, western blotting, and sequencing. To create strain GEM188 with 2-DSBs, we inserted a second HO-cut site into JKM179 at LYS2 locus by CRISPR/Cas9 (Anand et al., 2017) using a synthetic DNA template with 117 bp consensus HO recognition site. Non-phosphorylatable and phosphomimetic mutants of H2A were generated in a JKM179 background using CRISPR/Cas9 to target HTA1 and HTA2 genes at serine 129 and 80 nt templates to mutate serine to either alanine (non-phosphorylatable) or glutamic acid (phosphomimetic). H2B mutants were generated in a JKM179 background using CRISPR/Cas9 to target HTB1 and HTB2 genes at threonine 129 with 80 nt repair templates to mutate threonine to either alanine or glutamic acid.

The CEN/ARS plasmid pFZ052-pRAD9-AID-*9Myc::Trp1 (pRAD9-AID) was obtained by digesting the plasmid pFL36.1 (Lazzaro et al., 2008) with SmaI and AscI to excise the 3 HA tag on the C-terminal end of Rad9. A 9xMyc-AID PCR product generated from the plasmid pJH2892 (pKan-9xMyc-AID) was cut with AscI and sticky/blunt end cloned into the SmaI-AscI-digested pFL36.1 to add the 9xMyc-AID tag to the C-terminal end of Rad9. pRad9-AID was retained by growing cells in a Trp- media with 2% raffinose. The primers used for strain and plasmid creation are listed in the Key Resources Table, Supplementary file 1a, and Supplementary file 1b. Plasmids are listed in the Key Resources Table.

Strains containing degron fusions and galactose-inducible HO were cultured using standard procedures. Briefly, a single colony grown on a YEPD plate (1% yeast extract, 2% peptone, 2% dextrose, 2.5% agar) was inoculated in 5 ml YEP-lactate (YEP containing 3% lactic acid) and was grown for ~15 hr at 30°C with agitation. Next day, the overnight culture was used to inoculate a 500–100 ml of YEP-lactate culture such that the cell density reached an OD₆₀₀ of 0.5 the following day. After harvesting 15 ml liquid culture before treatment, HO expression was induced by galactose treatment with a 2% final concentration. Then, cultures were split either at 4 hr or 15 hr following induction with galactose. The split cultures were treated either with IAA or 5-Ph-IAA or an equivalent volume of 200 proof ethanol. IAA (Sigma-Aldrich, I3750) was resuspended in ethanol for a 500 mM stock and used at a 1 mM final concentration both for liquid media and for agar plates. 5-Ph-IAA was dissolved in ethanol for a 1 mM stock and used at a final concentration of 1 µM. 15 ml of liquid culture were harvested at various timepoints and prepared for microscopy or western blot analysis as described below.

To measure growth rate, strains were grown in 5 ml of YEPD with 2% dextrose at 30°C with an initial OD₆₀₀ of 0.1. The OD was measured 3, 5, 7, and 10 hr after the initial OD measurement using a Thermo Scientific NanoDrop 2000c Spectrophotometer. To measure the OD, 50 µl of culture was added to 950 µl of fresh YPD in a cuvette at a dilution of 1:20.

Protein extracts were prepared for western blot analysis by TCA extraction protocol as previously explained (Miller-Fleming et al., 2014). Briefly, 15 ml of liquid culture was spun down and the media was discarded. Harvested cells were incubated on ice in 1.5 ml microcentrifuge tubes with 20% TCA for 20 min. Cells were washed with acetone and the pellet was air-dried. 200 µl of MURBs buffer (50 mM sodium phosphate, 25 mM MES, 3 M urea, 0.5% 2-mercaptoethanol, 1 mM sodium azide, and 1% SDS) was added to each sample with acid-washed glass beads. Cells were lysed by mechanical shearing with glass beads for 2 min. Crude cell lysates were harvested by poking a hole in the bottom of the 1.5 ml microcentrifuge tube and spinning the tubes on a 15 ml conical tube. Samples were boiled at 95°C for 10 min prior to loading on SDS-PAGE.

