PARP7 mRNA levels are lower in ovarian cancer patient samples compared to non-cancerous tissue, but PARP-7 protein nonetheless contributes to several cancer-related biological endpoints in ovarian cancer cells . Global gene expression analyses in ovarian cancer cells subjected to PARP-7 depletion indicate biological roles for PARP-7 in cell-cell adhesion and gene regulation. To identify the MARylated substrates of PARP-7 in ovarian cancer cells, we developed an NAD+ analog-sensitive approach, which we coupled with mass spectrometry to identify the PARP-7 ADP-ribosylated proteome in ovarian cancer cells, including cell-cell adhesion and cytoskeletal proteins. Specifically, we found that PARP-7 MARylates α-tubulin to promote microtubule instability, which may regulate ovarian cancer cell growth and motility. In sum, we identified an extensive PARP-7 ADP-ribosylated proteome with important roles in cancer-related cellular phenotypes.PARP-7 (TiPARP) is a mono(ADP-ribosyl) transferase whose protein substrates and biological activities are poorly understood. We observed that  Cancer is a complex illness where changes inside healthy cells causes them to grow and reproduce rapidly. Specialized proteins called enzymes – which regulate chemical reactions in the cell – often help cancer develop and spread through the body. One such enzyme called PARP-7 labels other proteins by attaching a chemical group which changes their behavior. However, it was unknown which proteins PARP-7 modifies and how this tag alters the actions of these proteins.To investigate this, Parsons, Challa, Gibson et al. developed a method to find and identify the proteins labelled by PARP-7 in ovarian cancer cells taken from patients and cultured in the laboratory. This revealed that PARP-7 labels hundreds of different proteins, including adhesion proteins which affect the connections between cells and cytoskeletal proteins which regulate a cell’s shape and how it moves.One of the cytoskeletal proteins modified by PARP-7 is α-tubulin, which joins together with other tubulins to form long, tube-like structures known as microtubules. Parsons et al. found that when α-tubulin is labelled by PARP-7, it creates unstable microtubules that alter how the cancer cells grow and move. They discovered that depleting PARP-7 or mutating the sites where it modifies α-tubulin increased the stability of microtubules and slowed the growth of ovarian cancer cells.Ovarian cancer is the fifth leading cause of cancer-related deaths among women in the United States. A new drug which suppresses the activity of PARP-7 has recently been developed, and this drug could potentially be used to treat ovarian cancer patients with high levels of PARP-7. Clinical trials are ongoing to see how this drug affects the behavior of cancer cells in patients. Of these four tissues, PARP7 mRNA was expressed to the highest levels in ovary and the lowest levels in the pancreas  on OVCAR4 human ovarian cancer cells subjected to siRNA-mediated nockdown . We usednockdown . Gene onnockdown . The resnockdown , while inockdown .To understand how loss of PARP-7-mediated MARylation results in the altered cellular state and gene expression found following PARP-7 knockdown, we sought to identify the direct protein substrates of PARP-7. For these studies, we focused on cytoplasmic substrates because PARP-7 is predominantly localized in the cytoplasm of OVCAR4 cells . To iden+ binding pocket of a PARP protein is mutated to a smaller residue to create a void (‘hole’), which typically reduces the affinity of the PARP for NAD+ and, hence, its catalytic activity. Catalytic activity can be restored, however, by adding a bulky moiety onto NAD+ (‘bump’), which fills the void and restores binding  reactions  and two have azide-containing R groups    to facil3 and 4)  of MARyl analogs . The dif analogs .+ analogs T-NAD+ for our subsequent studies. PARP-7 S563G, referred to from here forward as asPARP-7, had the highest activity with 8-Bu(3-yne)T-NAD+, a well-characterized and readily available ‘click’-able NAD+ analog T-NAD+ and purified recombinant wild-type or asPARP-7 protein. 8-Bu(3-yne) T-ADP-ribose-labeled proteins were ‘clicked’ to an azido-TAMRA fluorophore and imaged using in-gel fluorescence T-NADrescence . We foun+ and asPARP-7, and then clicked the 8-Bu(3-yne)T-ADP-ribose–labeled proteins to azide-agarose. Following extensive washing of the agarose beads, we performed trypsin-based identification of the ADP-ribosylated proteins by mass spectrometry T-NADtrometry  and 4. Wtrometry . GO analtrometry . These rPARP7 knockdown and assayed by western blotting for the presence of α-tubulin, confirming that PARP-7 was required for MARylation of α-tubulin . Ectopic expression of the catalytically dead PARP-7 mutant resulted in a loss of α-tubulin MARylation , which we expressed using a Flag-tagged mouse Rylation . In addity assay  and the ty assay  T-ADP-ribose-labeled proteins from the azide-agarose beads described above using hydroxylamine and analyzed them by mass spectrometry. Hydroxylamine-cleaved ADPR modifications produce a 15.019 fication . Our anafication . We focufication , and ectty assay . These rCollectively, these results indicate that PARP-7 reduces cellular microtubule content following recovery from depolymerization, likely in part through direct MARylation of α-tubulin. Moreover, the PARP-7-mediated microtubule control may play a role in the regulation of cancer cell growth and motility.Studies over the past two decades have begun to elucidate the biological functions of PARP-7. The earliest studies focused on the role of PARP-7 as a component of the AHR signaling pathway in response to the environmental contaminant dioxin (TCDD) . This le+ analog-sensitivity comprising an NAD+ binding pocket PARP-7 mutant (S563G) paired with the NAD+ analog 8-Bu(3-yne)T-NAD+ T-NAD+ –5. We use)T-NAD+ . Interes+ , a defined cellular process (i.e. microtubule control), and a broader biological outcome (i.e. cancer-related phenotypes).Among the PARP-7 substrates that we identified in our analyses, we found other PARP family members, including PARP-1 (both replicates from OVCAR4 and HeLa cells), PARP-4 (one replicate from OVCAR4 cells and both replicates from HeLa cells), and PARP-2 (one replicate from HeLa cells) . PARPs 1TIPARP  was identified in a susceptibility locus for ovarian cancer in a recent genome-wide association study  is now in Phase I clinical trials for solid tumors . In contrast to the current FDA-approved PARP1/2 inhibitors, RBN-2397 is the first inhibitor of a MART, representing a previously unexplored therapeutic target.These observations stand in contrast to our results with ovarian cancer cells, in which invasion , as wellFinally, treatment with carboplatin/paclitaxel is the primary initial treatment regimen for the vast majority of ovarian cancers . InteresAB_2665469) (AB_2607074), α-tubulin , β-tubulin , SNRP70 , Flag , HA , rabbit IgG fraction , goat anti-rabbit HRP-conjugated IgG , goat anti-mouse HRP-conjugated IgG , and Alexa Fluor 488 goat anti-mouse IgG .The custom recombinant antibody-like MAR binding reagent (anti-MAR) was generated and purified in-house  . The anthttps://gtexportal.org/home/) project is a resource which allows users to study human gene expression and its relationship to genetic variation. We used GTEx Analysis Release ver. 8 (dbGaP Accession phs000424.v8.p2) to determine the expression of PARP7 in normal human tissues. We selected four tissue types: ovary (n = 180), breast (n = 459), pancreas (n = 328), and kidney cortex (n = 85). The expression values for each tissue, calculated in TPM , were downloaded from the GTEx portal and presented in violin plots showing expression values in log10(TPM+1).The Genotype-Tissue Expression (GTEx)  to generate box plots for comparing expression in the four cancer types across TCGA matched to normal samples from GTEx. The patients in each cohort group were as follows: ovary , breast , pancreas , kidney . The expression data were log2(TPM+1) transformed for differential analysis, which is plotted in our analysis.GEPIA is a portal for analyzing gene expression with a standardized pipeline using RNA-sequencing data from 9736 tumor and 8587 normal samples obtained from TCGA and GTEx . We comphttps://portal.gdc.cancer.gov/legacy-archive/search/f) in the Cancer Genome Atlas (TCGA) (https://confluence.broadinstitute.org/display/GDAC/Home). For mRNA expression analysis in relation to somatic mutations and copy number alterations, bar graphs from the ovarian cancer cohort of TCGA were downloaded from cBioPortal   .OVCAR4, OVCAR3, 293T, and HeLa cells were purchased from the American Type Cell Culture (ATCC). A704 cells were obtained from Dr. Laura Banaszynski. OVCAR4 and OVCAR3 cells were maintained in RPMI  supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin. HeLa, 293T and A704 cells were maintained in DMEM  supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin. Fresh cell stocks were regularly replenished from the original ATCC-verified stocks and confirmed as mycoplasma-free every three months using a commercial testing kit.PARP7 mRNA were as follows:The siRNAs targeting PARP-7 were from Dharmacon (see below) and the control siRNA (SIC001) was from Sigma. The siRNA oligos were transfected into OVCAR4 cells at a final concentration of 30 nM using Lipofectamine RNAiMAX reagent  according to the manufacturer’s instructions. The cells were used for various assays 24 hr after siRNA transfection. The siRNA oligos used to knockdown PARP7 siRNA 1: D-013948–08 (our siRNA #1)PARP7 siRNA 2: D-013948–09 (our siRNA #2)PARP7 siRNA 4: D-013948–07 (our siRNA #3)OVCAR4 cells were cultured as described above before the preparation of whole cell lysates, nuclear extracts, and cytoplasmic extracts.The cells were collected, washed twice with ice-cold PBS, and resuspended in Lysis Buffer  containing 1 mM DTT and 1x complete protease inhibitor cocktail . The cells were vortexed for 30 s in Lysis Buffer and then centrifuged to remove the cell debris. The supernatants were collected, aliquoted, flash frozen in liquid nitrogen, and stored at −80°C until used.2, 3 mM CaCl2, 0.3 M sucrose, with freshly added 1 mM DTT, and 1x protease inhibitor cocktail, 250 nM ADP-HPD, and 10 μM PJ34), incubated on ice for 15 min, and lysed by the addition of 0.6% IGEPAL CA-630 with gentle vortexing. After centrifugation, the nuclei were collected by centrifugation in a microcentrifuge for 1 min at 4°C at 1000 rpm. The supernatant was collected as the cytoplasmic fraction. The pelleted nuclei were resuspended in Nuclear Extraction Buffer  and incubated on ice for 30 min. The nuclear fraction was collected by centrifugation . The nuclear and cytoplasmic extracts were collected, aliquoted, flash frozen in liquid nitrogen, and stored at −80°C until used.The cells were collected, washed twice with ice-cold PBS, and resuspended in Isotonic Buffer . Volumes of lysates containing equal total amounts of protein were boiled at 100°C for 5 min after