ADP-ribosylation is a post-translational modification (PTM) process that is intrinsically associated with basal metabolic signaling pathways and has been recently identified as an essential regulator of DNA repair and cancer biology (Cohen and Chang, 2018; Crawford et al., 2018). An ADP-ribosyl reaction occurs whenever single or multiple ADP-ribose (ADPr) units present on the redox cofactor β-nicotinamide adenine dinucleotide (β-NAD⁺) are transferred to a substrate protein and when nicotinamide (Nam) is released (Cohen and Chang, 2018). The previously identified modifications occurred on receptor residues (Asp, Glu, Ser, Tyr, Arg, and Cys) linked by O-, N-, and S-glycoside bonds (Cohen and Chang, 2018). With advancements in the detection technology, ADP-ribosylation is no longer considered only as a protein modification, and these modifications have also been reported to occur onto phosphorylated nucleic acids such as the ends of DNA and RNA (Dölle and Ziegler, 2017; Munnur et al., 2019; Groslambert et al., 2021; Weixler et al., 2021).

In mammals, this biochemical reaction is mainly catalyzed by three families of enzymes: 1) clostridium toxin-like ADP-ribosyltransferase (ARTs) (ARTCs) catalyze extracellular ADP-ribosylation, 2) diphtheria toxin-like ARTs (ARTDs) catalyze intracellular ADP-ribosylation, and 3) sirtuins (namely, SIRT4, 6, and 7) catalyze ADP-ribosylation in different intracellular compartments. In this review, we focused on the 17ARTD family members in humans, following a recent consensus that “PARP” should be used as a separate term to describe various ARTD family members (Table 1) (Lüscher et al., 2021).

Poly (ADP-ribosyl) polymerase (PARP) family share a highly conserved ART folding region, wherein a binding pocket of NAD⁺ is situated, which contains a conserved His-Tyr-Glu (H-Y-E) triplet, also known as an ART signature sequence (Table 1, Figure 1) (Steffen et al., 2013). The sequence differences affect the ability of PARP molecules to transfer ADPr. The first two amino acids (histidine and tyrosine) are critical for NAD⁺ binding, whereas the glutamate residue is essential for elongation of the poly-ADP-ribose (PAR) chain. Glutamate in the catalytic domain of mono-ARTs is replaced by leucine, isoleucine, or tyrosine and restricted to the transfer of one ADPr unit. On the basis of the catalytic efficacy of enzymes, the PARP family can be divided into three groups: poly-ARTs, mono-ARTs, and inactive members. Analysis of self ADP-ribosylation indicates that only PARP1, 2, and Tankyrase (TNKS) 1, 2 can add multiple ADPr units, whereas the remaining 11 PARPs conjugate a single ADPr to amino acid residues (Vyas et al., 2014). Although PARP3, 4 contain H-Y-E patterns, they function as mono-ARTs (Table 1). PARP9, 13 do not have catalytic activity (Vyas et al., 2013). Furthermore, when PARP9 is linked to histone E3 ubiquitin ligase 3L (DT3XL), it can catalyze the action of mono-ARTs (Yang et al., 2017). In particular, the substitutions of His residues in the ART sequence interfere with the binding of NAD⁺, and such substitutions are observed in the catalytically inactive PARP family member, namely, PARP13 (Vyas et al., 2013).

In addition to differences in the catalytic domains, PARP family members also differ in their respective regulation domains. The main functional domains of mono-ARTs are WGR (Trp-Gly-Arg), macrodomains (Macro), WWE (Try-Try-Glu), CCCH Zn Finger, RNA recognition motif (RRM), and ubiquitin-interacting groups (UIM) (Figure 1).

