Genomic DNA is constantly associated with the various proteins that are involved in DNA folding and transactions. The association between DNA and proteins is reversible, and when prompted, proteins dissociate from or translocate along the DNA strand, leaving the open nucleotide sequence available for replication, transcription, and repair. This process ensures the faithful expression and propagation of genetic information. However, exposure of cells to DNA-damaging agents can cause proteins to become covalently trapped on DNA, generating DNA–protein cross-links (DPCs) [1, 2]. The formation of DPCs was originally demonstrated in bacterial and mammalian cells that had been heavily irradiated with ultraviolet light [3, 4]. It was subsequently shown that DPCs can be produced by various chemical and physical agents, such as aldehydes , metal ions , and ionizing radiation , and by certain types of anticancer agents [8-10].
DPCs are unique among DNA lesions, since they are extremely bulky and are likely to impose steric hindrances upon proteins involved in DNA transactions, and hamper their function. Despite the potential importance of DPCs as genomic damage, they have received less attention than other DNA lesions. Accordingly, much remains to be learned about how cells alleviate the toxic effects of DPCs and about what happens to cells if DPCs are left unrepaired.
The characteristics of DPCs vary considerably with respect to the size, physicochemical properties, biological function, and cross-linking bonds of the trapped proteins. The currently known DPCs can be subdivided into four groups (types 1–4) according to whether and how they are associated with flanking DNA nicks (Fig. 1) [2, 11]. Type 1 DPCs contain proteins that are covalently attached to an undisrupted DNA strand. They are the most common form of DPC found under physiological conditions and are produced by chemical and physical agents such as aldehydes, chromate, platinum compounds, ionizing radiation, and ultraviolet light . Type 2 DPCs, which were identified very recently
In this article we review the current knowledge regarding the formation, repair, and biological effects of type 1 and 2 DPCs (Fig. 1). There already exist extensive reviews and research papers on similar topics for the TOPO-inhibitor-induced type 3 and type 4 DPCs [15-17], and so these will not be dealt with herein.
2. Detection and characterization of DPCs
2.1. Overview of DPC detection
Analysis of the induction and removal of DPCs in the genome is indispensible when studying the repair and biological effects of DPCs. DPCs can be detected either directly or indirectly; while DNA purification is not required for the indirect detection method (Section 2.2), it is required for both direct detection (Section 2.3) and immunodetection (Section 2.4) methods. When required, DNA can be purified by conventional cesium chloride density gradient ultracentrifugation [10, 18, 19] or using the DNAzol-based method [20, 21]. Recently, the rapid and small scale purification methods of DNA were reported and used for immunodetection of DPCs [22, 23]. The methods of DPC detection and their principles are summarized below.
2.2. Indirect detection of DPCs
The indirect methods of detecting DPCs include the alkaline elution, nitrocellulose filter-binding, sodium dodecyl sulfate (SDS)/potassium ion (K+) precipitation, and single-cell gel-electrophoresis methods. The alkaline elution method is based on the different elutabilities of DNA without and with cross-linked proteins from a filter under alkaline conditions [24, 25]. In brief, cells are filtered onto a polyvinylchloride filter and lysed with sarkosyl; the DNA that is retained on the filter is eluted at pH 12.1. The adsorption of cross-linked proteins to the filter reduces the elutability of unwound single-stranded DNA, thereby changing its elution kinetics. The nitrocellulose filter-binding method depends upon the different abilities of DNA without and with cross-linked proteins to bind to a nitrocellulose filter [26, 27]. In this method, cells are lysed with sarkosyl and passed through the filter, which retains proteins and DNA with cross-linked proteins, but not free DNA. The amount of DNA that is retained on the filter via cross-linked proteins is then assayed for DPCs. The SDS/ K+ precipitation method is based on SDS binding tightly to proteins to form insoluble precipitates with K+ [6, 28]. Cells are lysed with SDS, and SDS-bound proteins and DNA with cross-linked proteins, but not free DNA, are selectively precipitated by KCl. The amount of DNA precipitated due to cross-linked proteins is then assayed for DPCs. The single-cell gel-electrophoresis method (the comet assay) detects retarded DNA migration due to a certain type of DPC [29, 30]. Pretreatment of lysed cells with proteinase K enables the distinction between DNA with and without DPCs in these methods.