addition of 1/4 vol of 4x SDS-PAGE Loading Solution , run on 10% polyacrylamide-SDS gels, and transferred to nitrocellulose membranes. After blocking with 5% nonfat milk in TBST, the membranes were incubated with the primary antibodies described above in TBST with 0.02% sodium azide, followed by anti-rabbit HRP-conjugated IgG (1:5000) or anti-mouse HRP-conjugated IgG (1:5000). Immunoblot signals were detected using an ECL detection reagent .PARP7-specific siRNAs at a final concentration of 30 nM in 10 cm diameter culture dishes. Twenty-four hours later, the cells were collected and plated at a density of 20,000 cells per well in a 24-well plate. The cells were fixed with 4% paraformaldehyde at each of the indicated time points and washed with water. The plates were stored at 4°C until the end of the time course when all samples could be analyzed simultaneously. After all samples were collected, the fixed cells were stained with crystal violet  for 30 min with gentle shaking at room temperature. The stained cells were washed several times with water and air dried. The crystal violet in each well was re-dissolved in 10% acetic acid and the absorbance at 570 nm was read using a spectrophotometer. The absorbance of a blank well was subtracted from the samples and the values were normalized to the values at day 1. Three independent experiments were performed with independent biological replicates to ensure reproducibility. Statistical differences were determined using Student’s t-tests at each time point.Cells were transfected with control or PARP7 mRNA. Twenty-four hours later, the cells were trypsinized and were seeded at a density of 100,000 cells into migration chambers  or invasion chambers  following the manufacturer’s protocols. Briefly, the trypsinized cells were resuspended in serum-free RPMI media and collected by centrifugation at 300 x g for 3 min at room temperature. The cell pellets were resuspended in serum-free RPMI media and 500 μL of the media containing 100,000 cells were plated into the top chamber. The chambers were then incubated in 750 μL of RPMI media with 10% FBS. For rescue experiments in OVCAR4 cells expressing wild-type or catalytically dead mutant PARP-7, the cells were plated as above and then treated with 1 µg/mL doxycycline (Dox). After 24 hr of incubation at 37°C, the cells in the top chamber were scraped and removed. The chambers were stained with crystal violet  for 30 min with gentle shaking. The chambers were washed with water, air-dried and the images of cells at the bottom of the membrane were collected using an upright microscope. Three independent biological replicates were performed for each condition. Statistical differences were determined using Student’s t-test.Boyden chamber assays were used to determine the migration and invasive capacity of cells as described below. The cells cultured in six-well plates were transfected with 30 nM control siRNA or two different siRNAs targeting PARP7 or control siRNAs as described above. Total RNA was isolated using the Qiagen RNAeasy Plus Mini kit  according to the manufacturer's protocol. Total RNA was reverse transcribed using oligo(dT) primers and MMLV reverse transcriptase  to generate cDNA. The cDNA samples were subjected to RT-qPCR using gene-specific primers, as described below. Target gene expression was normalized to the expression of RPL19 mRNA. All experiments were repeated a minimum of three times with independent biological samples to ensure reproducibility and a statistical significance of at least p<0.05. Statistical differences between control and experimental samples were determined using Student’s t-test. The primers used for RT-qPCR were as follows:OVCAR4 cells were plated in 6-well plates and transfected with 30 nM of RPL19 forward: 5’- ACATCCACAAGCTGAAGGCA-3’RPL19 reverse: 5’- TGCGTGCTTCCTTGGTCTTA −3’PARP7 forward: 5’- CCAAAACCAGTTTCTTTGGGAG −3’PARP7 reverse: 5’- CAGATTCCATCTACCACATCC −3’Two biological replicates of total RNA were isolated from each siRNA-mediated knockdown. Total RNA was isolated using the RNeasy kit  according to the manufacturer’s instructions. The total RNA was then enriched for polyA+ RNA using Dynabeads Oligo(dT)25 (Invitrogen). The polyA+ RNA was then used to generate strand-specific RNA-seq libraries as described previously . The RNAPARP7 knockdown samples compared to the control samples were integrated to find the commonly regulated gene set.The raw data were subjected to QC analyses using the FastQC tool . The reaPARP7 knockdown. The genes passing the filters were used to make heat maps using Java TreeView  Bioinformatics Resources tool for gene ontology analysis  for geneMultiple sequence alignment of PARP-7 (Q7Z3E1), PARP-1 (P09874), and PARP-12 (Q9H0J9) was performed using Clustal Omega . The catPARP-7 cDNA was then amplified by PCR from the cDNA pools described above, adding sequences encoding an N-terminal Flag epitope to the cDNAs using the primers below.cDNA pools from OVCAR4 cells were prepared by extraction of total RNA using Trizol reagent , followed by reverse transcription using superscript III reverse transcriptase  and an oligo(dT) primer according to manufacturer’s instructions. The human PARP-7 cDNA by PCR-based site-directed mutagenesis. The F547A mutant PARP-7 cDNA was subjected to additional site-directed mutagenesis to generate double mutants by altering the codon for S563 to V, T or I individually. The primers for site-mutagenesis are listed below.Base pair alterations changing the codon for phenylalanine 547 (F547) to glycine (G), alanine (A), or valine (V), or the codon for serine 563 (S563) to G, A, V, threonine (T), or isoleucine (I) were introduced into the BamHI and EcoRI, and ligated into BamHI- and EcoRI-digested pFastBac1 plasmid. All constructs were sequenced to ensure the fidelity of the sequences. The recombinant pFastBac1 bacmids were then prepared for transfection into Sf9 cells by transformation into the DH10BAC E. coli strain with subsequent blue/white colony screening using the Bac-to-Bac system (Invitrogen) according to the manufacturer’s instructions.The PCR products were digested using restriction enzymes Primers used for PARP-7 cDNA cloning:PARP-7 forward: 5’-TTAACCGGATCCATGGACTACAAAGACGATGACGACAAGAGCGAAATGGAAACCACCGAACCTGAGCCAG-3’PARP-7 reverse: 5’-CCTTCCGAATTCTTATCAAATGGAAACAGTGTTACTGACTTCTTCATATTGGATAAC-3’Primers used for site-mutagenesis:PARP-7 F547G forward: 5’-CAGGATGTGGTAGATGGAATCTGCAAACACAACGGTGACCCTCGAGTCTGTGGAAAGCATGC-3’PARP-7 F547G reverse: 5’-GCATGCTTTCCACAGACTCGAGGGTCACCGTTGTGTTTGCAGATTCCATCTACCACATCCTG-3’PARP-7 F547A forward: 5’-CAGGATGTGGTAGATGGAATCTGCAAACACAACGCTGACCCTCGAGTCTGTGGAAAGCATGC-3’PARP-7 F547A reverse: 5’-GCATGCTTTCCACAGACTCGAGGGTCAGCGTTGTGTTTGCAGATTCCATCTACCACATCCTG-3’PARP-7 F547V forward: 5’-CAGGATGTGGTAGATGGAATCTGCAAACACAACGTTGACCCTCGAGTCTGTGGAAAGCATGC-3’PARP-7 F547V reverse: 5’-GCATGCTTTCCACAGACTCGAGGGTCAACGTTGTGTTTGCAGATTCCATCTACCACATCCTG-3’PARP-7 S563G forward: 5’-GCTACAATGTTTGGACAAGGCGGTTATTTTGCAAAGAAGGC-3’PARP-7 S563G reverse: 5’-GCCTTCTTTGCAAAATAACCGCCTTGTCCAAACATTGTAGC-3’PARP-7 S563A forward: 5’-GCTACAATGTTTGGACAAGGCGCTTATTTTGCAAAGAAGGC-3’PARP-7 S563A reverse: 5’-GCCTTCTTTGCAAAATAAGCGCCTTGTCCAAACATTGTAGC-3’PARP-7 S563V forward: 5’-GCTACAATGTTTGGACAAGGCGTTTATTTTGCAAAGAAGGCAAGC-3’PARP-7 S563V reverse: 5’-GCTTGCCTTCTTTGCAAAATAAACGCCTTGTCCAAACATTGTAGC-3’PARP-7 S563T forward: 5’-GCTACAATGTTTGGACAAGGCACTTATTTTGCAAAGAAGGCAAGE-3’PARP-7 S563T reverse: 5’-GCTTGCCTTCTTTGCAAAATAAGTGCCTTGTCCAAACATTGTAGC-3’PARP-7 S563I forward: 5’-GCATGCTACAATGTTTGGACAAGGCATTTATTTTGCAAAGAAGGCAAGC-3’PARP-7 S563I reverse: 5’-GCTTGCCTTCTTTGCAAAATAAATGCCTTGTCCAAACATTGTAGCATGC-3’2, and stored at −80°C until used.Sf9 insect cells were cultured in SF-II 900 medium (Invitrogen) and transfected with 1 µg of recombinant bacmids for expressing Flag-tagged wild-type (WT) or analog-sensitive mutant human PARP 7 (asPARP-7) using Cellfectin transfection reagent (Invitrogen) according to the manufacturer’s instructions. After 3 days, the medium was supplemented with 10% FBS, penicillin, and streptomycin, and collected as a baculovirus stock. After multiple rounds of amplification of the stock, the resulting high titer baculovirus was used to infect fresh Sf9 cells to induce PARP-7 protein expression for 2 days. The PARP-7-expressing Sf9 cells were collected by centrifugation, flash frozen in liquid N2, 0.4 mM EDTA, 20% glycerol, 5 mM β-mercaptoethanol, 2x protease inhibitor cocktail (Roche)], and lysed by Dounce homogenization (Wheaton). The lysate was clarified by centrifugation, mixed with an equal volume of Flag Dilution Buffer , sonicated, and then clarified by centrifugation again. The clarified lysate was mixed with anti-Flag M2 agarose resin (Sigma), washed twice with Flag Wash Buffer #1 , twice with Flag PARP Wash Buffer #2 , and twice with Flag PARP Wash Buffer #3 . The Flag-tagged PARP-7 proteins were then eluted from the anti-Flag M2 agarose resin with Flag Wash Buffer #3 containing 0.2 mg/mL 3x Flag peptide (Sigma). The eluted proteins (~0.5 mg/mL) were distributed into single use aliquots, flash frozen in liquid N2, and stored at −80°C until use.For purification of PARP-7 proteins, the Sf9 cell pellets were thawed on ice, resuspended in Flag Lysis Buffer  with 250 µM NAD+ or NAD+ analogs at room temperature for 30 min.One microgram of purified recombinant PARP-7 proteins (wild-type or mutant) were incubated in ADP-ribosylation Reaction Buffer [30 mM HEPES pH 8.0, 5 mM MgClThe autoADPRylation reaction described above was stopped by the addition of one third of a reaction volume of 4x SDS-PAGE Loading Buffer  followed by incubation at 100°C for 10 min. The reactions were then loaded onto a 10% PAGE-SDS gel and transferred to a nitrocellulose membrane. After blocking with 5% nonfat milk in TBST, the membrane was then blotted with antibodies against PARP-7 (Invitrogen) or Flag (Invitrogen), or an ADP-ribose detection reagent , followed by blotting with anti-rabbit HRP-conjugated IgG (1:5000) or anti-mouse HRP-conjugated IgG (1:5000) secondary antibodies. Immunoblot signals were detected using an ECL detection reagent , and quantified using Image Lab 6.0 (Bio-Rad).4, 5 mM sodium ascorbate) in the dark at room temperature for 2 hr. The clicked proteins were collected by methanol:chloroform precipitation with centrifugation, dissolved in 1x SDS Loading Buffer , and incubated at 100°C for 10 min in the dark. The clicked proteins were run on a 10% PAGE-SDS gel in the dark. After washing twice with 10% methanol and twice with MilliQ H2O, the gel was then visualized on a Bio-Rad Pharos FX Plus Molecular Imager  (The autoMARylation reactions were stopped by methanol:chloroform precipitation with subsequent collection of the precipitates by centrifugation. The protein pellets were dissolved and ‘clicked’ to azido-rhodamine (Click Chemistry Tools) in Denaturing CC Buffer  .