One of the most well-known regions of PARPs is the WGR (Trp-Gly-Arg) domain, which is a conserved central motif essential for DNA-dependent activation (Langelier et al., 2012). Macrodomains, a family of protein domains, play a crucial role in the recognition and combination of ADPr groups (Perina et al., 2014). Multiple macrodomains are present in PARP9, 14, and 15. WWE (Try-Try-Glu) is a domain that has been named after its three most conserved amino acids, and sequence analyses have indicated its association with ADP-ribosylation. PARP7, 12, 13, and 14 have the WWE domain and E3 ubiquitin-protein ligase 1 in common (Barkauskaite et al., 2015). Previously, the WWE domain of RNF146 (ubiquitin E3 ligase) was reported to interact with the PAR chain (Zhang et al., 2011). In addition, the WWE structure of PARP11 binds to the PAR chain; although this interaction may differ from that of RNF146, it may combine with the terminal units of ADPr in the PAR chain. By contrast, PARP14 does not interact with PAR, and evidence indicates that the WWE domain mediates protein–protein interactions independent of protein modifications by PAR (Wang et al., 2012; He et al., 2012).

PARP7, 12, and 13 contain zinc finger domains of the CCCH-type, which are known as RBP binding to RNA (Guo et al., 2004). Zn CCCH is characterized by its ability to bind to both host and viral RNA (Todorova et al., 2014; Guo et al., 2007). Multiple RRMs within PARP10, 14 are involved in RNA binding with high affinity and sequence specificity. In addition, RRMs can interact with ADPr. For example, the RRM domain of the RNA-binding protein, NONO, can bind to PARP1 produced during DNA damage response (Gagné et al., 2012). PARP10 contains two ubiquitin-interacting motifs (UIMs), which can bind to the polyubiquitin chain via K63 and can promote interactions between PARP10 and proliferating cell nuclear antigen (PCNA) (Verheugd et al., 2013; Nicolae et al., 2014).

Some mono-ARTs can be classified into multiple groups, whereas other mono-ARTs cannot be classified because they contain domains that are either unique (PARP4, 16) or uncharacterized (PARP6, 8) (Figure 1). The specific functions and significance of these molecules remain further explored, and the development of detection and research tools will hopefully help in discovering other poorly studied mono-ARTs (Steffen et al., 2013).

ADP-ribosylation is a reversible modification controlled by ARTs (i.e., writers) and removed by the members of two protein families, namely, macrodomains (MacroD) and (ADP-ribosyl) hydrolases (ARHs, i.e., eraser) (Figure 2) (Rosenthal et al., 2013; Cohen and Chang, 2018; Rack et al., 2020).

MacroD1, MacroD2, and terminal ADP-ribose protein glycohydrolase 1 (TARG1) are three macrodomain-containing enzymes capable of reversing the MARylation of proteins and RNA (Munnur et al., 2019; Rosenthal et al., 2013). MacroD1, 2, and TARG1 preferentially remove the ADPr group from acidic residues and hydrolyze a sirtuin by-product, O-acetyl-ADP-ribose (Rosenthal et al., 2013; Chen et al., 2011; Jankevicius et al., 2013). MacroD1 is most prevalent in the mitochondria of skeletal muscles cells. MacroD2 is localized to the nucleolus and cytoplasm and is found only in neuroblastoma cells, whereas the more ubiquitously expressed TARG1 is present in the nucleoplasm, nucleolus, and stress granules (Žaja et al., 2020).

Among ARHs, ARH1 and inactive ARH2 are localized in the cytoplasm, whereas ARH3 is localized in the nucleus, cytosol, and mitochondria (Moss et al., 1992; Niere et al., 2008; Beijer et al., 2021). ARH3 is the main hydrolases of serine-MARylation and consequently plays a critical role in DNA damage response (Rack et al., 2020). In addition, ARH3 can remove the terminal ADP-ribose moiety from the protein substrate, a necessary step for the process of poly-ADP-ribosylation reversal (Rack et al., 2021). On the other hand, poly (ADP-ribose) glycohydrolase (PARG) is incapable of cleaving the terminal ADP-ribosyl bond (Slade et al., 2011), which is responsible for the bulk of polymer degradation.

The concentration of NAD⁺, an ADPr donor, varies across different cellular sub-regions because it does not exhibit transmembrane capability. The NMNAT family includes rate-limiting enzymes of the major NAD⁺ synthesis pathway that are located in different cellular compartments. For example, NMNAT1 is located in the nucleus, NMNAT2 is located in the Golgi membrane, and NMNAT3 is located in the mitochondria (Lau et al., 2009; Ryu et al., 2018). The specific subcellular location of these three NAD⁺ synthases leads to the segregation of NAD⁺ generation, which is critical to maintain NAD⁺ homeostasis within the cell (Figure 2).

