ADP-ribosylation (ADPRylation) is a regulatory post-translational modification (PTM) of proteins that results in the reversible attachment of ADP-ribose (ADPR) units on substrate proteins (Gupte et al., 2017; Lüscher et al., 2018). Members of the PARP family of ADP-ribosyltransferases (ARTs) (Amé et al., 2004; Vyas et al., 2013) function as ‘writers’ to catalyze the transfer of ADPR moieties from oxidized β-nicotinamide adenine dinucleotide (NAD⁺) to a variety of amino acids (Asp, Glu, Ser, Arg, and Lys) in substrate proteins (Gupte et al., 2017; Lüscher et al., 2018). Mono(ADP-ribosyl) transferases (MARTs) add a single ADPR moiety to their substrates through a process called mono(ADP-ribosyl)ation (MARylation) (Challa et al., 2021), whereas poly(ADP-ribosyl) transferases (PARPs) add branched or linear chains of multiple ADPR moieties through a process called poly(ADP-ribosyl)ation (PARylation) (Gupte et al., 2017; Lüscher et al., 2018). Site-specific ADPRylation of substrate proteins by PARP enzymes can have important functional consequences, including alteration of the biochemical or biophysical properties of the substrate protein or creation of new binding sites for ADPR binding domains (ARBDs) that drive protein-protein interactions (Gibson and Kraus, 2012; Gupte et al., 2017). As such, ADPRylation can control a wide variety of cellular and biological processes, including DNA repair, DNA replication, gene expression, and RNA biology, as well as inflammatory responses and cell differentiation (Challa et al., 2021; Gibson and Kraus, 2012; Gupte et al., 2017; Lüscher et al., 2018).

The various forms of ADPR are recognized and bound by an assortment of protein domains and motifs that are found in a variety of proteins with diverse functions and mediate many of the biological functions of ADPRylation (Barkauskaite et al., 2013; Gibson and Kraus, 2012; Teloni and Altmeyer, 2016). These ARBDs function as ‘readers’ of the various forms of protein-linked ADPR (MAR, PAR, branched, terminal residue, etc.). They include PAR-binding motifs (PBMs), macrodomains, PAR-binding zinc fingers (PBZs), and WWE domains (Ahel et al., 2008; Feijs et al., 2013; Gagné et al., 2008; Karras et al., 2005; Pleschke et al., 2000; Rack et al., 2016; Wang et al., 2012). Macrodomains recognize free ADPR, as well as the terminal ADPR moieties in PAR, allowing them to bind to both MAR and PAR (Karras et al., 2005; Timinszky et al., 2009). PBZ domains recognize branched forms of PAR (Chen et al., 2018). WWE domains recognize the iso-ADPR linkages joining ADPR monomers, restricting their binding to PAR (Kang et al., 2011; Wang et al., 2012; Zhang et al., 2011). In addition to the ‘writers’ and ‘readers,‘ ‘eraser’ enzymes, including PAR glycohydrolase (PARG) and ADP-ribosylhydrolase 3 (ARH3), recognize specific ADPR modifications through the ARBDs and catalyze PAR chain degradation through endo- and exoglycocidic activities. Their activities leave the terminal ADPR moiety attached to the acceptor amino acid residue of the substrate (Barkauskaite et al., 2013; Niere et al., 2012; Oka et al., 2006; Slade et al., 2011).

Although recent developments in mass spectrometry-based identification of ADPR-modified amino acids have enhanced the study of specific ADP-ribosylation events on target proteins (Daniels et al., 2015), the lack of a complete set of immunological tools that recognize the diverse forms of ADPR has hampered progress in studying ADPRylation. Anti-ADPR polyclonal antibodies have been reported, but the specificity and utility of these antibodies have not been assessed broadly (Bredehorst et al., 1978; Kanai et al., 1974; Kanai et al., 1978; Meyer and Hilz, 1986; Sakura et al., 1978). Instead, the PARP field has relied on the anti-PAR monoclonal antibody 10H, which binds to PAR chains longer than ten ADPR units (Kawamitsu et al., 1984). Although useful, this antibody has left the field blind to mono- and oligo(ADP-ribosyl)ation.

