DNA is exposed to both endogenous and exogenous DNA damaging agents that chemically modify it. To counteract the deleterious effects exerted by DNA lesions, eukaryotic cells have evolved a network of cellular pathways, termed DNA damage response (DDR). The DDR comprises both mechanisms devoted to repair DNA lesions and signal transduction pathways that sense DNA damage and transduce this information to specific cellular targets. These targets, in turn, impact a wide range of cellular processes including DNA replication, DNA repair and cell cycle transitions. The importance of the DDR is highlighted by the fact that DDR inactivation is commonly found in cancer and causes many different human diseases. The protein kinases ATM and ATR, as well as their budding yeast orthologs Tel1 and Mec1, act as master regulators of the DDR. The initiating events in the DDR entail both DNA lesion recognition and assembly of protein complexes at the damaged DNA sites. Here, we review what is known about the early steps of the DDR.

Introduction

Each cell in the human body is subjected to tens of thousands of DNA lesions per day. DNA damage can occur spontaneously during DNA metabolism or they can be induced by environmental agents [1]. The majority of endogenous DNA damage arises from hydrolytic and oxidative reactions of DNA with water and reactive oxygen species that are naturally present within a cell. Other spontaneous DNA alterations can be due to deoxyribonucleoside triphosphate (dNTP) or ribonucleoside triphosphate (rNTP) misincorporation during DNA replication, modification of DNA bases by alkylation, loss of DNA bases by depurination, interconversion between DNA bases by deamination, or single- and double-strand breaks (DSBs) produced, for example, by abortive topoisomerase actions. Exogenous DNA damage can be produced either by chemicals, such as alkylating and cross-linking agents, or by ultraviolet (UV) and ionizing (IR) radiations (Figure 1). These DNA lesions can compromise the survival of cells and organisms by blocking DNA transcription and replication. Furthermore, lesions that are either not repaired or unproperly repaired can lead to genome instability by causing mutations or genome aberrations.

Schematic representation of the major DNA lesions induced by external and endogenous factors, the DNA repair pathways and the related sensor proteins.

Figure 1.
Schematic representation of the major DNA lesions induced by external and endogenous factors, the DNA repair pathways and the related sensor proteins.

MMR, mismatch repair; RER, ribonucleotide excision repair; BER, base excision repair; NER, nucleotide excision repair; SSB repair, single-strand break repair; HR, homologous recombination; NHEJ, non-homologous end joining.

Figure 1.
Schematic representation of the major DNA lesions induced by external and endogenous factors, the DNA repair pathways and the related sensor proteins.

MMR, mismatch repair; RER, ribonucleotide excision repair; BER, base excision repair; NER, nucleotide excision repair; SSB repair, single-strand break repair; HR, homologous recombination; NHEJ, non-homologous end joining.

To counteract threats posed by DNA damage, cells have evolved mechanisms which detect DNA lesions, signal their presence and promote their repair [2,3]. These mechanisms are collectively termed the DNA damage response (DDR). The DDR is a spatiotemporally regulated process, where proteins involved in DNA repair and checkpoint are assembled at the sites of DNA damage in a sequential and co-ordinated manner to form discrete nuclear foci (Table 1) [2,3]. Hereditary defects in the DDR cause a variety of human diseases that are associated with cancer predisposition, developmental defects, infertility, immune deficiency, neurodegeneration and accelerated aging [4–6]. Localization of the DDR proteins to the sites of damage is initiated by sensor proteins that directly recognize DNA lesions and trigger a broad spectrum of post-translational modifications. Here, we review the initial recognition events that occur in the DDR.

Table 1.
Conserved DNA damage response proteins
FunctionS. cerevisiaeS. pombeH. sapiens
Checkpoint kinase Mec1 Rad3 ATR 
Mec1/Rad26/ATR interacting protein Ddc2 Rad26 ATRIP 
Checkpoint kinase Tel1 Tel1 ATM 
9–1–1 checkpoint clamp Ddc1–Rad17–Mec3 Rad9–Rad1–Hus1 RAD9–RAD1–HUS1 
9–1–1 checkpoint clamp loader Rad24–RFC Rad17–RFC RAD17–RFC 
DSB signaling; Tel1/ATM activator Mre11–Rad50–Xrs2 Mre11–Rad50–Nbs1 MRE11–RAD50–NBS1 
Activator of Mre11 nuclease Sae2 Ctp1 CtIP 
DNA-end binding complex Ku70/ku80 Ku70/Ku80 KU70/KU80 
Checkpoint mediator Mrc1 Mrc1 Claspin 
Checkpoint mediator; resection inhibitor Rad9 Crb2 53BP1 
Mec1/ATR activator Dpb11 Cut5/Rad4 TopBP1 
Nuclease involved in resection Exo1 Exo1 EXO1 
Helicase involved in resection Sgs1 Rqh1 BLM 
Nuclease involved in resection Dna2 Dna2 DNA2 
Effector kinase Rad53 Cds1 CHK2 
Effector kinase Chk1 Chk1 CHK1 
Chromatin remodeling Fun30 Fft3 SMARCAD1 
Scaffold protein; checkpoint inhibitor Slx4 Slx4 SLX4 
FunctionS. cerevisiaeS. pombeH. sapiens
Checkpoint kinase Mec1 Rad3 ATR 
Mec1/Rad26/ATR interacting protein Ddc2 Rad26 ATRIP 
Checkpoint kinase Tel1 Tel1 ATM 
9–1–1 checkpoint clamp Ddc1–Rad17–Mec3 Rad9–Rad1–Hus1 RAD9–RAD1–HUS1 
9–1–1 checkpoint clamp loader Rad24–RFC Rad17–RFC RAD17–RFC 
DSB signaling; Tel1/ATM activator Mre11–Rad50–Xrs2 Mre11–Rad50–Nbs1 MRE11–RAD50–NBS1 
Activator of Mre11 nuclease Sae2 Ctp1 CtIP 
DNA-end binding complex Ku70/ku80 Ku70/Ku80 KU70/KU80 
Checkpoint mediator Mrc1 Mrc1 Claspin 
Checkpoint mediator; resection inhibitor Rad9 Crb2 53BP1 
Mec1/ATR activator Dpb11 Cut5/Rad4 TopBP1 
Nuclease involved in resection Exo1 Exo1 EXO1 
Helicase involved in resection Sgs1 Rqh1 BLM 
Nuclease involved in resection Dna2 Dna2 DNA2 
Effector kinase Rad53 Cds1 CHK2 
Effector kinase Chk1 Chk1 CHK1 
Chromatin remodeling Fun30 Fft3 SMARCAD1 
Scaffold protein; checkpoint inhibitor Slx4 Slx4 SLX4 