In 1984, Kawamitsu et al. developed the first antibody against ADP-ribosylation, named 10H; however, it was thought to bind to only PAR, with a polymer length limit of approximately 10 ADPr units (Kawamitsu et al., 1984). Developing MAR-encoded immunogens that recognize MARylation without responding to PAR and adenosine-derived modifications such as adenylation is challenging.

Researchers have examined naturally occurring ADP-ribosylation-binding domains. The WWE domain recognizes only PARylation, whereas Macrodomain2 and 3 of PARP14 are effective in binding to the MARylation targets (Gibson et al., 2017). In 2017 Gibson et al. synthesized a recombinant antibody-like reagent that recognizes mono-, poly-, and oligo-ADP ribosylations based on the discovery of ADPr binding domains (e.g., Macro and WWE) (Gibson et al., 2017). These reagents can recognize all forms of ADP-ribosylation with different specificities collectively. Moreover, the Macro Af1521 domain of bacteria recognizes both MARylation and PARylation (Larsen et al., 2017). Hottiger Laboratory produced an engineered Af1521-macrodomain that was fused to a mouse Fc fragment (Nowak et al., 2020). After that, using MARylated peptides as antigens, Hottiger Laboratory developed a new commercially available polyclonal ADPr antibody specific for MARylated peptides (Hopp et al., 2021), whose efficacy for detecting ADP-ribosylation was demonstrated in a multi-cancer immunohistochemistry analysis (Aimi et al., 2021). For in-depth research, precisely modified peptides and site-specific antibodies must be developed. Next, an antibody was generated against a peptide with a MARylated lysine in 2019 (Lu et al., 2019). In 2020, Bonfiglio et al. developed a generation of pure Ser-ADP-ribosylated peptides and antibodies capable of detecting site-specific histone Ser-ADP-ribosylation (Bonfiglio et al., 2020). Many reagents are available for studying MARylation; however, none of these reagents has been evaluated by comparing (Table 2).

A recent study compared the currently available reagents, enabling researchers to make an informed decision regarding which reagent to use for their specific applications (Weixler et al., 2022). By using Western blot, Slot blot, and confocal living cell imaging, researchers detected ADP-ribosylation in cells on both protein and RNA substrates. Though no perfect method exists at the moment, future research is expected to focus on developing more efficient tools for detecting the specific ADPr-substrate bond.

The construction of ADPr site-specific antibodies requires an in-depth understanding of the modified specific amino acid sites. The ADP-ribosylation reaction is a highly dynamic reversible modification. Furthermore, similar to most PTMs, PARP expression is low in physiological states at the cellular level (Nagaraj et al., 2011), thus requiring the identification and enrichment of mono-ART substrates as a preliminary step before the use of mass spectrometry (MS) analysis. The expression of mono-ARTs is upregulated only under certain conditions such as stress, the presence of interferons, and other cancer-causing factors (Cohen and Chang, 2018). The unstable nature of ADP-ribosylation makes its study challenging. Moreover, it is present in many heterogeneous forms, and its polymers are large and charged.

Combining chemical and genetic engineering methods for the enrichment and identification of ADP-ribosylation sites has been improving over the last decade. A mono-ART substrate detection strategy involves a three-step process that monitors ADP-ribosylation in vitro and in vivo via various components of the labeling reactants. The following sections discuss the progress made in each category (Table 2, Figure 3).

In each of the aforementioned methods, affinity purification methods are used to induce ADP-ribosylation modification before proteomics analysis and can be classified into the following three types (Dani et al., 2009; Jungmichel et al., 2013): 1. affinity purification via labeled vectors, 2. reorganization of ADPr group, and 3. biotin–streptavidin binding system (Table 2, Figure 3).