+ or each NAD+ analog, the signal from a ‘water only control’ run simultaneously with the experimental conditions was subtracted from itself and each PARP-7 automodification signal . The signal from wild-type PARP-7 with NAD+ was then set to 100, The water control had a value of 0 because of the subtraction of the background signal from itself. Any negative values after the water background subtraction (i.e. less than background) were set to zero. The signals from the wild-type or mutant PARP-7 proteins with NAD+ or the NAD+ analogs were then expressed relative to the background-corrected signal from wild-type PARP-7 and NAD+. This yielded values that ranged from 0 to more than 100, with the highest values coming from pairings between PARP-7 mutants and NAD+ analogs that yielded more PARP-7 automodification than wild-type PARP-7 with NAD+.PARP-7 automodification in the in vitro assays was determined by western blotting as described above. Immunoblot signals were detected by ECL and quantified using Image Lab 6.0 (Bio-Rad). For determination of relative PARP-7 activity with NAD+ or 8-Bu(3-yne)T-NAD+ in ADP-ribosylation Reaction Buffer (see above) at room temperature for 30 min. The reactions were stopped by methanol:chloroform precipitation with subsequent collection of the precipitates by centrifugation. The protein pellets were ‘clicked’ to azido-rhodamine (Click Chemistry Tools) in Denaturing CC Buffer, run on an SDS-PAGE gel, and visualized using a Bio-Rad Pharos FX Plus Molecular Imager, as described above.OVCAR4 cytosolic S100 extract was prepared as previously described . One mic+ for 30 min. The reactions were stopped by methanol:chloroform precipitation. The protein pellets were resuspended in 1 mL of Urea Solubilization Buffer  and the insoluble proteins were pelleted by centrifugation for 1 min at maximum speed in a microcentrifuge.Twenty micrograms of analog-sensitive PARP-7 protein (wild-type or asPARP-7 mutant) were incubated sequentially with 1 mg of OVCAR4 cell or HeLa cell cytosolic extract in ADP-ribosylation Reaction Buffer at room temperature for 5 min and then after the addition of 250 μM 8-Bu(3-yne)T-NAD2O and were collected by centrifugation at room temperature for 1 min at 1000 RCF in a microcentrifuge.Proteins soluble in the Urea Solubilization Buffer were combined sequentially in a 2 mL tube with the following: 100 μL azido-agarose Beads (Click Chemistry Tools), 820 μL water, 40 μL of a 50:250 mM CuSO4:THPTA pre-formed catalytic complex, 20 μL of 500 mM aminoguanidine hydrochloride, and 20 μL of 500 mM sodium ascorbate. The reaction was incubated in the dark on a rotating mixer for 18 hr at room temperature. The beads were then collected by centrifugation at room temperature for 1 min at 1,000 RCF in a microcentrifuge and the reaction supernatant was aspirated. The beads were resuspended in 1.8 mL MilliQ HThe beads were then suspended in 1 mL of SDS Wash Buffer  with the addition of freshly made 1 mM DTT. The reaction was then heated to 70°C for 15 min and allowed to cool to room temperature. The reaction was then centrifuged at room temperature for 5 min at 1000 RCF in a microcentrifuge and the supernatant was aspirated. The resin was then resuspended in 1 mL of SDS Wash Buffer containing 40 mM iodoacetamide and incubated at room temperature for 30 min in the dark to alkylate the cysteine residues. The resin was then transferred to a 2 mL single use column (Bio-Rad) and washed as follows in sequential order: 10 washes with 2 mL of SDS Wash Buffer (see above), 10 washes with 2 mL of Urea Wash Buffer , and 10 washes with 2 mL of 20% acetonitrile.2, 10% acetonitrile). Trypsin digestion of bead-bound 8-Bu(3-yne)T-ADP-ribosylated proteins was performed by adding of 1 μg of trypsin (Promega) to the Trypsin Digestion Buffer, with incubation at room temperature overnight on a rotating mixer. The peptides from the tryptic digest were collected and lyophilized for storage at −80°C prior to LC-MS/MS analysis.The resin was resuspended in 500 μL of Trypsin Digestion Buffer T-ADP-ribosylation sites, the beads were transferred to a fresh 2 mL single use column (Bio-Rad) and washed as follows: 10 washes of 2 mL each with SDS Wash Buffer, 10 washes of 2 mL each with Urea Wash Buffer, 10 washes of 2 mL each with 20% acetonitrile, and 5 washes of 2 mL each with Peptide Elution Buffer . The beads were transferred to a microcentrifuge tube and hydroxylamine (Sigma) was added to 0.5 M to elute the glutamate- and aspartate-modified 8-Bu(3-yne)T-ADP-ribosylated peptides from the beads. The eluted peptides were prepared for LC-MS/MS by desalting on a C18 stage tip (Thermo) according to the manufacturer’s protocol and then lyophilized for storage at −80°C prior to LC-MS/MS analysis.Following solid-phase extraction cleanup with an Oasis HLB microelution plate (Waters), the trypsin samples were reconstituted in 2% (v/v) acetonitrile (ACN) and 0.1% trifluoroacetic acid in water such that the resulting concentration was 1 µg/µL. Hydroxylamine-eluted samples were reconstituted in 6 µL of the same buffer. One microliter of the trypsin digest samples were injected onto an Orbitrap Fusion Lumos mass spectrometer (Thermo Electron) coupled to an Ultimate 3000 RSLC-Nano liquid chromatography system (Dionex). Samples were injected onto a 75 μm i.d., 50 cm long EasySpray column (Thermo) and eluted with a gradient from 1–28% buffer B over 60 min. Buffer A contained 2% (v/v) ACN and 0.1% formic acid in water, and Buffer B contained 80% (v/v) ACN, 10% (v/v) trifluoroethanol, and 0.1% formic acid in water.The mass spectrometer was operated in positive ion mode with a source voltage of 1.5–2.4 kV and an ion transfer tube temperature of 275°C. MS scans were acquired at a resolution of 120,000 in the Orbitrap and up to 10 MS/MS spectra were obtained in the ion trap for each full spectrum acquired using higher-energy collisional dissociation (HCD) for ions with charges 2–7. Dynamic exclusion was set for 25 s after an ion was selected for fragmentation. Raw MS data files were analyzed using Proteome Discoverer v2.2 (Thermo), with peptide identification performed using Sequest HT searching against the Human database from UniProt. Fragment and precursor tolerances of 10 ppm and 0.6 Da were specified, and three missed cleavages were allowed. Carbamidomethylation of cysteine was set as a fixed modification, with oxidation of Met and hydroxamic acid addition (+15.0109 Da) to Asp and Glu set as variable modifications. The false-discovery rate (FDR) cutoff was 1% for all proteins and peptides.The ADPRylation site calls from the mass spectra for α-tubulin see  were manThe peptides identified from the mass spectrometry experiments were used for further analyses and the data were expressed in various formats, as described below.The peptide identifications (IDs) in each of the replicates for the two cell lines (OVCAR4 and HeLa) were used to generate a Venn diagram of overlapping peptide IDs using jvenn software . The uniGene ontology analyses were performed using the DAVID online bioinformatics resource  for the Immunoprecipitation (IP) of MARylated proteins from OVCAR4 cells was performed as follows. OVCAR4 cells plated on 15 cm diameter cell culture dishes were transfected with 30 nM control siRNA or two different siRNAs targeting PARP-7. Two 15 cm dishes of cells were used for each siRNA transfection. Forty-eight hours after transfection, the cells were collected, washed twice with ice-cold PBS, and resuspended in IP Lysis Buffer , and incubated at 4°C for 30 min with gentle shaking. The cell debris was cleared by centrifugation for 15 min at 4°C at 15,000 g. The supernatants were collected and the protein concentrations were measured using a Bradford assays.Aliquots of cell lysates containing equal amount of protein were used for each IP condition. Five percent of each cell lysate was saved for input. The cell lysates for IP were incubated with 3 µg of MAR detection reagent or rabbit IgG , and protein A agarose beads overnight at 4°C with gentle rotation. The beads were then washed five times with Lysis Buffer for 5 min each at 4°C with gentle mixing. The beads were then heated to 100°C for 10 min in 1x SDS-PAGE loading buffer to release the bound proteins. The immunoprecipitated material was subjected to western blotting as described above.The plasmid for expression of C-terminal HA-tagged α-tubulin was obtained from Sino Biologicals (HG14201-CY). Mutations of the sites of MARylation  were introduced into the pCMV3-C-HA-TUBA3C plasmid using a protocol adapted from the Quickchange site-directed mutagenesis kit (Agilent). D69N and E71Q were added first, then E77Q to the double mutant to generate the triple mutant using the primers listed below.Primers used for generatingα-tubulin MARylation site mutants:α-tubulin D69N/E71Q forward: 5’- GAGCAGTGTTTGTGAACCTGCAGCCCACTG −3’α-tubulin D69N/E71Q reverse: 5’- CAGTGGGCTGCAGGTTCACAAACACTGCTC −3’α-tubulin D69N/E71Q/E77Q forward: 5’- GTGAACCTGCAGCCCACTGTGGTCGATCAAGTG −3’α-tubulin D69N/E71Q/E77Q reverse: 5’- CACTTGATCGACCACAGTGGGCTGCAGGTTCAC −3’Immunoprecipitation (IP) of HA-tagged α-tubulin from OVCAR4 or 293 T cells was performed as follows:For Nocodazole or cold treatments, OVCAR4 cells were transiently transfected with an expression vector for HA-tagged α-tubulin. Forty-eight hours after transfection, the cells were either placed on ice for 45 min or treated with 6 μM nocodazole for 1 hr and then washed with warm medium three times. The cells were allowed to recover by incubation in normal growth medium at 37°C for 15 min, and then collected and extracted as described above.For the induction of PARP-7 expression in OVCAR4 cells, cells expressing wild-type or catalytically dead mutant PARP-7 were transiently transfected with HA-tagged α-tubulin and simultaneously treated with 1 µg/mL Dox. For expression of PARP-7 in 293 T cells, the cells were transiently transfected with Flag-tagged PARP-7 and wild-type or mutant α-tubulin constructs. Forty-eight hours after transfection, the cells were collected as described above.Cell lysates for IP were incubated with 1.5 µg of mouse monoclonal antibody against HA-tag and protein G agarose beads overnight at 4°C with gentle rotation. The beads were then washed three times with Lysis Buffer for 5 min each at 4°C with gentle mixing. The beads were then heated to 100°C for 5 min in 1x SDS-PAGE loading buffer to release the bound proteins. The immunoprecipitated material was subjected to western blotting as described above.Parp7 cDNA was prepared by extracting total RNA from 3T3-L1 cells using Trizol , followed by reverse transcription using Superscript III reverse transcriptase  and an oligo(dT) primer, according to the manufacturer’s instructions. The Parp7 cDNA was then amplified from the cDNA library with primers listed below and cloned into pCDNA3 using the primers listed below. A cDNA for the catalytically dead PARP-7 mutant (Y564A) was generated by site-directed mutagenesis using Pfu Turbo DNA polymerase  using the primers listed below. The PCR products were then amplified with primers encoding an N-terminal FLAG epitope tag and cloned into the pINDUCER20 lentiviral Dox-inducible