DNA repair pathways

To respond to the wide variety of DNA lesions, eukaryotic cells have evolved repair mechanisms that detect and remove alterations in the DNA chemistry, so that the integrity of the genome is restored (Figure 1) [7]. In particular, in addition to the direct reversal of some types of damage (such as the enzymatic photoreactivation of thymine dimers), cells possess multiple distinct mechanisms for excising damaged DNA regions and inserting new nucleotides to fill the gap, followed by ligation of the DNA pieces [7]. In case cells suffer breakage of both DNA chains, these DNA breaks are repaired by mechanisms that, based on their homology requirements, can be divided into non-homologous end joining (NHEJ), single-strand annealing (SSA) relying either on microhomology or extensive homology, and homologous recombination (HR) [8,9]. While NHEJ involves the direct ligation of DNA ends with little or no homology [9], HR refers to the exchange or transfer of information between DNA sequences with perfect or near-perfect homology [8]. The choice among these DSB repair pathways depends on the structure of the DNA ends and on the phase of the cell cycle.

Excision repair

In excision repair, biochemically distinct events excise from the DNA molecule different types of DNA damage that do not affect the sugar-phosphate backbone [7]. Removal of these lesions leaves DNA gaps that are filled-in by DNA synthesis. Since these processes use the complementary strand and not a sister chromatid, the repair can occur in any phase of the cell cycle. Excision repair pathways include base excision repair (BER), nucleotide excision repair (NER), ribonucleotide excision repair (RER) and mismatch repair (MMR) (Figure 1).

BER excises uracil and other inappropriate bases from the DNA, as well as many types of chemically modified bases, such as alkylated bases [10]. During BER, the damaged base is recognized by a specific DNA glycosylase enzyme that catalyzes base removal by hydrolyzing the N-glycosidic bond. The resulting abasic (AP) site is further processed by an AP endonuclease, which cleaves the DNA backbone 5′ to the abasic site, followed by completion of DNA repair through the action of nuclease, polymerase and ligase proteins. There are at least 11 nuclear DNA glycosylases in humans, and each recognizes a specific group of modified bases or mismatches [11].

NER is the process that leads to the excision of a variety of helix-distorting DNA lesions, including UV-induced 6–4 pyrimidine–pyrimidone photoproducts and cyclobutane pyrimidine dimers (CPDs), as well as bulky chemical adducts resulting from aromatic hydrocarbons or cisplatin exposure [12]. During NER, more than 30 different proteins are recruited in a sequential manner to the site of DNA damage, followed by DNA incision on both sides of the lesion, excision of a single-stranded oligonucleotide containing the damage, repair synthesis and DNA ligation. NER can operate via two subpathways depending on how the lesions are initially recognized: transcription-coupled NER (TC-NER), which removes lesions from the transcribed strand of active genes, and the global genome NER (GG-NER), which can repair lesions at any location in the genome [13]. Lesions in TC-NER are detected by stalled RNA polymerases at the damaged sites, followed by recruitment of TC-NER specific factors. In humans, recognition of DNA lesions in GG-NER relies on dedicated protein complexes such as XPC–RAD23B–CETN2 and UV-DDB. Subsequent recruitment of XPA, RPA, XPF–ERCC1 and XPG proteins allows the assembly of an incision complex, with the structure-specific XPF–ERCC1 and XPG endonucleases making incisions on the damaged strand 5′ and 3′ to the lesion, respectively [12]. Interestingly, the Saccharomyces cerevisiae ortholog of mammalian XPC, Rad4, was proposed to slide on the DNA scanning for intactness of the DNA structure via bending/twisting [14–17]. The crystal structure of Rad4 bound to DNA containing a CPD shows that Rad4 inserts a β-hairpin into the DNA helix, causing the flipping out of the two residues containing the CPD lesion [18]. In this recognition structure, Rad4 specifically interacts with the two nucleotides opposite the CPD in the nondamaged strand. Such an indirect mode of recognition might explain the ability of Rad4/XPC to recognize different types of DNA lesions.