The recent development of recombinant site-specific and broad-specificity antibodies to ADPR has been a major advance (Bonfiglio et al., 2020). We recently described the generation and characterization of a set of recombinant antibody-like ADP-ribose binding proteins, in which natural ARBDs have been functionalized with the Fc region of rabbit immunoglobulin (Gibson et al., 2017). Collectively, the ADPR detection tools described here are useful for cellular and biochemical assays, but they are not useful for exploring the dynamics of ADPRylation in cells and in animals. In fact, ADPRylation is a rapid process that can occur within minutes and can be removed by various ‘erasers’ including ARH3 and PARG (Barkauskaite et al., 2013; Gupte et al., 2017). Therefore, developing tools to measure ADPR dynamics in cells and in vivo is critical for better understanding the various biological processes mediated by ADPR.

To overcome this limitation, several split-protein reassembly approaches have been applied to PAR detection. With this approach, nonfunctional fragments of a split-fluorescent protein or luciferase are induced to reassemble through the direct interaction of fused ARBDs (Furman et al., 2011; Krastev et al., 2018; Lee et al., 2021; Serebrovskaya et al., 2020). These include the PBZ modules of aprataxin PNK-like factor (APLF) (Ahel et al., 2008) attached to each half of split firefly luciferase (split-Fluc) (Furman et al., 2011), PBZ modules with split Venus GFP (Krastev et al., 2018), and WWE domains with Turquoise and Venus, allowing for Förster resonance energy transfer (FRET) (Serebrovskaya et al., 2020). However, these tools have some limitations: (1) they can only detect PAR accumulation in vitro (Furman et al., 2011), (2) they can only detect PAR accumulation on specific target proteins (Krastev et al., 2018), or (3) they have modest dynamic ranges (Serebrovskaya et al., 2020). Moreover, none of these sensors has been shown to be capable of detecting PAR accumulation in vivo.

In the work described herein, we developed a set of PAR Trackers (PAR-Ts)—optimized dimerization-dependent and split-protein reassembly PAR detection tools that have broad utility for both in vitro and in vivo studies. The PAR-Ts contain a WWE fused to both parts of dimerization-dependent GFP (ddGFP) (Alford et al., 2012) or split Nano luciferase (NanoLuc) (Wang et al., 2020) with LSSmOrange. The ddGFP version (PAR-T ddGFP) allows for real-time assessment of dynamic PAR production in vitro and in living cells, while the split NanoLuc version (PAR-T NanoLuc) allows detection of PAR production in tissues in living mammals.

The use of functionalized ARBDs to detect ADP-ribosylation has been a useful approach (Forst et al., 2013; Gibson et al., 2017; Timinszky et al., 2009). In this study, we further developed our previous ADPR detection reagents (Gibson et al., 2017) to expand their utility to in vivo applications as ADPR sensors. To achieve this, we used a systematic approach including in vitro characterization of the sensors and validation of their utility in vivo.

Using dimerization-dependent GFP-based sensors to detect PAR in Vitro

Dimerization-dependent GFP is a genetically encoded sensor that was initially developed to study protein interactions (Alford et al., 2012). In this system, a pair containing a quenched GFP (ddGFPA) and a nonfluorogenic GFP (ddGFPB) form a heterodimer with improved fluorescence (Alford et al., 2012). The reversible complementation of ddGFP pairs, unlike irreversible split fluorophores, is ideal to monitor dynamic signaling events, such as PARylation (Villalobos et al., 2007). Hence, we sought to design ddGFP-based fluorescence sensors for PAR (fluorescent PAR-Trackers or PAR-T ddGFP) (Table 1) to enable us to perform live-cell imaging, with a high signal-noise ratio. To achieve this, we fused various ARBDs to ddGFP-A/B and purified the recombinant proteins (Figure 1A and B, Figure 1—figure supplement 1A). We performed in vitro ADP-ribosylation assays using recombinant PARP-1 (to generate PAR) and PARP-3 (to generate MAR) (Figure 1—figure supplement 1B and C). We observed that of all the ARBD-ddGFP pairs tested, the WWE domain from RNF146, macrodomain from AF1521, and a combination of these two performed well in specifically recognizing PARylated PARP-1 (Figure 1—figure supplement 1D and E). These sensors recognized PARylated-PARP-1, but not MARylated PARP-3, or the precursors of ADPR (Figure 1C).