While a major advantage of these aforementioned indirect methods is that they enable the detection of DPCs without purifying DNA, there is no linear relationship between the amounts of DNA and cross-linked proteins. This makes it difficult to quantitatively interpret the data derived from these indirect measurements of DPCs.
2.3. Direct detection of DPCs
Two techniques have been developed that allow direct and quantitative analysis of DPCs: the 125I-postlabeling and fluorescein isothiocyanate (FITC)-labeling methods. The 125I-postlabeling method is based on the specific incorporation of 125I into tyrosine residues that are associated with purified DNA . A recently reported FITC-labeling method has been shown to provide a more straightforward analysis. Cross-linked proteins in purified DNA are specifically labeled with FITC and directly assayed for the resulting fluorescence [18, 19, 32]. A key advantage of the FITC-labeling method is that the amount of DPCs is proportional to the fluorescence intensity of the labeling.
2.4. Immunodetection of DPCs
Most DPC-inducing agents are fairly nonspecific and covalently trap various proteins. However, DPCs can be detected directly by Western blotting if the identity of the cross-linked protein is known. DNA methyltransferase (DNMT), which is associated with type 1 DPC [10, 22], PARP-1, which is associated with type 2 DPC [12, 13], and TOPOs I and II, which are associated with type 3 and type 4 DPC, respectively , can be detected by Western blotting when they are covalently trapped in DNA by inhibitors.
2.5. Proteomic analysis of cross-linked proteins
Considerable insight into the biological effects and repair mechanisms of DPCs can be obtained by identifying the cross-linked proteins in DNA. Comprehensive proteomic analyses of the cross-linked proteins induced by ionizing radiation , formaldehyde , mechlorethamine (one of the antitumor nitrogen mustards) [34, 35], and butadiene diepoxide (a carcinogenic metabolite of 1,3-butadiene) [36, 37] have been performed. The identified cross-linked proteins include those participating in transcriptional regulation, translation, RNA processing, DNA damage response, DNA repair, cell cycle, homeostasis, cell signaling, and cell architecture. These proteomic approaches may have potential applications in the analysis of DNA-damage interactomes .
3. Formation of DPCs
DPCs are produced by various chemical and physical agents or during DNA transactions. Here we summarize the formation of DPCs by selected agents including aldehydes, bifunctional alkylating agents, and ionizing radiation. We also refer to DPC formation by abortive DNA metabolism and repair.
3.1. Formation of DPCs by DNA-damaging agents
Aldehydes are well-known inducers of type 1 DPCs. Humans are exposed to various aldehydes through anthropogenic and food sources. Aldehydes are also generated by lipid peroxidation and metabolism in cells . Aldehydes that have escaped from the detoxification systems of cells react with DNA, proteins, and other biomolecules and hamper cellular functions. The reactions between aldehydes and DNA result in the formation of DPCs [1, 5] and base adducts . Other DNA lesions such as DNA intrastrand cross-links, DNA interstrand cross-links (ICLs), SSBs, and DSBs may be formed concurrently to varying extents .
In light of the reactivity of aldehydes, the side chains of lysine, cysteine, and histidine residues in proteins react with aldehydes to form adducts (Fig. 2). The aromatic amines of DNA bases are weak nucleophiles and are less reactive to aldehydes. The resulting protein adducts react further with the amino group of DNA bases to form various types of cross-linking bond that have different chemical stabilities. The cytotoxicities of formaldehyde, chloroacetaldehyde, acrolein, crotonaldehyde,
Finally, it is also worth noting that the importance of DNA damage induced by endogenous aldehydes has been again acknowledged in recent studies. Studies involving mouse models have revealed that DNA damage induced by endogenous aldehydes is associated with the symptoms of Fanconi anemia (FA), which is a complex heterogenic disorder of genomic instability, bone marrow failure, and cancer predisposition [44, 45]. In a study involving chicken DT40 cells, it was found that the catabolism of formaldehyde is essential for cells deficient in the FA DNA-repair pathway . The identity of the aldehyde-induced DNA lesion that is responsible for FA remains elusive.