RER is a repair pathway dedicated to eliminate ribonucleoside monophosphates (rNMPs) that are erroneously incorporated by DNA polymerases during semi-conservative DNA replication [19,20]. During canonical RER, the RNase H2 enzyme, a metal-dependent endonuclease present in all branches of life, recognizes the junction between the ribonucleotide and the deoxyribonucleotide (RpD) and hydrolyzes at the 5′-end of the ribonucleotide, leaving it attached to the 5′-end of the DNA. The strand displacement activity of DNA polymerase δ starting from the nick creates a DNA flap containing the rNMP, which is then removed by a flap endonuclease, followed by sealing of the nick by a DNA ligase [21]. The crystal structure of Thermotoga maritima RNase H2 in complex with a DNA molecule containing a single ribonucleotide revealed a crucial role of a conserved tyrosine residue in the recognition and hydrolysis of the RpD structure [22]. This residue displaces the strand to be cleaved and allows the phosphate group between the ribose and deoxyribose to participate in the coordination of the divalent metal ion that is required for catalysis.

MMR detects and corrects mispaired DNA bases, as well as mono-, di- and trinucleotide insertion/deletion loops that cause expansion or contraction of DNA repeats [23]. In Escherichia coli, the newly synthesized strand remains transiently unmethylated within the GATC sequences. Then, the mismatch-activated MutS–MutL complex licenses MutH to incise the unmethylated GATC sequence. The generation of the MutH-catalyzed nick allows the loading of a helicase and of an exonuclease, which leads to the degradation of the error-containing strand beyond the mismatch in order to generate a single-stranded gap that is subsequently filled-in by the DNA polymerase III [24]. In eukaryotes, DNA methylation is not used for strand discrimination. Instead, members of the MutS homolog (MSH) protein family recognize the mismatch by forming an ATP-bound sliding clamp that moves along the DNA. Association of the mismatch-activated MLH/PMS proteins with proliferating cell nuclear antigen (PCNA) bound to the 3′ terminus of the leading strand activates a cryptic MLH/PMS endonuclease activity, which introduces a nick into the newly synthesized strand. This nick provides an entry site for nucleases that degrade the error-containing strand [25].

DNA double-strand breaks repair

DNA DSBs can be repaired by either NHEJ, which directly re-ligates the broken DNA ends, or HR, which uses homologous DNA on sister chromatids or homologous chromosomes as a template for repair [8,9]. In NHEJ, DSBs are recognized by the Ku70-Ku80 heterodimer, followed by ligation of the broken DNA ends by DNA ligase IV (Dnl4-Lig4 in yeast) [9]. If the DSB is not repaired by NHEJ, the 5′ strands at either side of the DSB are subjected to 5′–3′ nucleolytic degradation in a process called DNA end resection [26]. This degradation results in the formation of 3′-ended single-stranded DNA (ssDNA) ends that inhibit NHEJ and channel DSB repair into HR. Not surprisingly, resection is regulated during the cell cycle to ensure that commitment to HR occurs primarily in S and G2 phases of the cell cycle when a sister chromatid is available as a repair template [26].

During HR, the ssDNA is first coated by the ssDNA binding protein complex replication protein A (RPA). RPA is subsequently replaced by Rad51 to form a nucleoprotein filament that is used to search and invade a homologous duplex DNA, followed by resolution of the resulting cruciform DNA structures and DNA ligation [8,27].

The highly conserved MRX/MRN complex (Mre11–Rad50–Xrs2 in budding yeast; MRE11–RAD50–NBS1 in mammals) is rapidly recruited to DSBs [28], where it exerts several functions, including DSB detection, processing and signaling [29,30]. While Xrs2/NBS1 is only present in eukaryotes, orthologs of Rad50 and Mre11 are found in all kingdoms of life. Mre11 exhibits both 3′–5′ exonuclease and endonuclease activities [31,32]. In both yeast and mammals, the endonuclease activity of Mre11 is activated by the Sae2 (human CtIP) protein in order to cleave the 5′-terminated DNA strands at both DNA ends [33]. This step is followed by 3′–5′ nucleolytic degradation by Mre11, which proceeds back towards the DNA ends [34,35]. The resulting nick/gap provides an entry site for either the Exo1 (human EXO1) exonuclease or the combined activities of the Sgs1 (human BLM) helicase and the Dna2 (human DNA2) nuclease, which degrade DNA in the 5′–3′ direction away from the DSB ends [34–43] (Figure 2).

Model for sensing and processing DNA DSBs.

Figure 2.
Model for sensing and processing DNA DSBs.

When a DSB occurs, MRX, Sae2 and Ku are rapidly recruited to the DSB ends. MRX is required for the recruitment at the DSB of Tel1, which in turn stabilizes MRX retention at the DSB (double arrows). Upon ATP hydrolysis by Rad50, Mre11, with the support of Sae2, catalyzes an endonucleolytic cleavage of the 5′ strand. This incision allows processing by Exo1 and Sgs1–Dna2 in a 5′–3′ direction from the nick and by Mre11 in a 3′–5′ direction toward the DSB ends. The initiation of DSB resection leads to the removal of Ku from the DSB. The resulting 3′-ended ssDNA attenuates Tel1 signaling activity and, once coated by RPA, allows the recruitment of Mec1–Ddc2. Initiation of DSB resection also promotes the association to the ssDNA/dsDNA junctions of 9–1–1 and Dpb11, which promote the recruitment of Rad9 at the DSB. Rad9, in turn, transduces the checkpoint signal from Mec1 to Rad53 and limits the resection activity of Dna2–Sgs1. The inhibitory function of Rad9 on Sgs1–Dna2 is counteracted by Fun30. Rad53 activation by Rad9 leads to phosphorylation and inhibition of Exo1. Red dots indicate phosphorylation events. S. cerevisiae proteins are indicated.