expression vector .The Primers used for amplifying Parp7 cDNA:PARP-7 forward: 5’- ATGGATTATAAGGATGACGATGACAAGATGGAAGTGGAAACCACTG −3’PARP-7 reverse: 5’- GTTTAATTAATCATTACTACTTAAATGGAAACAGTGTTACTG −3’Primers used for cloning Parp7 cDNA into pCDNA3 vector:PARP-7 forward: 5’- CTGGCTAGCGTTTAAACTTAATGGATTATAAGGATGACG −3’PARP-7 reverse: 5’- AGCGGGTTTAAACGGGCCCTTTAAATGGAAACAGTGTTACTG −3’Primers used for cloning Parp7 cDNA into pINDUCER20 vector:PARP-7 forward: 5’- TCCGCGGCCCCGAACTAGTGATGGATTATAAGGATGACG −3’PARP-7 reverse: 5’- GTTTAATTAATCATTACTACTTAAATGGAAACAGTGTTACTG −3’Primers used for generating Parp7 catalytic dead mutant, Y564A:Parp7 Y564A forward: 5’- AGCTTGCCTTCTTTGCGAAAGCACTGCCTTGTCCAAACATTG −3’Parp7 Y564A reverse: 5’- CAATGTTTGGACAAGGCAGTGCTTTCGCAAAGAAGGCAAGCT −3’Cells were transduced with lentiviruses for Dox-inducible ectopic expression of PARP-7 (wild-type and mutant). We generated lentiviruses by transfection of the pINDUCER20 constructs described above, together with: (i) an expression vector for the VSV-G envelope protein , (ii) an expression vector for GAG-Pol-Rev , and (iii) a vector to aid with translation initiation  into 293 T cells using Lipofectamine 3000 reagent  according to the manufacturer’s instructions. The resulting viruses were collected in the culture medium, concentrated by using a Lenti-X concentrator , and used to infect OVCAR4 cells. Stably transduced cells were selected with G418 sulfate . The cells were treated with 1 µg/mL Dox for 24 hr to induce protein expression. Inducible ectopic expression of the cognate proteins was confirmed by western blotting.OVCAR4 cells grown in six-well plates were transfected with 30 nM control siRNA or two different siRNAs targeting PARP-7. Twenty-four hours after transfection, the cells were trypsinized and seeded into eight-well chambered slides . For rescue experiments in OVCAR4 cells expressing wild-type or catalytically dead mutant PARP-7, the cells were plated as above and then treated with 1 µg/mL Dox. Twenty-four hours later, the cells were either placed on ice for 45 min or treated with 6 μM nocodazole for 1 hr and then washed with warm medium. The cells were allowed to recover by incubation in normal growth medium at 37°C for 15 min and were then washed twice with PBS, fixed with 4% paraformaldehyde for 15 min at room temperature, and washed three times with PBS. The cells were permeabilized for 5 min using Permeabilization Buffer (PBS containing 0.01% Triton X-100), washed three times with PBS, and incubated for 1 hr at room temperature in Blocking Solution . The fixed cells were incubated with α-tubulin antibody  or HA antibody  in PBS overnight at 4°C. After washing three times with PBS, the cells were incubated with Alexa Fluor 488 goat anti-mouse IgG  in PBS for 1 hr at room temperature. After washing three times with PBS, coverslips were mounted with the VectaShield Antifade Mounting Medium with DAPI . All images were acquired using an inverted Zeiss LSM 780 confocal microscope and the fluorescence intensities were measured using Fiji ImageJ software. In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.Acceptance summary:Using chemical genetics approaches, this study along with the back-to-back study from the Cohen group represent the first systematic investigations on PARP-7 regulated proteome. Particularly, this study not only identifies PARP-7 substrates but also dissects the biological roles of α-tubulin ADP-ribosylation and PARP-7 catalytic activity in cancer phenotypes. Intriguingly, this study identifies part of the PARP-7 specificity being towards the acidic residues, in contrast to the cysetine specificity in the Cohen study. Given the recent findings on the role of PARP-7 in viral infection and the development of PARP-7 inhibitors for cancer therapy, this work is timely for the field and beyond.Decision letter after peer review:eLife. Your article has been reviewed by three peer reviewers, including Anthony K L Leung as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by Philip Cole as the Senior Editor. The following individual involved in review of your submission have agreed to reveal their identity: and Ivan Matic (Reviewer #2).Thank you for submitting your article "Identification of PARP-7 substrates reveals a role for MARylation in microtubule control in ovarian cancer cells" for consideration by The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.eLife".As the editors have judged that your manuscript is of interest, but as described below that additional experiments are required before it is published, we would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). First, because many researchers have temporarily lost access to the labs, we will give authors as much time as they need to submit revised manuscripts. We are also offering, if you choose, to post the manuscript to bioRxiv  along with this decision letter and a formal designation that the manuscript is "in revision at Summary:Parsons, Gibson, Challa et al. created a new variant of their chemical genetics methods to identify PARP7 targets. The authors identified two PARP7 gatekeeper residues and generated 10 different mutants, which were screened for their catalytic activity in the presence of four different NAD analogs. The authors used S563G mutant with the 8-Bu(3-yne)T-NAD+ analog to identify PARP7 substrates by mass spectrometry, which represents an important resource of PARP7 targets that can be explored in future studies. In addition, the authors reported the mRNA expression of PARP7 is lower in cancer than with normal tissue. Among the identified targets, they focus on α-tubulin, indicating that PARP7 is important for the maintenance of microtubules dynamics, suggesting a possible mechanistic bridge between ovarian cancer growth and PARP7 activity. Given that PARP7 inhibitor is now in Phase I clinical trials for solid tumors, this manuscript represents a significant advance and should be considered for publication with the following improvements:Essential revisions:1) Extend the mechanistic insights on PARP7 and microtubules: "PARP-7 mediated microtubule control may play a role in the regulation of cancer cell growth and motility" This statement is premature without further experimental data. The mechanism of the observed PARP7-dependent change in α-tubulin in two tubulin-sensitized conditions (cold and nocodazole treatment) is unclear. Would the level of α-tubulin and its ADP-ribosylation change at the basal, stressed (cold/nocodazole) and recovered in wild-type and PARP7 knockdown condition? Along the same lines, Figure 7C lacks immunofluorescence data of WT and PARP7 KD cells not treated with cold or nocodazole, which is an important control to aid in interpretation of the results.How does this observed change in α-tubulin status relate to cancer phenotype? It is hard to reconcile the observations that PARP7 depletion stabilizes microtubule structure with the decreased cell growth, migration, and invasion observed upon PARP7 knockdown. Is the ability to quickly polymerize and assemble microtubules important for efficient cell growth and migration in cancer cells? If so, it might indicate that the observed phenotype in this cell line is not dependent upon PARP7 ADP-ribosylation of α-tubulin, and may be partially hampered by it. Providing in the text a more thorough exposition of the consequences of α-tubulin dynamics and dysregulation in ovarian cancer may help resolve this apparent contradiction, especially for readers not familiar with the field.Lastly, without identifying α-tubulin residues that are MARylated by PARP7 and showing that their mutation has an impact on miroctubule stability, the authors cannot claim, at this stage, a direct link between PARP7 and microtubules. Therefore, the authors may need to limit their claims in the Title, Abstract and Discussion.2) Is the observed effect due to PARP7 catalytic activity? Although PARP7 knockdown resulted in less MARylation on α-tubulin, it is unclear whether the effect is direct or indirect. Can the author rescue the ADP-ribosylation of α-tubulin by wild-type PARP7 but not catalytically inactive PARP7 in the PARP7 knockdown cells? Would the effects identified in α-tubulin be reproducible using the PARP7-specific inhibitor such as RBN-2397? Would the inhibitor reduce proliferation in ovarian cancer cells through cell growth and motility?3) PARP7 expression in ovarian cancer cell and oncogenesis: Figure 1 legend states that expression levels across tissues were normalized under B) but not A). Could the authors describe how the data were normalized under B) and could they also show normalized data under A). Would the use of GEPIA as mentioned in the Materials and methods be sufficient in this normalization? Do the authors have data to support the case? TPMs presume that the total RNA content between the tissues is the same. This is, however, often not the case. Therefore, it would be good to normalize the data using housekeeping genes. The following article provides a list of genes that have been found to have stable expression over human tissues: https://www.cell.com/trends/genetics/pdf/S0168-9525(13)00089-9.pdfGene expression levels in an ovarian cancer cell line such as OVCAR4 can be very different compared to ovarian cancer tissue. Therefore, PARP7 expression levels should be compared between non-ovarian versus ovarian cells and ovarian normal vs ovarian cancer cells. As mentioned in the Discussion, it is puzzling how PARP7 expression levels are reduced in ovarian cancer tissue, which would suggest a tumor suppressor role for PARP7, and yet PARP7 silencing reduces proliferation and migration properties of OVCAR4 cells as reported in this manuscript, suggesting that it acts as an oncogene. This conundrum could be due to different PARP7 expression levels in ovarian tissue versus ovarian cell lines or specifically the ovarian cancer cell line OVCAR4 used in this study. One issue with the current analysis is that the authors are not comparing PARP7 expression in normal vs ovarian cancer cells. Instead they are comparing tissues, which consists of different cell types . In this case, the authors may want to check the expression of PARP7 in individual ovarian cell types from single cell data:Healthy: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC7052271/Ovarian cancer: https://www.nature.com/articles/s41591-020-0926-0If properly analyzed, these data should elucidate how PARP7 behaves during carcinogenesis.4) Would the observed effect be restricted to OVCAR4 ? The reported PARP7-dependent ADPr of α-tubulin, and its role as mediator of microtubule dynamics in ovarian cancer cells is intriguing, but it is not clear if this role is specific for ovarian cancer cell lines or a general property of PARP7-expressing cells. For successful translation of these results into a clinically-relevant settings, an important step is to clarify this point by comparing the α-tubulin behaviour of OVCAR4 cell line to that of other cell lines. Similarly, the authors show that