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Table 1

Nomenclature used in the manuscript	ARBDs used with fluorescent or luminescent protein fragments		Activity in vitro	Activity in cells	Figures
Fluorescent sensors	ddGFP-A	ddGFP-B
PAR-T ddGFP	WWE from RNF146	WWE from RNF146	+++	++++	Figure 1—figure supplement 1D, E
	Macrodomain from AF1521	Macrodomain from AF1521	++	N.D.	Figure 1—figure supplement 1D, E
	PBZ from APLF	PBZ from APLF	+	N.D.	Figure 1—figure supplement 1D, E
	Macrodomain from MH2A.1	Macrodomain from MH2A.1	+	N.D.	Figure 1—figure supplement 1D, E
	WWE from RNF146	Macrodomain from AF1521	+++	N.D.	Figure 1—figure supplement 1D, E
	Macrodomain from AF1521	WWE from RNF146	++++	–	Figure 1—figure supplement 1D, E
Firefly luciferase-based sensors	N-terminal of Firefly luciferase (FLucN)	C-terminal of Firefly luciferase (FLucC)
PAR-T fLuc	WWE from RNF146	WWE from RNF146	N.D.	+	Figure 4—figure supplement 1C
	Macrodomain from AF1521	Macrodomain from AF1521	N.D.	–	Figure 4—figure supplement 1C
	WWE from RNF146	Macrodomain from AF1521	N.D.	–	Figure 4—figure supplement 1C
	Macrodomain from AF1521	WWE from RNF146	N.D.	–	Figure 4—figure supplement 1C
Nano luciferase-based sensors	N-terminal of Nano luciferase (NanoLucN)	C-terminal of Nano luciferase plus LssmOrange (NanoLucC)
PAR-T NanoLuc	WWE from RNF146	WWE from RNF146	N.D.	++++	Figure 4C

We further tested the sensitivity of these sensors and their dynamic range using in vitro PARP-1 PARylation reactions with increasing concentrations of NAD⁺ (Figure 1—figure supplement 1F and G) and increasing time of reaction (Figure 1D and E). Similarly, we detected the degradation of PAR chains by the ADP-ribosylhydrolase ARH3 in a time (Figure 1F and G) and dose (Figure 1—figure supplement 1H) dependent manner. Further, we performed in vitro reactions by incubating the recombinant PAR-T sensors with lysates from HeLa cells treated with H₂O₂ to induce DNA damage and activate PARP-1, or an inhibitor of the PAR glycohydrolase PARG (i.e., PDD00017273) to increase PAR. We observed an increase in fluorescence when lysates from cells treated with either H₂O₂ or PARG inhibitor (PDD00017273) were used, as well as a profound increase in fluorescence when lysates from cells treated with both H₂O₂ and PARG inhibitor were used (Figure 1—figure supplement 1I and J). The H₂O₂- and PARG inhibitor-stimulated signals were reduced with PARP inhibitor treatment (Figure 1—figure supplement 1I and J). Taken together, these data suggest that the PAR-T ddGFP sensors can specifically recognize PAR, and that they exhibit a good dynamic range in vitro.