3.1.2. Bifunctional alkylating agents
Bifunctional alkylating agents have been generating considerable interest as both health hazards and anticancer drugs. Their biological activity relies on their capacity to cross-link biomolecules, resulting in inactivation of their function. With DNA, bifunctional alkylating agents react with DNA to form monoadducts. The remaining reactive site of these reagents can further react with either DNA to form a DNA–DNA cross-link or a protein to form a DPC .
Glyceraldehyde 3-phosphate dehydrogenase and histones are cross-linked to DNA by butadiene diepoxide
The nitrogen mustards containing
Mechlorethamine belongs to the member of the nitrogen mustards. The formation of DPCs together with ICLs upon the treatment of cells and nuclei with mechlorethamine and other cross-linking agents (nitrosoureas) was initially demonstrated using the alkaline elution method . Recent proteomic analyses have revealed the formation of DPCs in mechlorethamine-treated nuclear extracts and cells, demonstrating the involvement of functionally different proteins in DPCs [34, 35].
Mitomycin C is another class of bifunctional alkylating agent and is also used for cancer therapy. FK973, FK317, and FR900482 are substituted dihydrobenzoxazine derivatives and undergo reductive activation to form the reactive mitosene structures that are similar to that of mitomycin C. Alkaline elution analyses have shown that together with mitomycin C, FK973 forms concentration- and time-dependent ICLs and DPCs in cells, but not SSBs . In addition, chromatin immunoprecipitation analyses have revealed that FR900482 and FK317 cross-link minor-groove binding proteins such as HMGA1, HMGB1, and HMGB2, but not major-groove binding proteins such as NF-κB or Elf-1, to the promoter regions of the IL-2 and IL-2Rα genes to form DPCs
3.1.3. Ionizing radiation
Ionizing radiation causes damage to DNA via both direct and indirect mechanisms. In the direct mechanism, the radiation energy is deposited directly in DNA and produces DNA cation radicals, which are unstable and undergo decomposition. In the indirect mechanism, the radiation energy is deposited to water (i.e., the bulk medium of cells) and produces reactive oxygen species such as hydroxyl radicals, which in turn attack and damage DNA. Various types of radiation-induced DNA lesion have been identified: base damage, SSBs, DSBs, and DPCs. The most critical damage underlying the cell-killing effects of ionizing radiation is attributed to DSBs. The efficiency of DSB formation by ionizing radiation is decreased under hypoxic conditions relative to normoxic conditions, whereas that of DPC formation is increased under hypoxic conditions [1, 7]. Although the contribution of DPCs to the lethal events in irradiated cells remains to be clarified, the aforementioned opposing effects of oxygen on DSB and DPC formation point to the potential importance of DPCs for hypoxic cells, and in particular those present in tumors.
The induction of DPCs and their removal from the genome following irradiation of normoxic and hypoxic mouse tumors with carbon-ion beams were recently analyzed using the FITC-labeling method . The yield of DPCs was greater by 4-fold in hypoxic tumors than in normoxic tumors. Simultaneously, the yield of DSBs in hypoxic tumors was decreased to 1/2.4 relative to that in normoxic tumors. Interestingly, the carbon-ion beams produced two types of DPC that differed according to their rate of removal from the genome. The half-life of the rapidly removed component of DPCs was less than a few hours
It is possible that the rapidly removed DPCs are chemically unstable and reversed spontaneously by hydrolysis as is the case for aldehyde-induced DPCs (see Section 3.1.1 and Fig. 2). Alternatively, they may be peptide-containing DPCs, which are relatively small and are efficiently removed from DNA by nucleotide excision repair (NER) as described in Section 4.4.2. Slowly removed DPCs are virtually irreversible DPCs and are resistant to excision repair as evidenced by their long half-live
3.2. Formation of DPCs by abortive DNA metabolism and repair
3.2.1. DPC formation by inhibition of DNA-metabolizing and repair enzymes
Some classes of enzymes form transient covalent complexes with their substrates during catalysis. Those involved in DNA metabolism and repair are no exception to this, and considerable numbers of enzymes have been found that form a transient covalent complex with DNA as a reaction intermediate.