Because of the dynamic nature and reversibility of ADP-ribosylation, enrichment of ADP-ribosylated substrates by co-immunoprecipitation (Co-IP) would lead to the omission of peptides negatively. Therefore, researchers have used the Macro domain to directly enrich ADP-ribosylation (56). The Af1521 Macro domain has a preference to bind MARylated peptides and a relatively high affinity for ADPr (Kd ∼0.13 μM) (Karras et al., 2005). A study reported that random mutagenesis of wild-type Af1521 led to the development of engineered Af1521 (eAf1521) with a 1000-fold increase in the affinity for ADPr compared with that of the wild-type Af1521 (41). In the proteome ADP-ribosylation MS workflow, its use considerably improved the identification rates of ADP-ribosylation proteins and led to a greater modification coverage.

The biotin and streptavidin systems were developed in the last century. Streptavidin and avidin, collectively known as (strept) avidin, are structurally and functionally similar proteins and have exceptionally high affinity for biotin (Kd ∼10⁻¹⁴–10⁻¹⁶ M), and their interaction with biotin is much stronger than that of the Macro domain. In addition, they are stable in the presence of heat, denaturants, extreme pH, and proteolytic enzymes (Weber et al., 1989). The “click-it” chemistry technology is used to connect biotin to NAD⁺ analogs or ADPr probes, in addition to strep magnetic beads, for the enrichment of MARylate substrates.

The problem encountered in the identification strategy of proteomics is the similarity between ADPr and other abundant cellular molecules such as adenine nucleotides and nucleic acids. In addition, ADP-ribosylation is catalyzed by various ARTs, each with different enzyme activity and preference for amino acids. Of note, mono-ARTs can modify various amino acid residues (Gibson and Kraus, 2012; Schreiber et al., 2006). To date, ADP-ribosylation has been observed on a wide range of amino acid residues (Glu, Asp, Lys, Arg, His, Cys, and Ser) (Gupte et al., 2017). Currently, the following strategies are available for sample preparation and MS analysis (Table 2, Figure 3).

Because PARP1, 2 are mainly found in the nucleus, PARylation in the nucleus has garnered considerable scientific attention, whereas intranuclear mono-ARTs have been less studied. Recently, with the development of MS techniques and related antibodies and probes, different roles of MARylation in the nucleus have been identified, with PARP1-3, 7, 9, 10, and 14 currently reported to be localized in the nucleus (Table 2, Figure 4) (Sanderson and Cohen, 2020). All PARPs except for PARP1, 2 and TNKS1, 2 are mono ADPr moiety writers. Certain mono-ARTs, such as PARP3, 7, exhibit differential localization in the nucleus and the cytoplasm during different phases of the cell cycle (Vyas et al., 2013).

As mentioned previously, the writers of MARylation are distributed throughout the cell. In 2013, a study by Vyas et al. reported that mono-ARTs are predominantly present in the cytoplasm, and data suggest that PARP8, 12, and 16 potential functions for the assembly or maintenance of membranous organelles (Vyas et al., 2013). Vyas et al. knocked down the PARP family in HeLa cells, and mono-ARTs were discovered to be highly associated with cytoskeletal proteins and their functions, including the regulation of membrane structures, cell viability, cell division, and the actin cytoskeleton (Vyas et al., 2013). Similarly, a study by Kliza et al. reported that MARylation occurs primarily in the dynamic homeostasis of regulatory proteins and that the changes in proteostasis remarkably affect cancer development (Kliza et al., (2021). Protein stability in vivo is closely associated with RNA biology and is controlled at many stages, including ribosome biogenesis, ribosomal function, mRNA translation, protein stabilization, protein folding, and removal of misfolded proteins (Kors et al., 2019).

The transcriptional regulation of mono-ARTs by type I and type II interferons (IFNs) indicates their role in immune response and host defence against pathogens (Jiao et al., 2017). At least five human mono-ARTs possessing antiviral activity have been demonstrated on the basis of the following action mechanisms.

The PARP12, 13-mediated RNA degradation pathway is an effective mechanism of host cell defence (Figure 4). The N-terminal regions of PARP12, 13 contain CCCH-type zinc fingers that bind to RNA in a circular conformation which recognizes specific sequences in viral RNA and DNA and degrades retroviral RNA (Bick et al., 2003). PARP10, 12, 13, and 14 are all induced by IFNs and inhibit viral replication (Figure 4) (Atasheva et al., 2012).