Figure 2.
Model for sensing and processing DNA DSBs.

When a DSB occurs, MRX, Sae2 and Ku are rapidly recruited to the DSB ends. MRX is required for the recruitment at the DSB of Tel1, which in turn stabilizes MRX retention at the DSB (double arrows). Upon ATP hydrolysis by Rad50, Mre11, with the support of Sae2, catalyzes an endonucleolytic cleavage of the 5′ strand. This incision allows processing by Exo1 and Sgs1–Dna2 in a 5′–3′ direction from the nick and by Mre11 in a 3′–5′ direction toward the DSB ends. The initiation of DSB resection leads to the removal of Ku from the DSB. The resulting 3′-ended ssDNA attenuates Tel1 signaling activity and, once coated by RPA, allows the recruitment of Mec1–Ddc2. Initiation of DSB resection also promotes the association to the ssDNA/dsDNA junctions of 9–1–1 and Dpb11, which promote the recruitment of Rad9 at the DSB. Rad9, in turn, transduces the checkpoint signal from Mec1 to Rad53 and limits the resection activity of Dna2–Sgs1. The inhibitory function of Rad9 on Sgs1–Dna2 is counteracted by Fun30. Rad53 activation by Rad9 leads to phosphorylation and inhibition of Exo1. Red dots indicate phosphorylation events. S. cerevisiae proteins are indicated.

In vitro, the efficiency of the endonucleolytic cleavage catalyzed by MRX–Sae2 was shown to be strongly enhanced by the presence of protein blocks at DNA ends [33,44,45]. The nature of the block does not seem to be important, at least in vitro. In fact, such blocks can be, for instance, the Ku complex bound to DNA ends, a trapped topoisomerase, the RPA complex bound to either partially resected DNA ends or terminal hairpin structures, or even streptavidin bound to biotinylated DNA termini [33–35,46–50]. As the presence of protein-bound DNA molecules can make the DNA ends refractory to Exo1- and Sgs1–Dna2-mediated resection, the DNA-nicking and chew-back action of MRX–Sae2 at protein-bound DNA ends can be important to allow the access to the DSB end of the long-range resection machinery to initiate recombination.

Each Rad50 polypeptide possesses N- and C-terminal domains that assemble to form two complete ATPase sites on a Rad50 dimer [51]. Furthermore, Rad50 possesses long coiled coils that can form proteinaceous rings or rods, joined by a zinc hook at the apex [52–56]. The ATPase domains of Rad50, along with the Mre11 nuclease and DNA binding domains, form the catalytic core of the complex [57,58]. Structural studies of Mre11–Rad50 from bacteria and archaea have shown that, upon ATP binding by Rad50, this catalytic core retains a closed conformation in which the double-stranded DNA (dsDNA) is inaccessible to the Mre11 nuclease active site. In the ATP-hydrolyzed state, the Rad50 subunits are relatively open, suggesting that ATP hydrolysis drives the disengagement of the Rad50 dimer, which would allow the Mre11 nuclease active site to access the DNA [57–63]. However, a recent cryo-electron microscopy analysis of the E. coli Mre11–Rad50 complex has shown that DNA binding induces a conformational change in which the two Rad50 coiled-coils zip up and form a narrow clamp around the DNA molecule [64]. This structural change allows Mre11 dimer to move from the bottom to one side of Rad50 and to bind the DNA.

As described above, the Sae2/MRX-mediated nick generates an entry site for the resection nucleases Dna2 and Exo1, which degrade DNA in the 5′ to 3′ direction [34–43]. The nucleolytic processing catalyzed by these enzymes is negatively regulated by nucleosomes [65] and by nucleosome-associated factors, such as the budding yeast Rad9 protein or its functional human orthologue 53BP1 [66–72] (Figure 2). Rad9, which was originally identified as a component of the DNA damage checkpoint pathway [73–75], is bound to chromatin by an interaction with methylated histone H3 (H3-K79me) [76–78]. Its binding to the sites of damage is strengthened by an interaction with histone H2A (human H2AX histone variant) after it has been phosphorylated on serine 129 (γH2A) [79–84]. Finally, in both yeast and humans, recruitment of Rad9/53BP1 to the sites of DNA damage also relies on its interaction with the scaffold protein Dpb11 (human TopBP1) [85–87]. In yeast, this interaction depends on Rad9 phosphorylation at S462 and T474 residues [85,86] and a phosphorylation-dependent interaction can be detected also between human 53BP1 and TopBP1 [88] (Figure 3).

Mec1/ATR activators.

Figure 3.
Mec1/ATR activators.

The 9–1–1 complex, which is loaded to ssDNA/dsDNA junctions by the Rad24–Rfc2–5 clamp loader, promotes the localization of Dpb11 to DSBs. Dpb11, in turn, drive the association to DSBs of Rad9, which also interacts with modified histones. Rad9 acts as a resection barrier and its inhibitory function is counteracted by Fun30 and Slx4. Slx4 also inhibits Rad53 activation. Dna2 and Dpb11 stimulate Mec1 kinase activity. The Ddc1 subunit of the 9–1–1 complex directly stimulates Mec1 activation at least in budding yeast. Red dots indicate phosphorylation events. S. cerevisiae proteins are indicated.