The methylation of cytosine in 5’-CG-3’ sequences is an important carrier of epigenetic information in higher organisms; this methylation is performed by DNMTs including DNMT1 (maintenance methyltransferase), DNMT3a, and DNMT3b (both
Bifunctional DNA glycosylases and DNA repair proteins such as PARP-1 and Ku have an associated AP lyase activity and react with AP sites in DNA, forming covalent Schiff base intermediates [66-68]. The structure of these covalent Schiff base intermediates is similar to that of type 2 DPCs (Fig. 1). DNA polymerases that have an associated 5’-terminal 2-deoxyribose-5-phosphate (dRP) lyase activity also form covalent Schiff base intermediates [69, 70]. These intermediates mimic type 2 DPCs (Fig. 1), but the protein is tethered to the 5’ end of a SSB via dRP. With the exception of PARP-1, the covalent Schiff base intermediates of the aforementioned glycosylases, repair proteins, and polymerases cannot be isolated, but they can be stabilized by NaBH4-reduction and isolated as DPCs . Interestingly, the formation of stable DPCs containing PARP-1 (type 2)
Tyrosyl-DNA phosphodiesterase (Tdp1) is involved in the repair of TOPO I–DNA covalent complexes (see below), and its catalytic cycle involves a covalent reaction intermediate in which a histidine residue is connected to a DNA 3’-phosphate through a phosphoamide linkage . In the strand-breakage reaction catalyzed by TOPO I and TOPO II, a nucleophilic attack of a catalytic tyrosyl residue of the TOPO upon a DNA phosphodiester bond results in transient covalent attachment of the tyrosine to the DNA phosphate either at the 3’-end (TOPO I) or the 5’ end (TOPO II) of the broken DNA (Fig. 1) [15, 16]. TOPO inhibitors (poisons) such as camptothecin (a TOPO I inhibitor) and etoposide (a TOPO II inhibitor) freeze the covalent reaction intermediate and abort the subsequent rejoining of DNA ends, leaving TOPO cleaved complexes that contain strand breaks and protein covalently bound to DNA (type 3 and type 4 DPCs; Fig. 1) [15, 16]. Many chemotherapeutic drugs targeting the covalent TOPO reaction intermediates have been developed since type 3 and type 4 DPCs are complex DNA lesions containing DPC(s) and strand break(s) and would be effective at killing tumor cells.
3.2.2. Suicidal cross-linking DNA damage
Several DNA lesions have been shown to act as suicidal substrates by stably cross-linking base excision repair (BER) enzymes, although the cross-linking reactions have only been demonstrated
2-Deoxyribonolactone (dL) is an oxidized form of an AP site and is produced by many DNA-damaging agents. dL in DNA undergoes β-elimination to form α,β-unsaturated lactone at the 3’-terminus. Alternatively, dL can be incised by an AP endonuclease (e.g., APE1) to form a dL phosphate at the 5’-terminus. The lactone in these dL analogs can react with nucleophiles such as lysine to form a stable amide bond. It has been shown that of eight bifunctional DNA glycosylases tested, Nth/Endo III cross-links to dL in DNA, while Fpg and hNEIL1 cross-link to the β-elimination product of dL (Fig. 3A) [72, 73]. The dL phosphate at the 5’-terminus generated by the incision of APE1 also cross-linked to DNA polymerase (Pol) β . Other forms of oxidized 2-deoxyribose (dioxobutane and the C4-oxidized AP site) at the 5’-terminus of DNA produce transient covalent complexes with Pol β and Pol λ, but the complex containing the oxidized sugar and enzyme is subsequently released from DNA, resulting in inactivated free polymerases .