Using PAR-T ddGFP sensors to detect PAR in live cells

Having confirmed the specificity of the PAR-T sensors, we next sought to test their utility in live-cell imaging. We expressed the PAR-T sensors in HeLa cells using doxycycline (Dox) induction and performed live-cell imaging after subjecting the cells to H₂O₂-mediated PARP-1 activation (Figure 2—figure supplement 1A). The live cell PAR-T construct also expresses mCherry with a nuclear localization signal (NLS) to illuminate the nuclei and act as a control for the expression of the constructs (Figure 2—figure supplement 1A). When compared to ddGFP alone, ddGFP-conjugated to WWE detected PARP-1 activation in live-cell imaging (Figure 2A and B). Interestingly, even though the WWE-macrodomain combination sensor was able to detect PAR in vitro, this sensor combination failed to recognize PAR in cells (Figure 2—figure supplement 1B and C). Hence, we used the WWE-based PAR-T sensors for the experiments from this point onwards. Using the WWE-based ddGFP PAR-T sensor, we were able to detect accumulation of PAR after H₂O₂-treatment in real time. Treatment with PARP inhibitor blocked this accumulation (Figure 2C and D; Figure 2—video 1).

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Cancers are heterogenous tissues with spatial variation in nutrient availability and cell-extrinsic stressors (Dagogo-Jack and Shaw, 2018). PAR is enhanced by stressors, such as DNA damage or hypoxia, but the spatiotemporal dynamics of PAR production in cells have remained unclear due to a lack of efficient detection methods. Thus, we asked if PAR levels vary spatially in groups of cells. We performed live-cell imaging in three-dimensional (3D) cancer spheroids using MCF-7 human breast cancer cells expressing the WWE-based PAR-T-ddGFP sensor. We observed a heterogeneous distribution of PAR throughout the spheroid, which was inhibited by the PARP inhibitor, Niraparib (Figure 3A and B). We also performed a time course of Niraparib treatment in 3D cancer spheroids to visualize spatio-temporal changes in PAR levels over time. The results indicate that the PAR levels in cells at the core of the spheroids are relatively resistant to Niraparib treatment, since the PAR levels in these cells decrease at a lower rate compared to the PAR levels in the cells in the outer layer of the spheroid (Figure 3C and D). These results show that the PAR-T-ddGFP sensor can be used for live-cell imaging to evaluate spatial and temporal changes in PARylation in cancer cells.

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Developing a highly sensitive split-luciferase PAR-T sensor

Since we demonstrated that WWE domain-based PAR-T sensors can specifically detect PAR in biochemical assays and living cells, we sought to develop a set of highly sensitive PAR-T sensors that can detect PAR in vivo. Fluorescent sensors are not well suited for detection in vivo due to auto-fluorescence of tissues that can cause high background signals. Instead, luminescence-based approaches are preferred for in vivo applications due to the lack of auto-luminescence in tissues (Tung et al., 2016). Hence, we developed a set of luminescent PAR-T sensors to detect PAR levels in vivo. We first generated sensors based on split firefly luciferase (Maita et al., 2014) using various combinations of the ARBDs. As before, the WWE domains consistently performed better in detecting an increase in PAR levels with PARG inhibitor treatment and a decrease in PAR levels with a PARP inhibitor treatment using either cell lysates (Figure 4—figure supplement 1A) or live cells (Figure 4—figure supplement 1B-D). Luminescence from an unsplit Firefly luciferase remained unaltered with these treatments (Figure 4—figure supplement 1D and E).

Although split firefly luciferase-based approaches were capable of detecting signals from cells in vitro, the signal intensity from the split firefly luciferase-based PAR-T sensors were ~10,000-fold less than intact firefly luciferase, which makes it difficult to use this sensor in vivo. This limitation could be due to the bulkiness of firefly luciferase, which may interfere with the function of the domains fused to them in complementation assays (Wang et al., 2020; Yano et al., 2018). Therefore, we decided to employ a Nano luciferase (NanoLuc)-based split luciferase complementation system (Dixon et al., 2016; Hall et al., 2012), which is a smaller, brighter, and more stable luciferase compared to firefly luciferase (Figure 4A).