cell growth of OVCAR4 with PARP7 knockdown is impaired compared to WT, but how does PARP7 knockdown affect other highly-proliferative cell lines? Essential revisions:1) Extend the mechanistic insights on PARP7 and microtubules: "PARP-7 mediated microtubule control may play a role in the regulation of cancer cell growth and motility" This statement is premature without further experimental data. The mechanism of the observed PARP7-dependent change in α-tubulin in two tubulin-sensitized conditions (cold and nocodazole treatment) is unclear. Would the level of α-tubulin and its ADP-ribosylation change at the basal, stressed (cold/nocodazole) and recovered in wild-type and PARP7 knockdown condition? Along the same lines, Figure 7C lacks immunofluorescence data of WT and PARP7 KD cells not treated with cold or nocodazole, which is an important control to aid in interpretation of the results.We thank the reviewer for giving us an opportunity to address this important point. We have added new data showing that recovery from tubulin sensitizing treatments, such as nocodazole or cold, leads to enhanced expression of PARP-7 and increased MARylation of ⍺-tubulin . Together with our previous observation that PARP-7 regulates ⍺-tubulin MARylation, our results identify a novel regulatory pathway of ⍺-tubulin dynamics in which PARP-7 upregulation enhances ⍺-tubulin MARylation when the cells recover from tubulin stressors.More specifically, to address the reviewer’s concern, we added new experiments examining the ADPRylation of α-tubulin and microtubule stability under an additional set of conditions:1) α-tubulin MARylation under basal conditions and after nocodazole or cold treatment . These results show that the levels of α-tubulin MARylation and PARP-7 increase after treatment with the microtubule depolymerizing agents.PARP7 KD ± complementation with Wt or catalytically dead PARP7 . These results support the conclusion that PARP-7, through its catalytic activity, ADPRylates α-tubulin to destabilize microtubules.2) α-tubulin ADPRylation and microtubule stability under basal conditions and after nocodazole or cold treatment, with or without PARP7 KD cells not treated with cold or nocodazole  to Figure 7C, as requested. Interestingly, these control conditions suggest that PARP-7 catalytic activity is required for the microtubule destabilizing effects of nocodazole or cold treatment.3) We have added immunofluorescence data of Wt and Taken together, our new and previous results show a clear role for PARP-7-mediated α-tubulin ADPRylation in nocodazole- or cold-induced microtubule instability.How does this observed change in α-tubulin status relate to cancer phenotype? It is hard to reconcile the observations that PARP7 depletion stabilizes microtubule structure with the decreased cell growth, migration, and invasion observed upon PARP7 knockdown. Is the ability to quickly polymerize and assemble microtubules important for efficient cell growth and migration in cancer cells? If so, it might indicate that the observed phenotype in this cell line is not dependent upon PARP7 ADP-ribosylation of α-tubulin, and may be partially hampered by it. Providing in the text a more thorough exposition of the consequences of α-tubulin dynamics and dysregulation in ovarian cancer may help resolve this apparent contradiction, especially for readers not familiar with the field.We appreciate the reviewer’s concern and agree that we could have done a better job putting these results in a biological context. As the reviewers noted, at face value, our results suggest that enhanced microtubule stability upon depletion of PARP-7 or loss of its catalytic activity has negative consequences for cancer phenotypes . This suggests that in ovarian cancer cells, PARP-7 acts to MARylate α-tubulin to destabilize microtubules. How does this help cancer cells?One possibility, as the reviewer suggested, is that the ability to quickly depolymerize and disassemble microtubules is important for efficient cell growth and migration. Indeed, there is evidence for this in the literature. In fact, there is a class of anticancer drugs that stabilize microtubules , leading to the arrest of proliferation and mitosis. Moreover, previous studies have shown that well characterized tumor suppressors such as RASSF1A and APC act to stabilize microtubule polymerization (PMIDs: 14603253 and 11237529). We have now addressed more clearly the implications of our results for cancer biology and therapeutics in the Discussion.As described in our response below, our new results with an α-tubulin MARylation site mutant do indeed indicate that the observed phenotype is dependent upon PARP-7 MARylation of α-tubulin.Lastly, without identifying α-tubulin residues that are MARylated by PARP7 and showing that their mutation has an impact on miroctubule stability, the authors cannot claim, at this stage, a direct link between PARP7 and microtubules. Therefore, the authors may need to limit their claims in the Title, Abstract and Discussion.PARP7 knockdown  and expression of a catalytically dead PARP-7 mutant  in the microtubule stability assay . These results show that MARylation of α-tubulin is required for the observed effects of PARP-7.The reviewer raises a good point. To strengthen our conclusions, we have identified the sites of PARP-7-dependent MARylation on α-tubulin, mutated them, and performed functional analyses with the mutant . Our analysis identified three sites of MARylation: D69, E71, E77. We mutated each of the residues to similar, but non-modifiable residues (D à N and E à Q), generated expression vectors, and expressed the wild-type or mutant versions of α-tubulin in OVCAR4 cells. Mutation of these sites inhibited MARylation of α-tubulin . Importantly, expression of the α-tubulin MARylation site mutant phenocopied 2) Is the observed effect due to PARP7 catalytic activity? Although PARP7 knockdown resulted in less MARylation on α-tubulin, it is unclear whether the effect is direct or indirect. Can the author rescue the ADP-ribosylation of α-tubulin by wild-type PARP7 but not catalytically inactive PARP7 in the PARP7 knockdown cells? Would the effects identified in α-tubulin be reproducible using the PARP7-specific inhibitor such as RBN-2397? Would the inhibitor reduce proliferation in ovarian cancer cells through cell growth and motility?Parp7 cDNA to produce an shRNA-resistant mRNA that could be used in knockdown-addback experiments (the PARP7 shRNAs that we used were designed against the human PARP7 mRNA). Expression of the catalytically dead PARP-7 mutant resulted in a loss of α-tubulin MARylation . In addition, expression of the catalytically dead PARP-7 mutant phenocopied PARP7 knockdown and expression of the α-tubulin ADPRylation site mutant in the microtubule stability assay  and the cell migration assay  . These results show that PARP-7 catalytic activity is required for the observed effects of PARP-7. We preferred this approach to the use of a chemical inhibitor of PARP-7, such as RBN2397, to eliminate potential off target effects and uncertainty of proper dosing in OVCAR4 cells.We thank the reviewer for this important comment. We performed additional experiments to test whether PARP-7 catalytic activity is required for the observed phenotype. We used a catalytically dead PARP-7 mutant (Y564A) suggested by the reviewer . We expressed the mutant from an Flag-tagged mouse 3) PARP7 expression in ovarian cancer cell and oncogenesis: Figure 1 legend states that expression levels across tissues were normalized under B) but not A). Could the authors describe how the data were normalized under B) and could they also show normalized data under A). Would the use of GEPIA as mentioned in the Materials and methods be sufficient in this normalization? Do the authors have data to support the case? TPMs presume that the total RNA content between the tissues is the same. This is, however, often not the case. Therefore, it would be good to normalize the data using housekeeping genes. The following article provides a list of genes that have been found to have stable expression over human tissues: https://www.cell.com/trends/genetics/pdf/S0168-9525(13)00089-9.pdfWe appreciate the concerns raised by the reviewer, which we have addressed by clarifying the text and adding new analyses. First, we have edited the text to remove the impression that the analysis in Figure 1A was not normalized; this is not true. The GTEx data in Figure 1A were obtained from the GTEx portal and used as normalized TPM scores as described by Li et al., 2010. We have clarified this in the text. Second, we normalized the GTEx data in Figure 1A to 50 housekeeping genes, as suggested by the reviewer . This normalization did not fundamentally change the results.Gene expression levels in an ovarian cancer cell line such as OVCAR4 can be very different compared to ovarian cancer tissue. Therefore, PARP7 expression levels should be compared between non-ovarian versus ovarian cells and ovarian normal vs ovarian cancer cells. As mentioned in the Discussion, it is puzzling how PARP7 expression levels are reduced in ovarian cancer tissue, which would suggest a tumor suppressor role for PARP7, and yet PARP7 silencing reduces proliferation and migration properties of OVCAR4 cells as reported in this manuscript, suggesting that it acts as an oncogene. This conundrum could be due to different PARP7 expression levels in ovarian tissue versus ovarian cell lines or specifically the ovarian cancer cell line OVCAR4 used in this study. One issue with the current analysis is that the authors are not comparing PARP7 expression in normal vs ovarian cancer cells. Instead they are comparing tissues, which consists of different cell types . In this case, the authors may want to check the expression of PARP7 in individual ovarian cell types from single cell data:Healthy: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC7052271/Ovarian cancer: https://www.nature.com/articles/s41591-020-0926-0If properly analyzed, these data should elucidate how PARP7 behaves during carcinogenesis.PARP7 expression. We used the scRNA-seq data suggested by the reviewer to analyze PARP7 expression in normal and cancerous ovarian tissues at the single-cell level . We observed that the identifiable cell types in normal and cancerous ovarian tissues are quite different. The main cell types identifiable as “ovarian” in normal ovary  are not well represented in ovarian cancer. Thus, as the reviewer noted, it is difficult to compare PARP7 expression in the two tissues. By these metrics, PARP7 expression in the “ovarian” cell types varies quite a bit, and may actually be lower on average than in the “malignant” ovarian cancer cell types . In this regard, PARP7 gene gain and amplification is a common feature of ovarian cancers .The reviewer has raised a very good point. Our new analyses, as well as additional information about the origins of ovarian cancer from the literature, suggest that the mixed cell populations in normal and cancerous