Figure 3.
Mec1/ATR activators.

The 9–1–1 complex, which is loaded to ssDNA/dsDNA junctions by the Rad24–Rfc2–5 clamp loader, promotes the localization of Dpb11 to DSBs. Dpb11, in turn, drive the association to DSBs of Rad9, which also interacts with modified histones. Rad9 acts as a resection barrier and its inhibitory function is counteracted by Fun30 and Slx4. Slx4 also inhibits Rad53 activation. Dna2 and Dpb11 stimulate Mec1 kinase activity. The Ddc1 subunit of the 9–1–1 complex directly stimulates Mec1 activation at least in budding yeast. Red dots indicate phosphorylation events. S. cerevisiae proteins are indicated.

In budding yeast, Rad9 acts as a resection barrier by controlling at least two different mechanisms (Figure 2). One of them relies on Rad9-mediated activation of Rad53, which promotes phosphorylation and inhibition of Exo1 [89,90]. The other mechanism relies on the inhibition of Sgs1–Dna2 resection activity. In fact, the lack of Rad9 increases the resection efficiency [67,69] and this enhanced resection activity is mainly dependent on Sgs1, whose recruitment at DSBs is inhibited by Rad9 [91,92]. Further support for a Rad9-mediated inhibition of Sgs1 comes from the recent identification of a hypermorphic allele of SGS1 (sgs1-G1298R), which behaves like a rad9Δ phenocopy with respect to the resection efficiency because it escapes Rad9-mediated inhibition [91].

Interestingly, the inhibitory function of Rad9 on DSB resection is counteracted by the ATP-dependent chromatin remodeler Fun30 [93–95] (Figures 2 and 3). Also, the Fun30 human ortholog, SMARCAD1, antagonizes 53BP1 and promotes DNA end resection, suggesting evolutionary conservation [95–97]. The molecular mechanism of the antagonism between Rad9/53BP1 and Fun30/SMARCAD1 is currently unknown. Fun30 might overcome the inhibitory function of Rad9 on DSB resection either by directly removing Rad9 from the DNA damage sites or by interfering with Rad9 recruitment. Interestingly, in both yeast and mammals, Rad9/53BP1 and Fun30/SMARCAD1 interact with Dpb11/TopBP1 and the formation of these complexes seems to facilitate the localization of Rad9/53BP1 and Fun30/SMARCAD1 to damaged chromatin [97]. Fun30 and Rad9 seem to share the binding site for Dpb11, suggesting that a direct competition for Dpb11 binding between Fun30 and Rad9 can contribute to their antagonism [86,87,96].

DNA damage checkpoint

Damaged DNA also elicits a cellular response, termed DNA damage checkpoint, which activates an elaborate signaling network that regulates a variety of cellular processes such as DNA replication, repair and cell cycle transitions [2,3]. Furthermore, if the damage cannot be removed, chronic DDR signaling triggers cell death by apoptosis or terminal differentiation through senescence [2,3].

Upon DNA damage recognition, the checkpoint response induces a signaling cascade driven by protein phosphorylation, which promotes DNA damage repair aided by a slowing down of cell cycle progression. The mammalian proteins ATM (ataxia–telangiectasia-mutated) and ATR (ATM- and Rad3-related), as well as their S. cerevisiae orthologs Tel1 and Mec1, respectively, are the most upstream checkpoint kinases that directly recognize aberrant DNA structures and activate the checkpoint cascade [98]. They belong to a family of phosphatidylinositol 3-kinase (PI3K)-like kinases that also includes DNA-dependent protein kinase (DNA-PK), the mammalian target of rapamycin (mTOR), TRRAP/Tra1 and SMG1. All these PI3K kinases contain a canonical two-lobed kinase domain. The smaller N-lobe contains a glycine-rich loop and contributes to nucleotide binding, while the larger C-lobe possesses the catalytic and activation loops [98]. An increasing number of additional conserved and functional elements within the kinase domain have been identified including a PIKK regulatory domain called PRD [99].

In humans, biallelic mutations in ATM lead to the syndrome ataxia-telangiectasia (AT), whose clinical phenotypes include neurodegeneration, immunodeficiency, radiosensitivity, premature aging and predisposition to cancer [100,101]. Furthermore, mutations that reduce ATR protein levels lead to the Seckel syndrome, a hereditary form of microcephalic dwarfism [102].

Both Tel1/ATM and Mec1/ATR recognize DNA damage, but their DNA damage specificities are distinct [103]. Whereas Tel1/ATM is activated by DSBs, Mec1/ATR responds to a broad spectrum of DNA lesions. Upon DNA damage recognition, these apical kinases trigger a phosphorylation cascade that leads to the activation of the downstream protein kinases Rad53 (human CHK2) and Chk1 (human CHK1) in order to enforce the DNA damage checkpoint response [2]. Although Rad53 is more similar to CHK2 by homology, CHK1 is considered functionally analog to Rad53 in humans, at least during S phase. Transduction of the checkpoint signal from Mec1/ATR to Rad53/CHK2 requires mediator proteins, among which are Rad9 and Mrc1 [73–75, 104–106]. While Rad9 controls the checkpoint throughout the cell cycle, Mrc1 associates with components of the replication fork and promotes DNA damage checkpoint activation specifically during S phase [104,106].