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To quantitatively assess the activity of this luminescent PAR-Tracker (PAR-T NanoLuc), we expressed it in a Dox-dependent manner in human breast cancer cells that also stably express firefly luciferase (MDA-MB-231-Luc cells) (Figure 4A). In this way, we had an internal standard (i.e., the signal from the firefly luciferase), which allowed us to account for changes in cell viability or tumor size in these experiments. We first tested if there was cross-reactivity of the two luciferases (NanoLuc and firefly luciferase) to the substrates; we observed specific detection of firefly luciferase with D-Luciferin and NanoLuc with furimazine with no cross-reactivity (Figure 4B-D). Moreover, the luminescence of PAR-T NanoLuc is only 30-fold lower than intact firefly luciferase (Figure 4C). PARP-1 depletion reduced the luminescence from PAR-T NanoLuc with little effect on the luminescence of firefly luciferase (Figure 4E-G). Interestingly, knockdown of PARP-2 had no effect on luminescence from PAR-T NanoLuc. Nevertheless, the luminescent PAR-T NanoLuc sensor is highly sensitive and can be used to detect PAR in 1000 cells with a dynamic range of approximately threefold (minimum to maximum) (Figure 4—figure supplement 2).

DNA damaging agents, such as UV irradiation and gamma irradiation, activate PARP-1 and promote auto and trans PARylation of PARP-1 and other DNA damage repair proteins, respectively, that are recruited to sites of DNA damage (Ray Chaudhuri and Nussenzweig, 2017). Since the PAR-T sensor can detect H₂O₂-induced PARP-1 activation (Figure 2), we assessed whether it can detect radiation-induced PARP-1 activation. We subjected the MDA-MB-231-Luc cells to Dox-induced expression of PAR-T NanoLuc and then exposed the cells to UV radiation. We observed that UV radiation-induced PARP-1 activation as assessed by an accumulation of PAR on Western blots (Figure 5A). This was further enhanced by inhibition of PARG, whereas inhibition of PARP-1 blocked the UV-induced PARP-1 activation (Figure 5A). UV radiation of PARG inhibitor-treated cells enhanced PAR-T luminescence, whereas UV radiation of PARP inhibitor-treated cells reduced the PAR-T luminescence (Figure 5B and C). None of these treatments affected the luminescence from firefly luciferase (Figure 5B and C).

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In a similar manner, we also performed a time course of UV-mediated PARP-1 activation using live cell luminescence assay with the PAR-T NanoLuc sensor. We subjected the MDA-MB-231 cells to Dox-induced expression of PAR-T NanoLuc or intact Nano luciferase and then exposed the cells to UV radiation. Consistent with the previous experiment, we observed a time-dependent increase in PAR-T NanoLuc signal in vehicle-treated cells, but not in Niraparib-treated cells. Interestingly, UV-mediated increases in PARP-1 activation were more spontaneous in PARG inhibitor-treated cells (Figure 5D). The PAR levels under basal (–UV) conditions were low, resulting in only a 50% decrease in PAR-T NanoLuc signal with Niraparib treatment (Figure 5A-C). The decrease in PAR-T NanoLuc signal was greater when UV-treated cells were pre-treated with Niraparib, which is consistent with the results from Western blot analysis (Figure 5A).

Comparison of assay performance using the PAR-T sensor and conventional PAR detection reagents

Next, we performed a set of assays to compare the performance of the PAR-T sensors to conventional PAR detection reagents (WWE-Fc and PAR antibody) in a variety of assays. In sum, we compared: (1) Western blotting with WWE-Fc versus fluorescence assay with PAR-T ddGFP, which were performed in conjunction with ARH3-mediated degradation of PAR in vitro (Figure 6A); (2) Western blotting with WWE-Fc versus live-cell luciferase assay using PAR-T NanoLuc, which were performed in conjunction with UV-induced DNA damage in MDA-MB-231 cells (Figure 6B); (3) enzyme-linked immunosorbent assay (ELISA) with PAR antibody versus fluorescence assay with PAR-T ddGFP, which were performed using immobilized PAR (Figure 6C); and (4) immunofluorescence with WWE-Fc versus live-cell imaging using PAR-T ddGFP, which were performed using H₂O₂-mediated PARP-1 activation in 293T cells (Figure 6D and E).