ovarian tissues are a complicating factor when analyzing and comparing PARP7 expression in normal and cancerous ovarian tissues may be like comparing apples and oranges: (1) the cellular origins of the tissues are different and (2) the cellular compositions are different. Thus, while the results presented in Figure 1B are correct, the require some context and qualification. We have now provided this through (1) new Figure 1—figure supplement 1A and B, (2) presentation of these data in the appropriate section of the Results, and (3) discussion of the results in the Discussion.Furthermore, when contemplating these results, we were drawn to a growing body of work indicating that ovarian cancers may actually arise from cells in the Fallopian tubes. Thus, attempts to compare 4) Would the observed effect be restricted to OVCAR4 ? The reported PARP7-dependent ADPr of α-tubulin, and its role as mediator of microtubule dynamics in ovarian cancer cells is intriguing, but it is not clear if this role is specific for ovarian cancer cell lines or a general property of PARP7-expressing cells. For successful translation of these results into a clinically-relevant settings, an important step is to clarify this point by comparing the α-tubulin behaviour of OVCAR4 cell line to that of other cell lines. Similarly, the authors show that cell growth of OVCAR4 with PARP7 knockdown is impaired compared to WT, but how does PARP7 knockdown affect other highly-proliferative cell lines?PARP7 knockdown in three additional cancer cell lines: HeLa , OVCAR3 (ovarian cancer), A704 (kidney cancer), whose PARP-7 levels are comparable to OVCAR4 cells. In all three cell lines and all three assays, PARP7 knockdown caused an inhibition of the cancer-related phenotypes  and microtubule dynamics  similar to what we observed in OVCAR4 cells. Thus, the observed effects of PARP-7 on cancer-related endpoints are not restricted to OVCAR4 cells.To address the reviewer’s concern, we performed key cancer phenotypic assays  upon


We produced various binding domains to replace the binding domain of IgG from Escherichia coli and the Fc domain of human IgG from Brevibacillus choshinensis as split-intein fusions. We showed that in vitro protein ligation could produce various Fc-fusions at the N-terminus in vitro from the independently produced domains from different organisms. We thus propose an off-the-shelf approach for the combinatorial production of Fc fusions in vitro with several distinct binding domains, particularly from naturally occurring binding domains. Antiviral lectins from algae are known to inhibit virus entry of HIV and SARS coronavirus. We demonstrated that a lectin could be fused with the Fc-domain in vitro by protein ligation, producing an IgG-like molecule as a “lectibody”. Such an Fc-fusion could be produced in vitro by this approach, which could be an attractive method for developing potential therapeutic agents against rapidly emerging infectious diseases like SARS coronavirus without any genetic fusion and expression optimization.Monoclonal antibodies, engineered antibodies, and antibody fragments have become important biological therapeutic platforms. The IgG format with bivalent binding sites has a modular structure with different biological roles, i.e., effector and binding functions, in different domains. We demonstrated the reconstruction of an IgG-like domain structure in vitro by protein ligation using protein  H, CH1, CH2, and CH3), the light chain has two immunoglobulin domains (VL and CL)  and the activation of complement pathways in immunity. Thus, domain structures of antibodies can be dissected based on their functional roles. Antibodies from camelids (like camel and llama) lack light chains, and are therefore termed as heavy-chain antibodies  is a multidomain protein that consists of four polypeptide chains. Two identical heavy chains and two identical light chains form a Y-shaped molecule connected by disulfide bonds a. While  and CL) a 1]. Va. VaH, CHtibodies b 2]. An. AnH, CHications .HH can be obtained by in vitro selection with directed evolution . Sg  of the naturally split DnaE intein from Synechocystis species, strain PCC6803 (SspDnaEC16)  were constructed in this study (We dissected the IgG format at two different sites within the hinge region a. The hiDnaEC16) b. We choposition . Naturalfragment c 21]. T. TC) of  inteins c 22]. W. WC) of is study b.Escherichia coli -GB1 c 36,37]N . We hypothesized that carbohydrate-binding proteins like lectins could be able to replace the role of the variable domain of VHH as a binding domain in IgG format when fused at the N-terminus of Fc domain  under reducing reaction conditions [E. coli, as previously reported [Scytonema varium as a model system. SVN binds to various viruses, including HIV glycoproteins, SARS coronavirus, and Ebola virus [In this study, we demonstrated the feasibility of producing N-terminal fusion proteins of the Fc domain in vitro by PTS. PTS successfully ligated the Fc domain made in nditions ,42. Redunditions ,42. Thisnditions . We succreported ,28. Howereported ,37. We bla virus ,44. It mla virus ,45,46. Tla virus .Npu/SspDnaE split intein with a cysteine residue at the +1 position, which is required for protein splicing reactions and remains in the ligated product. We replaced the cysteine with a serine residue at the +1 position in one of the IntC-Fc precursors (IntC+1S-Fc). This construct, however, still requires reducing conditions for PTS reaction, despite the lack of cysteine in both the intein and hinge regions. This result suggests that other cysteines, such as the first residue of the intein, which is responsible for the first step of the splicing reaction, still require reducing conditions. However, the produced Fc domain retained the capability of protein A binding, indicating the proper three-dimensional structure of the Fc domain by preserving the protein A binding surface. The structure was preserved presumably because two buried intramolecular disulfide bridges in Ig folds (CH2 and CH3) were not influenced by a reducing agent such as 0.5 mM TCEP , and E. coli Rosetta-gami 2 (DE3) pLacI  and T7 Express  were used for protein expression. MT medium, MT supplemented with 1.6 mM MgCl2, and 2SY media were used for culturing B. choshinensis [E. coli was cultured in Luria-Bertani (LB) medium. B. choshinensis was transformed by electroporation with a protocol adjusted from the previous report [hinensis . E. colis report .E.coli expression vector for H6-SUMO-IntN was constructed by transferring the gene of IntN from pHYDuet36 [BamHI and HindIII restriction sites, resulting in pHYRSF53-36. The gene encoding TRX-H6-SVN was amplified from pET(SVN) by PCR with two oligonucleotides, HK901: 5′-TACACCATGGGAAGCGATAAAATTATTCAC and HK900: 5′-TAGGATCCAC TACCACCTCCCGCAGCCGC GTGACCCGC, followed by cloning into pHYRSF53-36 between NcoI and BamHI sites, resulting in pAJRSF25A bearing the gene of TRX-H6-SVN-IntN (NpuDnaEΔC15).For the N-terminal target proteins, an HYDuet36  into pHYHYDuet36  (addgeneC-Fc fusions in B. choshinensis were cloned as follows. The gene encoding the DnaE C-intein (IntC) from Synechocystis sp. (SspDnaEC16) was constructed as described previously and transferred into the pBAD vector, resulting in pSABAD7PG [H2 and CH3from human immunoglobulin G1 was amplified from the L1-Fc vector (addgene #15124) and cloned into pSABAD7PG, resulting in pGVBAD049. The N-terminal hexahistidine tag followed by a thrombin cleavage site introduced by cloning the gene into pRSF vector with PstI site introduced by PCR using two oligonucleotides of GV065: 5′- CAGGTAGCCATCATCATCATCATCACAGCAGC and GV066: 5′-CAGTATATCTCCTTATTAAAGTTAAACAAAATTATTTCTACAGG, resulting in pGVRSF55-049#2. The lower hinge sequence of ‘PAPELLGG’ was introduced by PCR assembly using the four oligonucleotides GV065, AJ001: 5′-CCAGCACCTGAACTCCTAGGGGGACCGTCAGTCTTCCTC, J002: 5′-CCCTAGGAGTTCAGGTGCTGGGTTAAAACAGTTGGCGGCGATAG, and HK548: 5′-TACAAGCTTATTTACCCGGAGACAGGG, and cloned into pGVRSF55-049#2 and pNCM02 vector using PstI-HindIII restriction sites, resulting in pAJRSF18A and pAJNCM18P, respectively for the expression of IntC-Fc. For the production of IntC+1S-Fc in B. choshinensis, the vector pAJNCM18P was mutated by inverse PCR using a pair of the two primers HK866: 5′- GGTGCTATCGCCGCGAATTCTTTTAACCCAGCACCTG and HK867: 5′-CAGGTGCTGGGTTAAAAGAATTCGCGGCGATAGCACC, resulting in pAJNCM20A. For IntC-CPPC-Fc, the core hinge ‘CPPC’ was introduced into pAJRSF18P by inverse PCR using the two primers HK881: 5′-TTGCCCACCGTGCCCAGCACCTGAGCTCCTAGGGGGAC and HK882: 5′-GGTGCTGGGCACGGTGGGSAATTAAAACAGTTGGCGGCG, resulting in pAJRSF21B. The gene corresponding to IntC-CPPC-Fc was transferred from pAJRSF21B to pNCM02 or pNY326, resulting in pAJNCM21A and pAJNY21F vectors with two different promoters. The pAJNCM18P, pAJNCM20A, pAJNCM21A, pAJNY21F, pHYRSF53-36, and pAJRSF25A will be available from Addgene (www.addgene.org) with addgene #153166, #153167, #153168, #153169, #153170, and #153171, respectively.Expression vectors of IntSABAD7PG . The genB. choshinensis for protein expression and incubated in 4 mL of TM media at 30 °C, 200 min−1 overnight. The precultures were then inoculated by a 100-fold dilution and incubated in 4 mL 2SY medium at 30 °C. After 3–4 days, the culture supernatants were collected and subjected to SDS-PAGE analysis of the protein incubation. The culture supernatants were collected, centrifuged , and loaded onto a 5-mL HisTrap FF column . Unbound contaminants were washed off with 35 mM imidazole, and bound proteins were eluted with a 15- or 20-mL linear gradient of imidazole (35 mM–250 mM) in 50 mM sodium phosphate, 300 mM NaCl, pH 8.0. Fractions were collected and monitored by measuring absorbance at 280 nm (A280), and their purity was confirmed by SDS-PAGE. The fractions containing target proteins were pooled and dialyzed overnight at 4 °C against 20 mM sodium phosphate, pH 6.5 (protein A binding buffer) using 6–8 kDa molecular weight cut-off (MWCO) membrane. The dialyzed samples were concentrated using a centrifugal filter device  and clarified by centrifugation  prior to loading on a 1-mL HiTrap Protein A HP column. After washing, the bound proteins were eluted with a 2-mL linear gradient of 100 mM sodium citrate, pH 3.5 (0–100%). Fractions were collected and monitored by measuring A280 and by SDS-PAGE examination. The fractions containing the target proteins were immediately neutralized with 20% (v/v) 1 M Tris, pH 9.5, pooled, and dialyzed overnight at 4 °C against 10 mM Tris, 500 mM NaCl, 1 mM EDTA, pH 7 (ligation buffer). When necessary, the dialyzed samples were concentrated using a centrifugal filter device .The plasmids pAJNCM18P, pAJNCM20A, and pAJNCM21A were transformed into E. coli Rosetta-gami 2(DE3)pLacI. A single transformant was grown overnight in 50 mL LB medium, at 30–37 °C, 220 min−1. The overnight culture was diluted into 2 L of LB medium in three shake flasks and cultured at 37 °C, 220 min−1. When OD600 reached 0.5–0.6, protein expression was induced