Oxanine (Oxa) is produced by nitrosative damage of guanine  and has a reactive lactone-like structure. It was shown that of seven DNA glycosylase tested, Fpg, Nei/Endo VIII, and hOGG1 (bifunctional glycosylases) and AlkA (monofunctional glycosylase) cross-link to Oxa in DNA to form type 1 DPCs (Fig. 3B) . The glycosylases trapped by Oxa are notably different from those trapped by dL (Nth), suggesting distinct interactions with DNA damage in the active site. Histones also react with Oxa to produce DPCs, but at much lower rates, indicating the importance of specific interactions with proteins in DPC formation. With dL and Oxa, it seems that the catalytic amino acids in the active site (lysine or proline) of the enzymes attack the carbonyl carbon of dL or Oxa, resulting in stable amide bond formation (Fig. 3AB).
5-Hydroxy-5-methylhydantoin is produced by oxidation of thymine, and the carbanucleoside of 5-hydroxy-5-methylhydantoin (cHyd) has been shown to covalently trap Fpg, Nei/Endo VIII, and hNEIL1, serving as a suicidal substrate (Fig. 3C) . The crystal structure of the cHyd-DNA and Fpg covalent complex directly revealed the cross-linking between the N-terminal proline and the C5 of cHyd.
The cross-linking efficiencies of dL, Oxa, and cHyd for glycosylases or polymerases are relatively low. Thus, the biological significance of these lesions as suicidal substrates for repair enzymes remains to be assessed
4. Repair of DPCs
4.1. DPC repair in bacterial cells
The genes involved in the repair of DPCs have been elucidated by analyzing the sensitivity of a panel of repair-deficient
Since large DPCs are not removed from DNA by NER, they stall the replication fork. As mentioned above, the genetic data show the essential role played by HR plus RR in the reactivation of a stalled replication fork by DPCs. The HR of
The precise molecular mechanism underlying reactivation of the DPC-induced stalling of the replication fork by HR remains to be established. Inactivation of some replication proteins (DnaB, Rep helicases, DNA Pol III) by mutations results in fork breakage and DSBs in a
4.2. DPC repair in yeast
Yeasts such as
Reexamination of the sensitivity of individual strains to chronic low-dose exposure to formaldehyde has indicated that cell survival is mainly conferred by proteins of the HR pathway and those related to that process [
Interestingly, the requirement of repair gene to mitigate the cytotoxic effect of formaldehyde following acute high dose exposure (60 mM, 15 min) differed significantly from that following chronic low-dose exposure to formaldehyde (1.0–1.5 mM for 48 h) . The NER-deletion strains (
It has been demonstrated very recently that the metalloprotease Wss1 in
4.3. DPC repair in chicken DT40 cells
The chicken B lymphocyte cell line DT40 has a high rate of gene targeting and has been used as a model system for reverse genetics studies in higher eukaryotes . The genes involved in the repair of DPCs have been elucidated by assessing the formaldehyde sensitivity of DT40 cells with targeted mutations in various DNA repair genes . In total, 22 mutants involved in different DNA repair pathways were studied: FA (
Interestingly, the DT40
4.4. DPC repair in mammalian cells
4.4.1. Sensitivity to DPC-inducing agents
Chinese hamster ovary (CHO) cells deficient in HR (
The sensitivity of FA-pathway-deficient mammalian cells to formaldehyde has also been examined. One study involving mouse and CHO cells found that cell survival was dependent on
The removal of DPCs induced by formaldehyde and other aldehydes has been analyzed using NER-proficient and NER-deficient (
4.4.2. Repair capacity of mammalian NER for DPCs
The capacity of mammalian NER to repair DPCs has been studied
As with conventional bulky lesions, the 5’-incision sites were around the 21st phosphodiester bond 5’ to DPCs, and 3’-incision site was at the 6th phosphodiester bond 3’ to DPCs, which indicates that the incision sites are independent of the size of cross-linked proteins . The CFEs from NER-deficient cells exhibited no incision activity for DPCs. Despite significant differences in protein components, bacterial and mammalian NER shares an activity optimum for a cross-linked protein size of around 1.6 kDa, which is several times larger than the sizes of conventional bulky lesions. It would be interesting to know whether this is simply due to a mechanistic reason or if it has some evolutional significance.