Figure 6

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We compared the dynamic ranges of the PAR-T sensors with the other reagents (WWE-Fc and PAR antibody) in various PAR detection assays, such as Western blotting and ELISA (Figure 6F). We found that Western blotting with WWE-Fc had the highest dynamic range for detection of PAR (eightfold), but the dynamic range of live-cell luciferase assay with PAR-T NanoLuc was comparable (sixfold) (Figure 6F). While PAR-T ddGFP in a modified fluorescence assay had a larger dynamic range than PAR antibody in an ELISA (6-fold vs. 3.5-fold) when the assays were performed using immobilized PAR, it had a lower dynamic range when used for live-cell imaging (4.4-fold). This can be explained, in part, by the higher autofluorescence of cells, which can diminish the dynamic range of the PAR-T ddGFP sensors. Thus, this sensor requires further optimization to increase the signal-noise ratio. Nevertheless, the performance of PAR-T ddGFP in live cells is comparable to that of an immunofluorescence assay with WWE-Fc (4.4-fold vs. 5-fold).

Detection of PAR production from PARP-1 activation under physiological conditions

We previously showed that PARP-1 catalytic activity decreases during the initial differentiation of preadipocytes (Huang et al., 2020; Luo et al., 2017; Ryu et al., 2018). Thus, adipogenesis is a unique biological process to study the dynamics of PAR accumulation from changes in PARP-1 activity under physiological conditions. We used the PAR-T NanoLuc sensor to investigate changes in PARP-1 activity during early adipogenesis of murine preadipocytes (i.e., 3T3-L1 cells). We observed a decrease in the signal from PAR-T NanoLuc by 12 hr of differentiation and a greater reduction in PAR-T NanoLuc signal by 24 hr of differentiation (Figure 7), consistent with our previous observation that PARP-1 activation decreases precipitously during adipogenesis (Huang et al., 2020; Luo et al., 2017; Ryu et al., 2018). These results further highlight the high sensitivity of PAR-T NanoLuc sensor, which can be used to study physiological changes in PAR levels during biological processes, such as adipogenesis.

Figure 7

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Detection of PAR production from PARP-1 activation in vivo

To assess the in vivo utility of PAR-T NanoLuc, we established xenograft tumors in NOD/SCID/gamma (NSG) mice using MDA-MB-231-Luc cells and induced the expression of PAR-T NanoLuc with Dox (Figure 8A). Similar to the in vitro experiments, we detected an increase in PAR levels with the PAR-T NanoLuc sensor when the gamma-irradiated mice were treated with PARG inhibitor, but the signal was decreased when the gamma-irradiated mice were treated with PARP inhibitor (Figure 8B and Figure 8—figure supplement 1A). We normalized the signal from PAR-T to luminescence from firefly luciferase to confidently measure the differences in PAR levels, while accounting for the variability in tumor sizes (Figure 8C).

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We also measured PAR accumulation in breast cancer cells injected into C57/BL6 mice without establishing xenograft tumors over a 24-hr time course post injection, with or without PARG inhibitor treatment (Figure 8—figure supplement 1B-D). PAR accumulation was readily detected in the breast cancer cells injected into the mice in the absence of treatment. Upon treatment with PARG inhibitor, the luminescence from PAR-T NanoLuc increased significantly by 6 hr and then diminished by 24 hr. These results demonstrate that the PAR-T NanoLuc sensor has sufficient sensitivity to detect dynamic changes in PAR production in tissues of living animals in vivo.

Naturally occurring ARBDs have been invaluable tools for developing novel ADPR detection reagents and sensors (Forst et al., 2013; Gibson et al., 2017; Timinszky et al., 2009). In this study, we developed a set of PAR sensors that are useful tools for in vitro assays, live cells, and tissues in living animals (Table 1). To this end, we constructed a ddGFP-based fluorescent PAR-Tracker (PAR-T) that can be used for in vitro assays and live-cell imaging, to track dynamic changes in PAR levels in a single cell (Figure 2) and spatially resolve PAR accumulation in tissue-like spheroids (Figure 3). In addition, we made a split NanoLuc-based luminescent PAR-T containing LSSmOrange that can be used to track physiological changes in PAR levels during differentiation (Figure 7). It is extremely sensitive and can be used to detect PAR in xenograft tumors in living mice (Figure 8).