Once activated, Tel1/ATM and Mec1/ATR elicit a signaling cascade through the phosphorylation of numerous downstream substrates, among which histone H2A/H2AX [79–81]. Phosphorylated H2A marks the chromatin surrounding the DNA break and helps the recruitment of DNA repair proteins and chromatin-remodeling complexes [79–81,84].

Tel1/ATM

Tel1/ATM is the apical kinase responsible for sensing and signaling unprocessed DSBs [107,108]. Activation of Tel1/ATM clearly requires the MRX/MRN complex [109–111], but how MRX/MRN exerts this function is still not fully understood. Works in S. cerevisiae, Schizosaccharomyces pombe, Xenopus laevis and human cells have shown that MRX/MRN drives the localization of Tel1/ATM to the site of damage through direct interaction between Tel1/ATM and the C-terminal domain of the Xrs2/NBS1 subunit [112–114]. However, also the Mre11–Rad50 subcomplex contributes to Tel1/ATM activation. In fact, Mre11 and Rad50 individually interact with Tel1/ATM, suggesting an extensive MRX–Tel1 interaction interface [115]. Furthermore, in both S. pombe and mammals Mre11–Rad50 is sufficient to stimulate the activity of Tel1/ATM, as well as its binding to DNA [110,111,116]. In budding yeast, analyses of the individual subunits and of different MRX heterodimeric pairs in the presence of DNA revealed that Tel1 activation requires that the Rad50 protein is proficient in ATP-binding [115,117], suggesting that MRX/MRN activates Tel1/ATM when it is present in the ATP-bound conformation. This hypothesis is supported by the identification of the S. cerevisiae rad50-A78T mutant allele, which specifically abolishes Tel1 activation without impairing MRX functions in DSB repair [118]. Molecular dynamics simulations have shown that while the wild type Mre11–Rad50 subcomplex bound to ATP lingers in a closed conformation, the Mre11–Rad50A78T subcomplex undergoes opening, suggesting that the defective Tel1 activation by Mre11–Rad50A78T results from the destabilization of the ATP-bound conformation [118]. In any case, dsDNA and MRX activate budding yeast Tel1 synergistically [115], suggesting that DNA-induced conformational changes within the MRX complex are responsible for the activation of Tel1 as a kinase.

Cryo-EM structures of S. cerevisiae and Chaetomium thermophilum Tel1 in complex with an ATP analog show that the PRD domain, which is a PI3K-specific feature that was originally identified as an essential element for ATR activation by TopBP1 [99], sits on top of the kinase activation loop and restricts the peptide substrate binding site [119,120]. Thus, it is possible that MRX/MRN binding could affect the PRD domain conformation, possibly releasing inhibition for substrate binding and active site access.

Interestingly, a screen for S. cerevisiae mutants that require Tel1 to survive to genotoxic treatments has shown that Tel1, once it is loaded on dsDNA by MRX, promotes/stabilizes MRX association to the DSB in a positive feedback loop [121]. Tel1 exerts this function independently of its kinase activity, suggesting that it plays a structural role in promoting/stabilizing MRX retention to DSBs.

Mec1/ATR

In contrast with Tel1/ATM that responds primarily to DSBs, Mec1/ATR is activated by a much wider range of genotoxic lesions, including those induced by dNTP depletion, ultraviolet radiation, topoisomerase poisons, DNA polymerase inhibition, alkylating agents and DNA cross-linkers [122,123]. These DNA lesions have been traced to a common DNA structure that ATR can recognize. This structure consists of ssDNA, which also triggers the SOS response in bacteria [124]. Like in bacteria, where the SOS response is elicited by the ssDNA binding protein RecA [124], Mec1/ATR is activated by stretches of ssDNA that are covered by RPA [125]. Such RPA-coated ssDNA tracts are generated during several DNA metabolic processes, such as transcription, DNA replication and telomere maintenance. Furthermore, they are intermediates in many DNA repair mechanisms, such as during DSB resection in HR and after the incision of UV-induced lesions by NER endonucleases.

In particular, inactivation of NER in both yeast and mammals prevents Mec1/ATR activation following UV irradiation [126–129]. Furthermore, in E. coli, UV-induced DNA lesions cause SOS induction only when cells attempt to replicate UV-damaged DNA [130]. These findings suggest that NER function is required to activate an UV-induced checkpoint response outside of the S phase, whereas replication of UV-damaged DNA is sufficient by itself to generate enough ssDNA for checkpoint activation. Consistently with this hypothesis, an electron microscopy analysis of chromosomal replication forks has revealed that S. cerevisiae cells uncouple leading and lagging strand replication at irreparable UV lesions, thus generating long ssDNA regions on one side of the fork [131]. Interestingly, activation of the checkpoint after UV irradiation in non-cycling yeast cells requires the activity of Exo1, which generates extended ssDNA regions by enlarging the gaps generated by NER proteins [132,133]. This checkpoint activation is enhanced by impediments in repair DNA synthesis, suggesting that a defective refilling reaction by DNA polymerases allows Exo1 to generate extended ssDNA gaps that activate a Mec1/ATR checkpoint response.