8–20 µl of denatured protein samples prepared by TCA extraction were loaded onto a 10% or 8% SDS-PAGE gels. Proteins were separated by applying constant voltage at 90 V until the 37 kDa protein standard band reached the bottom of the gel. Gels were transferred to an Immun-Blot PVDF membrane (Bio-Rad) using a wet transfer apparatus set to 100 V constant voltage for 1 hr at 4°C. Membranes were then blocked with OneBlock blocking buffer (Genesee Scientific, Cat# 20-313) for 1 hr at room temperature, After three 10 min washes with 1× TBS-T, blots were incubated with either anti-Myc [9E11] (Abcam, ab56) to detect TIR1 and AID fusions, anti-Rad53 [EL7.E1] (Abcam, ab166859), anti-Pgk1 (Abcam, ab30359), or anti-Rad9 (Usui et al., 2009) for 1 hr at room temperature or at 4°C overnight. Blots were washed three times with 1× TBS-T and incubated with anti-mouse HRP (GE Healthcare, Cat# NXA931) or anti-rabbit HRP secondary antibody (Sigma-Aldrich, Cat# A6154) for 1 hr at room temperature. After washing the membranes three times with 1× TBS-T, Amersham ECL Prime Western Blotting Detection Reagent was added to fully coat the blots and left to incubate for 5 min at room temperature with gentle agitation. Blots were imaged using a Bio-Rad ChemiDoc XR+ imager and prepared for publication using ImageLab 6.1 software (Bio-Rad) and Adobe Photoshop CC 2017. The reagents used are listed in the Key Resources Table.

Aliquots from YEP-Lac cultures were taken either 4 hr or 15 hr after adding galactose, diluted 20-fold in sterile water, and plated on a YEP-Agar with 2% galactose with or without 1 mM IAA or 1 µM 5-Ph-IAA. Cells were counted on a light microscope with a ×10 objective, examined, and binned into three categories: unbudded, small buds, and G₂/M-arrested cells with large buds. For each timepoint, >250 cells were analyzed. For DAPI staining, 450 µl of culture was added to 50 µl of 37% formaldehyde and incubated in the chemical hood at room temperature for 20 min. Samples were spun down at 8000 rpm for 5 min and washed with 1× PBS three times. Cells were resuspended in 50 µl of DAPI mounting media (VECTASHIELD Antifade Mounting Medium with DAPI H-1200-10) and incubated at room temperature for 10 min, away from direct light. The samples were imaged by using a Nikon Ni-E upright microscope equipped with a Yokogawa CSU-W1 spinning-disk head, an Andor iXon 897U EMCCD camera, Nikon Elements AR software, a ×60 oil immersion objective, and a 358 nm laser. Fifteen z-stacks with a thickness of 0.3 µm were collected per image. In the morphology assays, at least three biological replicates were used for each strain.

We performed adaptation assays as previously described (Eapen et al., 2012; Lee et al., 1998). Cells grown in YEP-Lac overnight were diluted 20-fold in sterile water and plated on a YEP-agar plate containing 2% galactose. Using micromanipulation, 50 G₁ cells were isolated and positioned in a grid followed by incubation at 30°C. To quantify the percentage of adapted cells, the number of cells that re-entered cell cycle and grew to a microcolony (3+ cells) after 24 hr was divided by the total number of cells. For auxin plating assays, damage was induced in a YEP-Lac liquid culture by adding galactose at a final concentration of 2%, as described above. Cells were then transferred onto YEP-agar plates containing 2% galactose and 1 mM IAA or 1 µM 5-Ph-IAA 4 hr or 15 hr after adding galactose. For each timepoint, >250 cells were scored and categorized as described above for the adaptation assay from at least three biological replicates.

Graphs were prepared using GraphPad Prism 10 (Dotmatics). Statistical analysis for differences in the percentage of large budded (G₂/M-arrested) cells at different timepoints listed in Table 1 was done using a one-way ANOVA in GraphPad Prism 10. Protein quantification of Ddc2-myc blots was done using ImageLab 6.1 (Bio-Rad). To categorize DAPI-stained cells based on their morphology and number of DAPI signals, images were captured as described above and viewed using ImageJ with the Fiji addon.