An alternative model of DPC repair by mammalian NER has been proposed. In this model cross-linked proteins are initially degraded to short peptides by the proteasome, and due to the robust activity of mammalian NER for DPCs containing short peptides
4.4.3. DPC tolerance by HR
Since NER is virtually unable to repair DPCs in mammalian cells, the replication fork will run into the DPC site and become stalled. Given the high sensitivity of HR-deficient CHO mutants to formaldehyde and azadC, HR is pivotal with respect to activation of the DPC-stalled replication fork. Indeed, the formation of RAD51 nuclear foci, which is reminiscent of HR, was observed following treatment with formaldehyde and azadC . Accumulation of DSBs was observed in HR-deficient CHO cells, but not in HR-proficient CHO cells after treatment with formaldehyde or azadC, suggesting that HR for DPCs is initiated by fork breakage to generate one-sided DSBs .
When CHO cells are treated with replication inhibitors such as hydroxyurea and aphidicolin, accumulation of DSBs due to fork breakage is observed even in HR-proficient cells [111, 112], suggesting mechanistic differences in the formation of DSBs by DPCs and replication inhibitors. The MUS81-EME1 and MUS81-EME2 structure specific endonucleases are implicated in fork breakage in mammalian cells [113-115]. However, whether they are also involved in fork breakage at DPCs remains to be elucidated.
The data for DT40 cells suggest that TLS is a crucial component of DPC tolerance , but the sensitivity of mammalian TLS mutants to DPC-inducing agents has not been tested. Mouse embryonic stem cells deficient in two major DNA glycosylases (Nth1 and Ogg1) did not exhibit significant sensitivity to formaldehyde and azadC (Ide et al., unpublished data). Regarding the DNA damage response, it was shown that DPCs activate both ATM (ataxia-telangiectasia mutated) and ATR (ATM and Rad3-related) pathways in the late and early stages of damage response, respectively, in human cells .
Studies of the tolerance/repair of DPCs have been hampered by the facts that DPC-inducing agents such as aldehydes, bifunctional alkylating agents, and platinum anticancer drugs concurrently produce ICLs, which are also potent lethal lesions, and that tolerance/repair mechanisms for DPCs and ICLs are partly overlapping at least with respect to the requirement of HR and some structure-specific endonucleases. In the replication-dependent ICL repair mechanism, the FA core complex recognizes and binds an ICL that stalled the replication fork. Then, the FA core complex monoubiquitinates FANCD2 and FANCI. The ubiquitinated FANCD2-I heterodimer localizes to the ICL and recruits structure-specific endonucleases (XPF-ERCC1, MUS81-EME1, FAN1, SLX4) that incise the DNA on either side of the lesion to create a DSB. The complementary strand containing the unhooked cross-link is replicated by a TLS polymerase, and downstream FA proteins assist in coordination of HR to repair the DSB . ICLs are also repaired by the replication-independent mechanism involving structure-specific endonucleases and TLS polymerases . Similarly, DPCs can be repaired in a replication-independent manner if NER coupled with DPC-specific proteases (functional Wss1 homologs) operates in mammalian cells (see Section 4.2). The replication-independent repair of DPCs will be important for the survival of nonproliferating cells such as neurons since it ensures faithful gene expression and maintains cellular homeostasis.
5. Biological effects of DPCs
Some types of DPC generated by chemical and physical agents are stable and are not spontaneously reversed (Section 3). Furthermore, only small DPCs are actively removed from the genome by NER in bacterial cells (Section 4). Thus, a significant portion of DPCs persist in the genome and can affect various aspects of DNA transactions such as replication, transcription, repair, and recombination. This section focuses on the cytotoxic effects of DPCs through DNA replication and transcription.
5.2. Effects on DNA replication
5.2.1. The replisome
DNA is replicated by the replisome, which comprises the replicative DNA helicase, DNA polymerase, and other factors [118, 119]. The replicative helicases unwind the parental double-stranded DNA into two single strands, and DNA polymerases synthesize leading and lagging strands in continuous and discontinuous modes, respectively, using the separated strands as templates. The mechanism is well conserved from phages and bacteria through to higher organisms [118, 119]. The replisome proceeds through the barrier of DNA-associated proteins such as nucleosomes and site-specific DNA-binding proteins. The replicative helicases disrupt nucleosomes in eukaryotes, probably with the aid of histone modifications and chaperones . They can also unwind DNA that is associated with DNA-binding proteins with variable efficiencies [120, 121], and dislodge proteins from double-stranded DNA . Thus, the replisome has an intrinsic capacity to proceed through the protein barrier as long as it is reversible. However, many DNA-damaging agents generate DPCs and immobilize proteins in DNA.