PARP7 can bind to TANK-binding kinase 1, a major kinase involved at the beginning of the pathogen-associated molecular pattern pathway, which results in transcription of type I IFN genes. IFN-type I binds to the IFN-α/β receptor and signals via the Janus kinase signal transducer and activator of transcription (JAK/STAT) pathway to induce the expression of hundreds of IFN-stimulated genes, which regulate cellular functions upon the recognition of nucleic acids. The genomic instability of cancer cells can lead to the accumulation of aberrant cytosolic nucleic acids, which, in turn, can activate the pattern recognition receptors (PRRs) (Mackenzie et al., 2017; Paludan et al., 2019). In response to cytosolic nucleic acids accumulated because of pathogens or injury, the PRR pathways, which include cyclic GMP–AMP synthase-stimulator of IFN genes (cGAS-STING) and retinoic acid-inducible gene I, activate type I IFNs to promote innate immunity (Paludan et al., 2019; Ivashkiv and Donlin, 2014; Barber, 2015; Härtlova et al., 2015). At low levels, inflammatory signaling may facilitate cancer growth, whereas, at high levels, it may trigger cell death or immune recognition (Cheon et al., 2014).

As a negative regulator of nucleic acid sensing, PARP7 expression is upregulated in cancer to downregulate IFN signaling. PARP7 inhibitors can cause tumors to release IFN, resulting in tumor regression and persistent immunity (Gozgit et al., 2021). The combination of PARP inhibitors and programmed cell death protein 1 (PD-1)/PD-1 ligand (PD-L1) can activate antigen-presenting cells, such as dendritic cells, via the cGAS-STING pathway and is effective in BRCA1-deficient tumors (Jiao et al., 2017). In IFNγ-stimulated THP-1 cells, proteomics studies found high levels of PARP9 and PARP14, which were increasingly ADP-ribosylated (Higashi et al., 2019). Interestingly, PARP9, 14 exert anti-inflammatory and pro-inflammatory effects on macrophages, respectively, thereby regulating macrophage activation (Iwata et al., 2016). The expression of PARP9 is controlled by the IFNγ-JAK2-STAT1-IFN regulatory factor 1 signaling pathway, which is essential for the survival of cells in diffuse large B-cell lymphoma (DLBCL), where the host inflammatory response is activated (Juszczynski et al., 2006). In non-interleukin (IL)-4-stimulating conditions, PARP14 represses transcription by recruiting histone deacetylase (HDAC) 2 and HDAC3 to the IL-4-responsive promoter. In the presence of IL-4, the catalytic activity of PARP14 releases HDACs from promoters, thus enabling STAT-6 to bind to promoter regions of its target genes and activating STAT6-dependent transcription (Mehrotra et al., 2011).

In addition to altering proteins, ADP-ribosylation reversibly targets DNA and RNA. ADP-ribosylation of DNA was first reported in 2001 with Pierisin-1, which was capable of the MARylation of double-stranded DNA at the N2 position of guanine (Takamura-Enya et al., 2001). In vitro experiments have indicated that PARP3 catalyzed MARylation at the 5′-phosphate terminal of nicked DNA, which can serve as a substrate for DNA ligases (Figure 4) (Belousova et al., 2018). Furthermore, a recent study showed that the PARP2-HPF1-mediated bridging of two DNA breaks activates the PARP2 PARylation of proteins, and it would be interesting to determine whether the same activation mechanism also applies to ADP-ribosylation of DNA ends (Bilokapic et al., 2020).

DNA is not the only nucleic acid substrate that can be ADP-ribosylated. Because phosphorylated RNA ends are chemically similar to phosphorylated DNA ends, these modifications can also be targeted in vitro, thus expanding the range of substrates for ADP-ribosylation (Munnur et al., 2019). The RRM domain of PARP10 potentially contributes to its catalytic activity toward nucleic acids (Jankevicius et al., 2013). A study showed that PARP10, 11, and 15 MARylate single-stranded RNA (ssRNA) at its 5′ end (Todorova et al., 2014). The use of (32P)-labeled NAD⁺ as an ADPr donor indicated that PARP10 ribosylates 5′ and 3′ ends of phosphorylated ssRNA (Chen et al., 2011) (Figure 4).