with IPTG in the final concentration of 40 μg/mL. After 4-h protein expression, cells were harvested by centrifugation  and resuspended into 50 mM sodium phosphate, 300 mM NaCl, pH 8.0. The cell suspension was flash-frozen with liquid nitrogen and stored at −70 °C. After thawing, cells were lysed by sonication. Cell debris was removed by centrifugation , and the expressed proteins were further purified by polyhistidine tag IMAC with a linear gradient of 50–250 mM imidazole. Elution from the HisTrap FF column was dialyzed overnight at 4 °C against 10–20 mM Tris, 500 mM NaCl, 0–1 mM EDTA, pH 7–8 using 6–8 kDa MWCO membrane.Expression vectors pHYDuet36 , pHYRSF5For ligation reactions, equal amounts of the two precursor proteins were mixed at a final concentration of 10–18 µM. The reactions were conducted using a bench-top shaker  at 25 °C with mild shaking (350 rpm) in either reducing or nonreducing conditions. For reducing conditions, 0.5 mM TCEP and 5 mM DTT were tested. The reactions were monitored periodically after mixing by reducing SDS-PAGE. Each sample for SDS-PAGE was mixed 1:1 with 2× SDS sample buffer containing 200 mM DTT and heated at 95 °C for 5 min to stop the reaction. The samples were either loaded on a gel immediately or stored overnight at −20 °C.trans-splicing reaction conditions on intramolecular disulfide bonds. Proteins AJNCM18P, AJNCM20A, and AJNCM21A at a concentration of 15 µM were incubated in the presence of 0.5 mM TCEP or 5 mM DTT at 25 °C, 350 rpm . Control reactions were incubated without a reducing agent. After 3 h, samples were mixed with the SDS sample buffer and analyzed on a gel. AJNCM21A at a concentration of 14 µM was incubated in the presence of 0.5 mM TCEP for 1.5 h and stored overnight at 4 °C. The buffer was exchanged for a protein A binding buffer in a centrifugal filter device , and the sample was loaded onto the protein A column.SDS-PAGE and protein A affinity chromatography were used to analyze the effect of protein E. coli were ligated in vitro with the C-terminal Fc domain produced in Brevibacillus choshinensis. The in vitro product could constitute an IgG-like Y-shape structure with bivalent binding sites. In nature, there are many natural bioactive proteins, peptides, and domains like lectins that bind to various receptors and ligands but which have not been widely used as a binding domain in IgG-like format. The production of fusion proteins with such bioactive domains in vitro using an off-the-shelf approach paves a new way for the rapid production of IgG-like molecules with known therapeutic potential without any time-consuming genetic engineering and optimization. The off-the-shelf approach using premade domains could be advantageous for drug development against rapidly emerging infectious diseases.We demonstrated an off-the-shelf approach to producing Fc fusion proteins at their N-terminus by in vitro protein ligation using PTS. Independently prepared N-terminal domains made in


This review covers the main synthetic routes to and the corresponding mechanisms of phosphoramidate formation. The synthetic routes can be separated into six categories: salt elimination, oxidative cross-coupling, azide, reduction, hydrophosphinylation, and phosphoramidate-aldehyde-dienophile (PAD). Examples of some important compounds synthesized through these routes are provided. As an important class of organophosphorus compounds, the applications of phosphoramidate compounds, are also briefly introduced. Phosphoramidates (P-N) are a class of organophosphorus compounds known for the presence of a single covalent bond between the tetracoordinate P(V) atom and N(III) atom. There are generally three types of phosphoroamidates, which are distinguished according to the substitution on the P and N atoms  1]..1].2P(O)NR’2 , is a stable phosphoryl bond (P=O). Both P and N atoms are key physiological elements present in genetic material, energy transfer, enzymes, and other biomolecules and are required for various life processes. As such, molecules containing P-N linkages are found in a large array of biologically active natural products (1–7) (1) (Escherichia coli . F,3. F2P(O1–7) (1)  is an anhia coli . Dinoguncoli . Fdiseases . Lastly,5 and 7)  are nuclectively ,10.Due to the inherent physiochemical properties of the phosphorus atom including its polarizability, multivalency, varying oxidation states, and low coordination number, a diverse range of compounds containing the P-N motifs have been synthesized . These PThis class of organophosphates  has been widely used in pest management for crop protection  19,20,2,220,21. P-Ns have been used as additives in both oil and water-based lubricants . It has m/z range of the spectrum, thus interfering with analyte signals. To tackle this, various approaches such as using additives in the organic matrix, high mass matrix molecules, or matrices based on inorganic compounds have been employed . Fo. Fo42]. employed . Further signals . In addi signals . The enh signals . To this signals .N,O-chelating ligands  and phenyl dichlorophosphate (PdCP). Upon reacting the mixture with gaseous or aqueous NH3 in 1941 or primary amine in 1942, P-N and N-P-N were formed . In 1945, P-Ns were made by Atherton, Openshaw, and Todd through a salt elimination process using disubstituted H-phosphonates (H-P), amines, and chlorinating agents (B) Br catalyst (E) azepine (DBU) as a base in the process, the elimination of large amounts of stoichiometric waste, and comparatively lower yields of products are some of the limitations.A synthetic P-N was fortuitously prepared by Atherton, Openshaw, and Todd in 1945 by the salt elimination method using H-phosphonates (H-P) ((RO)f a base  route B base (B) . The sconitiator  95]. Th. Th2P(O)2N)3PBr2). In the reported method, (Me2N)3PBr2 was prepared in situ in a one-pot two-step synthesis of P-Ns from (EtO)2P(O)OH and amines (With reference to the limitations (such as low yield and the formation of organophosphorus anhydrides side products) in the method reported by Appel and Einig , Zwierzad amines  90]. Th. Th2N)3Pn-Bu4N]Br as a catalyst, Zwierzak and Osowska-Pacewicka synthesized P-Ns from (EtO)2P(O)H (diethyl H-P) and various amines ethyl)amine were explored to demonstrate the broad scope of this synthetic transformation. The P-Ns were isolated in 83–100% yield after recrystallization. In a similar synthesis, using [BnEt3N]Cl as the catalyst instead of [n-Bu4N]Br, Zwierzak accomplished the phosphorylation of amines using dialkyl H-phosphonate (RO)2P(O)H R’’’) were used containing a wide range of aryl-substituents 2Ph and 2,6-(CH3)2Ph). Either column chromatography (on silica) or recrystallization were used to purify the P-Ns to obtain isolated yields in the range of 30–80%.In another salt elimination route, this time using inorganic bases and [s amines a 89]. A. An-Bu4N acidity c 96]. D. Dn-Bu4N2P(O)H; R = alkyl) to (RO)2P(O)Cl  in the presence of readily available inorganic bases, thereby eliminating the use of Et3N. However, this method has limitations including a pre-functionalization step, the production of large amounts of stoichiometric waste, and the use of hazardous halogenating reagents.Inorganic salt elimination methods stimulated an era of catalytic transformations of dialkyl H-P ((RO)2P(O)H  to afford (RO)2P(O)Cl  was possible upon reacting with trichloroisocyanuric acid , which was then treated with an equal mixture of amines, R’R’’NH  and Et3N, to form the P-N in 82–92% yield, which was determined as conversions by 31P NMR spectroscopy. However, the isolation of (RO)2P(O)NR’R”  product gave 79% yield. A similar method, under base-free reaction conditions, was achieved by Kaboudin et al. (2P(O)H; R = Et, i-Pr) and amine . The crude product was purified by column chromatography (on silica) to yield isolated pure products in the range of 22–92%.A one-pot synthesis of P-Ns was reported by Gupta et al. a 97]. A. A97]. An et al. b 98]. T. T2P(O)HdPPA) and an amine in the presence of both a chlorinating agent (Cl3CCN) and a base (Et3N), which was added to sequester the acid formed during the reaction (2P(O)OH; R = OEt, OBu, OBn) in the presence of triphenylphosphine and CCl4 was also used to synthesize P-Ns (Jang and coworkers reported a facile synthetic route to form P-Ns using diphenyl phosphoric acid (reaction c 99]. T. TdPPA) ize P-Ns d 86]. T. TdPPA) 4 and Cl3CCN, generate large stoichiometric wastes, and involve a pre-functionalization step in the synthetic process.P-N synthesis via oxidative cross-coupling using chlorinating agents offers a short reaction time, low temperature synthesis, and a diverse choice of phosphorylating reagents. Nevertheless, the methods use hazardous chlorinating agents such as CCl3; R= Me, Et, i-Pr) in the presence I2 was reported by Chen et al. . . 2) has 2 was used in the presence of H2O2 as a catalyst at 20 °C for the dehydrogenative cross-coupling of diethyl H-P and amine or sulfoximines to form P-Ns (2P(O)H, R = Et, i-Pr, Ph) were used as the substrate along with cyclic aromatic secondary amines , derivatives of heterocyclic amines , aliphatic cyclic secondary amines , tertiary amines , or sulfoximine. According to the article, using other sources of I2 such as KI or N-bromosuccinimide (NBS) for the reaction resulted in decreased yield and trace quantities of the desired products, respectively.Iorm P-Ns c 102]. . 2 was u4 in the synthetic processes. The method also uses inexpensive oxidants without additives and generates water or alkanol as stoichiometric waste. However, the synthetic routes are prone to the formation of many undesired side products, suffer from low yields, and sometimes have limited applicability.Iodine-mediated oxidative cross-coupling methods offer better routes to P-N compounds by eliminating toxic halogenating agents such as CCl2P(O)H  in the presence of copper catalysts to synthesize P-Ns was reported by Hayes and coworkers (12–15) (The aerobic oxidative coupling of amines and disubstituted H-phosphonates (H-P) (RO)oworkers , route a (12–15) .2P(O)H  in the presence of CuBr (5 mol%) to give P-Ns  as well as alkylamines were very poor. The effectiveness of the reaction with diaryl H-phosphonates was not reported.Another copper-catalyzed aerobic oxidative cross-coupling reaction reported by Wang et al. in 2013 involved the coupling of aryl amines (1.5 eq.) and dialkyl H-phosphonates (RO)ive P-Ns , route b3O4@MgO nanoparticles were used as a catalyst to synthesize P-N from primary amines and H-P via the Atherton–Todd coupling reaction with CCl4 H  and amine in the presence of CuCl2 and Cs2CO3 d Cs2CO3 , route dN-acylphosphoramidates (AcP-N) from (RO)2P(O)H  and an amide via an oxidative cross-coupling reaction according to Mizuno et al. (2 gave similar cross-coupling results as Cu(OAc)2, while other Cu catalysts such as CuCl2 and CuSO4 and other transition metal catalysts 2.4H2O, Fe(OAc)2) were not selective or completely ineffective.A Cu(II) catalyst, a base, and air as an oxidant were used to synthesize o et al. , route e2P(O)H  