5.2.2. Host-cell reactivation assays
The effects of DPCs on replication were studied
5.2.3. Effects on DNA polymerases
The effects of DPCs on DNA polymerases have been studied
5.2.4. Effects on replicative helicases
The impediment of the replisome by DPCs is more closely associated with replicative helicases that unwind DNA at the front of the replication fork than with replicative polymerases. Replicative helicases such as the phage T7 gene 4 protein (T7gp4), simian virus 40 large T antigen (Tag), and
The effects of DPCs on the DNA-unwinding reaction of replicative helicases have been elucidated
In addition, the results obtained for helicase suggest the distinct fates of replisomes upon encountering conventional bulky damage and large DPCs. Conventional bulky damage both in the translocating and nontranslocating strands are cleared by helicases and arrest DNA polymerase (Fig. 7A). This can further lead to functional uncoupling of polymerases and helicases as well as that of leading and lagging polymerases. In eukaryotes, the functional uncoupling of polymerases and helicases activates a checkpoint kinase ATR (ATM and Rad3-related), which directs the DNA damage response . DPCs in the translocating strand block the helicase, immediately halting leading- and lagging-strand synthesis (Fig. 7B). This will preclude functional uncoupling of polymerases and helicases and of leading and lagging polymerases. In contrast, DPCs in the nontranslocating strand do not block the helicase, and act like conventional bulky damage. Accordingly, the mechanism underlying stalled fork-processing and the concurrent events of damage signaling may differ significantly for DPCs in the translocating and nontranslocating strands.
Stalled DnaB, T7gp4, and Mcm467 helicases exhibit limited stability and dissociate from DNA with a half-life of 15–36 min
In yeast, replisomes stalled by tight (but reversible) DNA–protein complexes are stable
5.3. Effects on transcription
5.3.1. RNA polymerase
Viral, prokaryotic, and eukaryotic RNA polymerases (RNAPs) have an ability to transcribe through nucleoproteins and site-specific DNA binding proteins, although the read-through efficiencies vary depending on the roadblocking proteins . ATP-dependent chromatin remodeling complexes, histone chaperones, and covalent histone modifications promote the transcription through nucleosomes . It has also been shown that the trailing RNAP stimulates forward translocation of the stalled leading RNAP through reversibly bound proteins [143, 144], as well as through naturally occurring pausing sites [145, 146].
RNAPs open the downstream DNA duplex at the DNA entry site to generate a transcription bubble, in which the transcribed strand (TS) is delivered deep into the active site and used for nascent RNA synthesis, while the nontranscribed strand (NTS) is relatively exposed to the surface of RNAP [147, 148]. Resolution of the crystal structure of yeast RNAP II revealed that conventional bulky lesions such as a cyclobutane pyrimidine dimer, a cisplatin intrastrand cross-link, and a monofunctional platinum adduct in the TS are delivered to the active site or its proximal position and then arrest transcription [149-151]. Conversely, DNA lesions in the NTS impose much less serious problems for transcription than do those in the TS .
5.3.2. Reporter assays
Luciferase-based reporter assays are widely used as a tool to study gene expression at the transcriptional level. To assess the effects of DPCs on transcription, histone H1 was cross-linked by formaldehyde to a pGL4.50 plasmid harboring the luciferase gene (Fig. 8A). The pGL4.50 containing histone H1-DPCs was transfected into HeLa cells and the bioluminescence resulting from the expressed luciferase was measured (Ide et al., unpublished data). The luciferase activity was found to decrease with increasing amounts of cross-linked histone H1 protein, indicating that transcription of the luciferase gene by RNAPII was inhibited by DPCs
5.3.3. Effects on T7 RNAP
T7 RNAP is a single subunit RNAP and is structurally unrelated to bacterial and eukaryotic multisubunit RNAPs, but all share many functional characteristics in the initiation and elongation phases of transcription .