In addition, homologues of human TRPT1 found in fungi, archaea, and bacteria may form a non-classical structure at the 5′-phosphorylated end of the RNA (22). This non-classical RNA cap may enhance the stability of RNA by protecting its ends from degradation by nucleases. It can also recruit proteins involved in RNA signaling, similarly to the m7GpppN cap of mRNA, which recruits EIF4E to initiate translation. ADP-ribosylation caps are also thought to inhibit translation, and modifications to RNA by TRPT1 and PARP10 have been reported to increase the resistance of oligonucleotide substrates to CIP treatment, which could be related to “RNA capping” (19, 22).

These modifications have not been detected in vivo because of technical challenges, although pieces of evidence suggest that ADP-ribosylation is a potential nucleic acid modification. To conclude, ADP-ribosylation of DNA and RNA promotes the beneficial physiological effects of this modification, thereby revealing its novel cellular functions.

Based on the principle of synthetic lethality, PARP inhibitors can effectively kill tumors that harbor mutations in BRCA1 or BRCA2 genes (Helleday, 2011). Because of the successful results with the use of PARP inhibitors, researchers have begun to pay attention to the association between PARP family compounds and malignancies.

A recent study by Fabio et al. using a newly developed anti-ADP-ribosylation antibody showed heterogeneous ADP-ribosylation staining patterns with a predominant cytoplasmic ADP-ribosylation appearance in most cancers (Aimi et al., 2021), including breast, ovarian, colon, lung, and prostate. In colorectal cancer, the intensity of cytoplasmic ADP-ribosylation staining correlates with metastatic development. In breast and advanced ovarian cancer, cytoplasmic ADP-ribosylation is related to overall patient survival; however, the association is not significant in prostate or lung cancer (Aimi et al., 2021). The results of IHC imply that cytoplasmic ADP-ribosylation is tumor-specific.

The function and expression of mono-ARTs may differ depending on the type of cancer. For example, PARP3, 9 are overexpressed in different types of human cancers, including BRCA1-associated cancers (Beck et al., 2019) and DLBCL (Aguiar et al., 2000). PARP6 is a new member of the PARP family that plays a dual role in various cancers. Evidence suggests that PARP6 expression in human colorectal cancer is associated with a positive prognosis (Qi et al., 2016). Alternatively, another study indicated that the treatment with PARP6 inhibitors might cause apoptosis in breast cancer cells because PARP6 contributes to the maintenance of centrosome integrity in breast cancer cells via MARylation of checkpoint kinase 1 (Wang et al., 2018). PARP7 expression is decreased in cancers such as breast, liver, colorectal, and other types of cancers. A study pointed out that PARP7 favours tumor progression in ovarian cancer (Palavalli Parsons et al., 2021). PARP14 has been shown to be critical for human multiple myeloma cell survival, and PARP14 levels are strongly linked with cancer progression and poor prognosis (Cohen and Chang, 2018). The expression of PARP family molecules in various cancer types has been discussed in detail and summarized in another review (Sha et al., 2021).

Several recent studies have provided an in-depth understanding of the precise molecular mechanisms and the relationship between ADP-ribosylation and malignancy. PARP4, also known as vPARP, along with major vault protein, is involved in cellular transport, cell signaling, immune response, and multidrug resistance (Berger et al., 2009; Siva et al., 2001; Mossink et al., 2003). PARP7 has been implicated in a variety of biological processes, and a subsequent study on PARP7 reported that PARP7 induces microtubule protein MARylation, thereby promoting microtubule instability and possibly regulating the growth and motility of ovarian cancer cells (Palavalli Parsons et al., 2021). In addition, PARP7 functions as a negative feedback regulator for some oncogenic transcription factors, including HIF-1, c-Myc, and estrogen receptor (ER) (Rasmussen et al., 2021). A theme emerging from the literature is that the localization of PARP7 is context-dependent (Figure 4). PARP7 is primarily localized to the nucleus, and infection with the Sindbis virus (SINV) causes PARP7 to accumulate in the cytoplasm (Kozaki et al., 2017).