ssDNA recognition by Mec1/ATR is mediated by Ddc2 (human ATRIP), which forms a constitutive complex with Mec1/ATR and promotes its activation [125,134–136]. However, localization of Mec1–Ddc2 to the sites of DNA damage is not sufficient to convert it into a catalytically active complex. Specific sensors transduce the DDR signal to Mec1 and stimulate its kinase activity. In budding yeast, kinase activation depends on the binding to Mec1 of at least three different activator proteins: the ATP-dependent helicase/nuclease Dna2, the checkpoint protein Ddc1 (human RAD9) and the multi-BRCT domain protein Dpb11 [137–140] (Figure 3). The human Dpb11 ortholog, TopBP1, stimulates Mec1/ATR kinase activity as well [141–143]. In both yeast and mammals, Dpb11/TopBP1 recruitment to DSBs and the consequent stimulation of Mec1/ATR activity requires that Dpb11/TopBP1 interacts with the Ddc1/RAD9 subunit of the 9–1–1 protein complex (Ddc1–Rad17–Mec3 in budding yeast and RAD9–RAD1–HUS1 in humans). This complex is a ring-shaped heterotrimer that is loaded onto ssDNA–dsDNA junctions by the Rad24 (human RAD17)–replication factor C subunits 2–5 (RFC2–5) clamp loader [139,144]. In budding yeast, the interaction between Dpb11 and 9–1–1 requires phosphorylation by Mec1/Tel1 of Ddc1 threonine 602 and this phospho-dependent Dpb11–Ddc1 interaction is conserved in mammals [139,142,145]. In budding yeast, Ddc1 is also able to directly stimulate Mec1 activation, a function that does not seem to be conserved in humans [139,144].

In mammals, full ATR activation requires not only TopBP1 and the 9–1–1 complex (at least to allow TopBP1 loading at DNA ends), but also ETAA1 protein that, unlike TopBP1, is directly recruited to RPA-coated ssDNA through a direct interaction with RPA [146–148].

Little is known about why there are multiple Mec1/ATR activators. They may direct Mec1/ATR towards distinct substrates, recognize different DNA structures in specific phases of the cell cycle and/or amplify the robustness of the DDR. In budding yeast, the employment of the three Mec1 activators differs depending on the phase of the cell cycle [145,149]. Similarly, in humans, TopBP1 seems to activate ATR during replication stresses, whereas ETAA1 promotes ATR activation during S/G2 [150,151].

Interestingly, although these Mec1/ATR activators are structurally unrelated, they all contain an unstructured Mec1-activating domain (MAD) or ATR-activating domain (AAD), which is absolutely required for Mec1/ATR activation [99,139,142,143,149,152]. Although what makes Mec1/ATR more active when it is bound to a MAD/AAD is not known, the mechanism of activation by Mec1/ATR-activating proteins could be similar because all the yeast and vertebrate MADs/AADs identified to date contain hydrophobic amino acids that are essential for Mec1/ATR binding and activation.

Switch from Tel1/ATM to Mec1/ATR signaling

The generation of RPA-coated ssDNA not only promotes Mec1/ATR activation, but also progressively attenuates Tel1/ATM signaling. In fact, it was shown that blunt dsDNA ends, as well as dsDNA ends with short ssDNA overhangs, are preferential substrates for mammalian ATM [153,154]. As the single-stranded tail increases in length, ATM activation is simultaneously attenuated [154], suggesting that ssDNA generation promotes a switch from Tel1/ATM- to Mec1/ATR-dependent checkpoint signaling (Figure 2). A similar mechanism has been proposed for budding yeast Tel1, whose signaling activity is progressively lost when the DSB ends are nucleolytically degraded [155].

Interestingly, the lack of S. cerevisiae Tel1 slightly reduces the efficiency of DSB resection [155]. Furthermore, inactivation of mammalian ATM leads to diminished ATR signaling at DSBs [156–158], suggesting that Tel1/ATM promotes the activation of Mec1/ATR by inducing ssDNA generation. How Tel1/ATM promotes DSB resection is unknown. Interestingly, once loaded at the DSBs by MRX, Tel1 supports MRX persistence at the DSBs in a positive feedback loop [121], suggesting that Tel1 facilitates DSB resection by promoting MRX function.

In contrast with the absence of Tel1, the lack of Mec1 accelerates the generation of ssDNA at the DSB [69], suggesting that Mec1 inhibits DSB resection. This Mec1-mediated inhibition of DSB resection may avoid excessive ssDNA generation and thus ensure a rapid checkpoint switch off when the DSB is repaired. Mec1 exerts this function by acting at least in two manners: (i) it induces Rad53-dependent phosphorylation of Exo1 and this leads to the inhibition of Exo1 activity [89,90] and (ii) it promotes the accumulation of the DSB resection inhibitor Rad9 at DNA DSBs, possibly through γH2A generation [69].