The effects of DPCs on transcription have been analyzed
Stalling of leading T7 RNAP by TS-DPCs caused congestion of the trailing T7 RNAPs. Interestingly, sequence analysis of runoff transcripts has shown that stalled leading and trailing T7 RNAPs become highly error prone and generate untargeted mutations in the upstream intact template regions (Fig. 9); 40–75% of runoff transcripts contained mutations in the region . In contrast, no mutations were induced in runoff transcripts when NTS-DPCs were used. This contrasts with the transcriptional mutations induced by conventional DNA lesions, which are delivered to the active site or its proximal position in RNAPs and cause direct misincorporation.
Another interesting observation is that the trailing RNAP stimulates forward translocation of the stalled leading RNAP, promoting the translesion bypass of DPCs . The cooperation of T7 RNAPs enhances transcription through DPCs by a factor of 5.2–17. It has been proposed that bacterial and eukaryotic RNAPs cooperate during elongation so that the trailing RNAP assists in the transcription of the leading RNAP through reversibly bound proteins and pausing sites, by reducing the backtracking of the stalled/paused leading RNAPs [143-146]. Accordingly, similar cooperating mechanism may be working for transcription through DPCs.
How bacterial and eukaryotic multisubunit RNAPs respond to DPCs
DPCs are superbulky DNA lesions that affect replication, transcription, and repair via mechanisms that differ from those involving conventional bulky lesions.
The findings from
In DNA repair, NER is the major mechanism for the repair of conventional bulky lesions. NER exhibits a robust activity for DNA–peptide cross-links, but a poor to negligible activity for DPCs. The initial recognition of DPCs by NER factors appears to be critical, and is compromised due to the steric hindrance of DPCs. However, the proteolytic degradation of DPCs by proteases may enable NER and TLS to participate in DPC repair. HR plays a principal role in the repair/tolerance of DPCs, but the molecular mechanism by which the DPC-stalled replication fork is reactivated through HR remains to be established.
Studies of the cytotoxic effects and repair of DPCs have been hampered by the facts that DPC-inducing agents concurrently produce other lethal lesions, such as ICLs (aldehydes and bifunctional cross-linking agents) and DSBs (ionizing radiation), and that repair pathways for DPCs, ICLs, and DSBs are partially overlapping. These also pose challenges for studies of the mutagenic and carcinogenic effects of DPCs, which were not addressed in this review. Future research will overcome these limitations and clarify the importance of DPCs in DNA damage together with the underlying molecular mechanism of the repair/tolerance of DPCs.
AGT: O6-alkylguanine-DNA alkyltransferase; AP: apurinic/apyrimidinic; azadC: 5-aza-2’-deoxycytidine; azarC: 5-azacytidine; BER, base excision repair; CHO. Chinese hamster ovary cells; cHyd: carbanucleoside of 5-hydroxy-5-methylhydantoin; dL: 2-deoxyribonolactone; DNMT: DNA methyltransferase; DPC: DNA-protein cross-link; dRP: 2-deoxyribose-5-phosphate; DSB: DNA double-strand break; FA: Fanconi anemia; FITC: fluorescein isothiocyanate; HR: homologous recombination; ICL: interstrand cross-link; Mcm: minichromosome maintenance; NER: nucleotide excision repair; NTS: nontranscribed strand; Oxa: oxanine; PARP-1: poly(ADP-ribose) polymerase-1; PRR: post-replication repair; RNAP: RNA polymerase; RR: replication restart; SDS: sodium dodecyl sulfate; SSB: DNA single-strand break; T7gp4: phage T7 gene 4 protein; Tag: simian virus 40 large T antigen; Tdp1: tyrosyl-DNA phosphodiesterase; TLS: translesion synthesis; TOPO: topoisomerase; TS: transcribed strand; XRCC: X-ray repair cross-complementing protein.
This work was partly supported by KAKENHI from the Japan Society for Promotion of Science and the Ministry of Education, Culture, Sports, Science and Technology in Japan (grant numbers: 22131010 and 26550030 to H.I., and 26340023 to T.N.).