PLK1 phosphorylates PARP10 and inhibits PARP10-mediated ubiquitination of NEMO, consequently increasing the activity of NF-κB transcription (Tian et al., 2020). In contrast, MARylation of polo-like kinase 1 (PLK1) inhibits the kinase activity and oncogenic function of PLK1 in hepatocellular carcinoma (HCC) (Tian et al., 2020). PARP10 mono-ADP-ribosylates Aurora A and inhibits its kinase activity, thereby playing an essential role in tumor proliferation and metastasis suppression (Zhao et al., 2018). PARP14 maintains low PKM2 activity in HCC cells by suppressing JNK1, which promotes the Warburg effect and promotes cancer cell proliferation and survival (Iansante et al., 2015). Recently, Challas et al. demonstrated that NMNAT-2 increases the catalytic activity of PARP16, which promotes protein homeostasis in ovarian cancer cells regulating the translation of specific mRNAs to avoid harmful protein aggregation (Challa et al., 2021).

With a better understanding of the role of mono-ARTs in carcinogenesis and progression, these molecules are gradually emerging as potential targets for cancer treatment. The development of selective inhibitors of mono-ARTs is garnering increasing attention. Presently, selective inhibitors of MARylating PARPs are available for PARP4, 6, 7, 10, 11, 14, and 16, and only one PARP7 inhibitor, RBN-2397 (Gozgit et al., 2021), is currently under phase I clinical trial (ClinicalTrials.gov identifier: NCT04053673).

By screening a library of compounds for the ability to induce mitotic defects, researchers have identified AZ0108 as a potent PARP6 inhibitor, which exerts antitumor effects in vivo and induces cell death in breast cancer cells in vitro (Wang et al., 2018).

PARP10 is an intriguing target for cancer treatment because it regulates cell proliferation through various processes, including the regulation of ß-catenin and the alleviation of replication and oxidative stress (Schleicher et al., 2018; Wu et al., 2020). Venkannagari et al. established the conditions conducive to the screening of ART inhibitors and identified OUL35, a selective inhibitor of PARP10 (Venkannagari et al., 2016). Murthy et al. modified OUL35 and developed a compound, 4-(benzyloxy) benzamide derivative, which is potent (IC50 = 230 nM) and selective, and like OUL35, it could rescue HeLa cells from PARP10-induced cell death (Murthy et al., 2018). Based on OUL35, Maksimainen et al. developed mono-ART inhibitors by supplementing the TIQ-A scaffold (PARP1 inhibitor) with slight structural changes, which changed the selectivity of the inhibitors from poly-ARTs to mono-ARTs (Maksimainen et al., 2021).

In 2018, Kirby et al. generated a PARP11 selective inhibitor, ITK7, by exploiting structural differences in the active regions of PARPs that facilitate MARylation versus PARylation (Kirby et al., 2018). Recently, Kirby et al. developed a selective PARP4 inhibitor, AEP07, by utilizing structural bioinformatics approaches to target a unique threonine residue (Thr484) in the PARP4 nicotinamide sub-pocket (Kirby et al., 2021).

Owing to the structural similarity of the catalytic domains of the numerous PARP family members, identifying selective PARP inhibitors might be challenging. Thus, addressing other distinctive structural properties of PARPs, such as large domains, may offer a further avenue for developing inhibitors. All of the inhibitors target the PARPs’ catalytic domain, except for the PARP14 inhibitor GeA-69, a kinase inhibitor, which acts as an inhibitor of PARP14 Macrodomain 2 (Moustakim et al., 2018). Potential PARP16 inhibitors decrease the phosphorylation of PERK and IRE1αgenerated by ER stress, eventually promoting cell death (Wang et al., 2017).

A multidisciplinary approach expands the chemical space of mono-ART inhibitors and provides new leads for understanding selectivity in mono-ART inhibition. Wigle et al. developed an active site probe, NanoBRET, which can be used to investigate cellular residence times of PARP inhibitors in live cells (Wigle et al., 2020). The development of more clinically effective and selective mono-ART inhibitors will be helpful in cancer treatment. Of note, these inhibitors may be used in the treatment of non-oncological disorders such as protection against oxidative stress, reduction of inflammatory responses, and neurological diseases (Marcus et al., 2021).