with primary or secondary amines  was reacted with butylamine (13% isolated yield) due to oxidative conversion to phosphonic acid (PP-A). Using this aerobic copper-catalyzed dehydrocoupling method, Zhou et al. synthesized a phosphoryl amino compound (16) und (16) .in situ to an organic azide using a Cu-based catalyst and subsequently used for the synthesis of P-Ns via a coupling reaction, according to Dangroo et al. , etc., n = 1, 2), and air as an oxidant. In addition, this route uses environmentally friendly H-P (RO)2P(O)H  and amines as starting materials in a one-pot single-step process without any need for purification of the starting material, thereby making the process cost- and time-effective. Apart from eliminating hazardous starting materials, the method uses fewer chemical additives and generates mainly water as a stoichiometric waste. Furthermore, the methods offer a possibility of catalyst recycling, which is an important attribute for any green chemical process in terms of mass production. However, a notable limitation of the synthetic route is the formation of various undesired by-products from the reactivities of H-P and amine with the catalysts under oxidative conditions . A. A2P(O)H2P(O)H (H-P) in the presence of lithium iodide-tert-butyl hydroperoxide (LiI/TBHP) was used for the synthesis of P-N and other compounds by Reddy et al. (An oxidative cross-coupling reaction of amines and (EtO)y et al.  111]. T. T2P(O)H2P(O)H  or (RO)2P(O)Cl  and amine as the P- and N-sources, respectively. As a result, derivatives of P-N compounds synthesized through these routes are limited mainly to substituents on the amine starting materials. Due to increased applicability, it has become necessary to synthesize P-N compounds with diverse functionalities. In order to achieve this, nitrene insertion from organic azides is becoming a popular one because the route uses phosphoryl azide, which acts a dual source of P and N. In this case, it is possible to add (or insert) diverse molecules on the P-N moiety through C-N bond formation.Many conventional routes to P-N synthesis as discussed above have used H-P (RO)3, and tert-butyl were some of the substituents on the benzamide. Biologically active phosphoramidates (17 and 18) were synthesized through this method  from aldehydes and phosphoryl azide through nitrene insertion -based catalyst was also used to synthesize P-Ns by Zhu et al. c 114]. . 114]. T2 gas as the only stoichiometric by-product, the process uses expensive catalysts in addition to many chemical additives. Moreover, apart from the hazards associated with the handling of phosphoryl azide, the route requires a long reaction time, high temperature, and does not favor electron-deficient starting materials.Although nitrene insertion provides a viable route to P-N synthesis and generates N3P  for the synthesis of P-N by Sangwan et al. n et al.  The crud19) was synthesized using the proposed method  relative to other routes and generate N2 as stoichiometric waste. However, the method relies on the pre-functionalization and preparation of hazardous azide precursors in multi-step processes.Routes via organic azide generation use readily available, less toxic organic halides and (RO)t-butylamine), substituted primary amines , and heterocyclic amines . Although the method circumvents the use of metal catalysts in the synthesis, the use of a hazardous azide and high temperature are its limitations.Transition metal-free synthesis of P-N from phosphoryl azide and amines was accomplished by Chen et al. . The synN-arylphosphoramidates from nitrobenzene and (RO)3P    118]. T. TN-aryl3P  after Lewis acid-catalyzed rearrangement angement  119]. T. T3P  to achieve the formation of a P-N featuring a P-C-N-P linkage in which the dialkyl phosphite group of (RO)2PONa was added to the carbon and nitrogen atoms of the nitriles  using sodium dialkyl phosphite ((RO)nitriles  120]..2PONa, R6H4CH2, C6H4CH2, C6H4COCH3, 4-BrC6H4, phenyl, and methyl. The unsaturated aldehyde derivatives that were used were 3,7-dimethylocta-2,6-dienal, 3-methylbut-2-enal, pent-2-enal, and hex-2-enal. This newest route for P-N synthesis uses inexpensive catalysts, generates only H2O as a by-product in stoichiometric amounts, and demonstrates an important route to the synthesis of polyfunctionalized P-N compounds.Chabour et al. reported a diastereoselective P-N synthesis via the PAD process using a TsOH acid catalyst and a dienophile, maleimide  1]. Fla. Fla1]. A wealth of information on the mechanisms for the synthetic routes to P-Ns is available. For each of the main categories highlighted above, researchers have provided mechanistic insight into the reaction by determining the main sources of oxidation as well as electronic and steric factors provided from the substituents that influence the reaction rate. Several examples of postulated catalytic cycles will be provided below, illustrating the key steps, similarities, and differences between them.2P(O)H  and CCl4 in the presence of Et3N − . The H-P2 before it abstracts chlorine from CHCl3 to form dACP 3 is oxidized to hexamethyltriaminodibromophosphorane ((Me2N)3PBr2) which is ionized in solution (2N)3PBr2 and diethyl hydrogen phosphate undergo ligand exchange in the presence of Et3N to form the intermediate, diethyl (phosphaneyloxy)phosphonate amino-hydrazine-bromide salt complex. Nucleophilic attack from the amine onto the positively charged P results in the diethyl phosphoramidate bromide salt, which reacts with Et3N to give the desired product with a concomitant elimination of triethylammonium bromide. Apart from the stoichiometric Et3N salt waste generated, the synthetic process is prone to the formation of significant quantities of pyrophosphate or phosphanetetraamine bromide salt side products, resulting in a low yield.In the plausible mechanism for the direct conversion of diethyl hydrogen phosphate, hexamethyltriaminophosphine P(NMesolution  90]. Su. Su2)3 i3, K2CO3, and [n-Bu4N]Br as outlined in the paper and a recently proposed mechanism for an Atherton–Todd type reaction, a probable mechanism can be postulated 2P(O)H, R = alkyl) is deprotonated by the cationic Cl3C+ to yield CHCl3 and anionic phosphate, (RO)2P(O)−. Next, the anionic phosphate reacts with HCO3ClH− to form (RO)2P(O)Cl  and H2CO3. In the following step, a nucleophilic substitution between the amine and dACP results in the formation of the desired product. Meanwhile, K2CO3 serves as the dehydrating agent in the reacting system. The plausible mechanism is also similar when NaOH and [BnEt3N]Cl are used in place of KHCO3 and [n-Bu4N]Br, respectively.The mechanistic steps for the synthesis of P-N via a salt elimination route were not proposed by Zwierzak et al. ; howeverstulated  121]. I. I3, K2C2P(O)H, R = alkyl) to form (RO)2P(O)Cl  . T. T103]. 2) and other Cu catalysts 2·H2O) were inactive or inefficient for the catalysis. It was further suggested that although a CuCl catalyst enabled reaction, it was not as effective as CuBr and conducting the reaction under anaerobic conditions  did not lead to the desired product.There were no suggested mechanisms for the reaction in the report by Wang et al. . However3O4@MgO nanoparticle-mediated synthesis of P-Ns as per the procedure reported by Habibi et al. . . 2P(O)Cl another ,123.2CO3 first deprotonates the amide, affording unpaired electrons on a Cu-amide complex . S. S109]. 2, the anionic dye radical is reoxidized to the organic dye, while the anionic superoxide that is formed in the process reacts with diethyl H-phosphonate (H-P) to generate anionic hydrogen peroxide and a diethyl phosphoryl radical. Then, the radical reacts with the cationic-amine radical to form a protonated P-N, which is then deprotonated by the anionic hydrogen peroxide. The authors reported that after an initial 74% isolated product, the organic dye catalyst was recycled a second time using an acid–base extraction, and 67% isolated product was obtained the second time.In a suggested mechanism for the organophotocatalytic synthesis of P-N, a photon from the visible light source is absorbed by the organic dye to become an excited radical  110]. B. B110]. 2O, and the desired product is formed in the final step from the reaction of phosphoryl iodide and lithium amide.In the overall mechanism suggested for the alkali–metal catalyzed oxidative cross-coupling reaction, a phosphoryl iodide intermediate and LiOH are generated in the first step from a reaction of LiI and the H-phosphonate (H-P) in the presence of aq. TBHP  111]. T. T111]. (III)-based catalyst to generate an iridacyclic intermediate, which then reacts with the phosphoryl azide to produce a coordinated phosphoryl azide–iridacyclic adduct -based catalyst affords the desired product while regenerating the catalyst.The mechanism of the reaction of nitrene insertion according to Chang, Kim, and coworkers follows activation of the C-H bond cleavage on the amide or ketone substrate by the Irc adduct  112]. B. B(III)-(III)-based catalyst in the presence of AgOAc and AgSbF6, which abstracts an H atom from the 2-phenylpyridine derivative to cyclo-(2-phenylpyridine)iridium(III) complex . I. I3P , eliminating N2, in the first step amino)trialkoxyphosphonium complex, cleaving off BF3. Nucleophilic attack on the trialkoxyphosphonium complex from a neighboring phosphorimidate completes the catalytic cycle, affording a P=O bond of the desired P-N product.The plausible two-step mechanistic process suggested by Hackenberger’s group for the synthesis involves the formation of phosphorimidate from the reduction of azide in the presence of (RO)rst step  119]. I. I3P  reacts with alkyl nitrile to form an imine (C=N), which reacts further with a second equivalent of (RO)2PONa to form the 1,1-addition product, sodium alkyl)amide yl)amide  120]. I. I2PONa,2P(O)NH2 reacts with (or is activated by) acetic anhydride to form diethyl acetylphosphoramidate and acetic acid phosphoramidate. The subsequent direct addition of the maleimide derivative results in a P-N adduct intermediate, which is deacetylated in the presence of acetic acid to afford the desired product.Chabour et al. proposed a mechanism of the reaction in which (EtO)tic acid  1]. Mea. Mea2P for more sustainable, efficient, atom economic, selective, and milder synthetic procedures toward P-Ns. In addition, the discussed mechanisms require a deeper understanding of the associated phosphorus chemistry in order to have better synthetic control and to optimize the yield and selectivity toward P-N products. To overcome these challenges, it is predicted that better catalysts, improved reaction conditions, and new solvent solutions for separating the product from the by-products and potentially improving selectivity are in the forecast. For example, there is a sharp rise into synthetic research under solvent-free conditions or using ionic liquids (ILs) and deep eutectic solvents (DES). These can be environmentally benign, highly versatile, selective, and recoverable solvents for use in catalysis.