All unprocessed Western blot image data and data from individual replicates from the fluorescent and luminescent assays can be found on Mendeley at https://data.mendeley.com/datasets/x9j73tdb5r/1. The work does not contain any high complexity, high content "omics" data.

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Author details

1. Sridevi Challa

   1. Laboratory of Signaling and Gene Regulation, Cecil H. and Ida Green Center for Reproductive Biology Sciences, University of Texas Southwestern Medical Center, Dallas, United States
   2. Division of Basic Research, Department of Obstetrics and Gynecology, University of Texas Southwestern Medical Center, Dallas, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Validation, Visualization, Writing – original draft, Writing – review and editing

   Contributed equally with

   Keun W Ryu

   Competing interests

   K.W.R., S.C., and W.L.K. have a patent pending for the PAR-T sensors described herein

2. Keun W Ryu

   1. Laboratory of Signaling and Gene Regulation, Cecil H. and Ida Green Center for Reproductive Biology Sciences, University of Texas Southwestern Medical Center, Dallas, United States
   2. Division of Basic Research, Department of Obstetrics and Gynecology, University of Texas Southwestern Medical Center, Dallas, United States

   Present address

   Cancer Biology & Genetics Program, Memorial Sloan Kettering Cancer Center, New York, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Validation, Visualization, Writing – review and editing

   Contributed equally with

   Sridevi Challa

   Competing interests

   K.W.R., S.C., and W.L.K. have a patent pending for the PAR-T sensors described herein

3. Amy L Whitaker

   1. Laboratory of Signaling and Gene Regulation, Cecil H. and Ida Green Center for Reproductive Biology Sciences, University of Texas Southwestern Medical Center, Dallas, United States
   2. Division of Basic Research, Department of Obstetrics and Gynecology, University of Texas Southwestern Medical Center, Dallas, United States
   3. Program in Genetics, Development, and Disease, Graduate School of Biomedical Sciences, University of Texas Southwestern Medical Center, Dallas, United States

   Contribution

   Data curation, Formal analysis, Investigation

   Competing interests

   No competing interests declared

4. Jonathan C Abshier

   1. Laboratory of Signaling and Gene Regulation, Cecil H. and Ida Green Center for Reproductive Biology Sciences, University of Texas Southwestern Medical Center, Dallas, United States
   2. Division of Basic Research, Department of Obstetrics and Gynecology, University of Texas Southwestern Medical Center, Dallas, United States

   Contribution

   Investigation

   Competing interests

   No competing interests declared

5. Cristel V Camacho

   1. Laboratory of Signaling and Gene Regulation, Cecil H. and Ida Green Center for Reproductive Biology Sciences, University of Texas Southwestern Medical Center, Dallas, United States
   2. Division of Basic Research, Department of Obstetrics and Gynecology, University of Texas Southwestern Medical Center, Dallas, United States

   Contribution

   Data curation, Formal analysis, Investigation, Methodology

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-1723-579X

6. W Lee Kraus

   1. Laboratory of Signaling and Gene Regulation, Cecil H. and Ida Green Center for Reproductive Biology Sciences, University of Texas Southwestern Medical Center, Dallas, United States
   2. Division of Basic Research, Department of Obstetrics and Gynecology, University of Texas Southwestern Medical Center, Dallas, United States

   Contribution

   Conceptualization, Funding acquisition, Project administration, Supervision, Writing – review and editing

   For correspondence

   LEE.KRAUS@utsouthwestern.edu

   Competing interests

   W.L.K. is a founder and consultant for Ribon Therapeutics, Inc and ARase Therapeutics, Inc He is also coholder of U.S. Patent 9,599,606 covering the ADP-ribose detection reagent used herein, which has been licensed to and is sold by EMD Millipore

   "This ORCID iD identifies the author of this article:" 0000-0002-8786-2986

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