Altogether, these data are consistent with a model whereby the binding of MRX/MRN to DNA ends promotes the recruitment of Tel1/ATM, which facilitates the generation of ssDNA by promoting MRX/MRN association to DNA ends (Figure 2). MRX/MRN, in turn, incises the 5′-terminated strands at Ku-bound DNA ends. Then, MRX/MRN proceeds back toward the DSB end using the Mre11 3′–5′ exonuclease activity, whereas Exo1 or Sgs1–Dna2 nuclease degrade DNA in the 5′–3′ direction. The generation of RPA-coated ssDNA ends, which are no longer recognized by Tel1/ATM, induces the recruitment of Mec1/ATR. Initiation of DSB resection also promotes the association to ssDNA/dsDNA junctions of both the 9–1–1 complex and Dpb11/TopBP1, which not only stimulate Mec1/ATR kinase activity, but also promote the recruitment of Rad9/53BP1. Rad9/53BP1 association to DSBs transduces the checkpoint signal from Mec1/ATR to Rad53/CHK2 and limits the resection activity of Exo1 and Dna2–Sgs1 (Figure 2).

Quantitative mechanisms of Mec1/ATR activation

Given that cells are never entirely free of DNA lesions and that ssDNA is inherently generated during DNA replication and transcription, Mec1/ATR activation should occur only when a threshold of a tolerable amount of ssDNA is exceeded. The existence of signaling thresholds can be observed in the bacterial SOS response, where the relieve of LexA-mediated repression of DNA repair genes occurs concomitantly with RecA–ssDNA generation [159]. Interestingly, in budding yeast it was shown that proteins phosphorylated by Mec1–Ddc2 respond differently to quantitatively different ssDNA signals [160]. In fact, the checkpoint response leading to γH2A generation is already fully active at relatively low (<1.1 kb) levels of ssDNA, but it is unresponsive to increased amounts of ssDNA. In contrast, phosphorylation/activation of the effector kinase Rad53 requires more than 20 kb of ssDNA and it quantitatively responds to the ssDNA length [160,161]. Indeed, H2A is an integral component of chromatin and therefore it can be immediately available for phosphorylation by Mec1/ATR. In contrast, Rad53 phosphorylation/activation requires the recruitment to the sites of DNA damage of several proteins, such as 9–1–1, Dpb11 and Rad9, raising the possibility that multiple ATR activators can provide an additional way to regulate or amplify Mec1/ATR signaling quantitatively. Consistently with this hypothesis, the association of 9–1–1, Dpb11 and Rad9 to a DSB is influenced by the extent of DNA end resection [160]. Furthermore, artificial hyperactivation of the 9–1–1 axis triggers Rad53 hyperactivation even under conditions of reduced Mec1–Ddc2 association [160], suggesting that the amount of 9–1–1 bound at a DSB contributes to Rad53 activation in a quantitative manner. Finally, in budding yeast, Rad53 activation is counteracted by the Slx4 protein, which constitutively interacts with the multi-BRCT domain protein Rtt107. Slx4 dampens Rad53 signaling by binding to Dpb11 and counteracting Dpb11–Rad9 interaction [87,162,163]. Since Rad9 acts as an inhibitor of DSB resection, Slx4 favors the nucleolytic processing of DSBs [164]. In any case, as Rad53 phosphorylation leads to a checkpoint-mediated cell cycle arrest, the finding that its activation quantitatively responds to ssDNA ensures that it occurs only when the amount of DNA damage is so high that DNA repair is not guaranteed [160].

Perspectives

  • The DDR, which includes both DNA repair and checkpoint pathways, is essential for the maintenance of genomic integrity. Defects in the DDR are implicated in several human diseases, including cancer.

  • The initiating events in the DDR entail the recognition of the DNA lesion and the assembly of DDR protein complexes at sites of damage. DNA damage signaling can be driven by protein phosphorylation catalyzed by apical protein kinases, which play key roles in activating the DDR.

  • One future challenge is to understand how the DDR impacts on cellular functions and how the repair and checkpoint pathways are connected. Such knowledge will help to design strategies to control deleterious consequences on human health. Understanding these processes will enable us to design more focused therapies for cancer and other diseases associated with genome instability.

Competing Interests

The Authors declare that there are no competing interests associated with the manuscript.

Funding

This work was supported by Fondazione AIRC under IG 2017 — ID. 19783 project — P.I. Longhese Maria Pia and Progetti di Ricerca di Interesse Nazionale (PRIN) 2017 to M.P.L. C.V.C. was supported by a fellowship from Italian Ministry of University and Research (MIUR) through the grant ‘Dipartimenti di Eccellenza-2017’.

Author Contribution

All authors contributed to, read and approved the final version of the manuscript.

Acknowledgements

We thank Giovanna Lucchini for critical reading of the manuscript.

Abbreviations

     
  • AP

    apurinic/apyrimidinic

  •  
  • AT

    ataxia–telangiectasia

  •  
  • ATM

    ataxia–telangiectasia-mutated

  •  
  • ATR

    ATM- and Rad3-related

  •  
  • BER

    base excision repair

  •  
  • DDR

    DNA damage response

  •  
  • DSB

    double-strand break

  •  
  • dsDNA

    double-stranded DNA

  •  
  • GG

    global genome

  •  
  • HR

    homologous recombination

  •  
  • MMR

    mismatch repair

  •  
  • MRN

    MRE11–RAD50–NBS1

  •  
  • MRX

    Mre11–Rad50–Xrs2

  •  
  • NER

    nucleotide excision repair

  •  
  • NHEJ

    non-homologous end joining

  •  
  • RER

    ribonucleotide excision repair

  •  
  • RPA

    replication protein A

  •  
  • SSB repair

    single-strand break repair

  •  
  • ssDNA

    single-stranded DNA

  •  
  • TC

    transcription-coupled

  •  
  • UV

    ultraviolet

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