Trypanosoma brucei, most mitochondrial mRNAs undergo editing, and 3′ adenylation and uridylation. The internal sequence changes and terminal extensions are coordinated: pre-editing addition of the short (A) tail protects the edited transcript against 3′-5′ degradation, while post-editing A/U-tailing renders mRNA competent for translation. Participation of a poly(A) binding protein (PABP) in coupling of editing and 3′ modification processes has been inferred, but its identity and mechanism of action remained elusive. We report identification of KPAF4, a pentatricopeptide repeat-containing PABP which sequesters the A-tail and impedes mRNA degradation. Conversely, KPAF4 inhibits uridylation of A-tailed transcripts and, therefore, premature A/U-tailing of partially-edited mRNAs. This quality check point likely prevents translation of incompletely edited mRNAs. We also find that RNA editing substrate binding complex (RESC) mediates the interaction between the 5′ end-bound pyrophosphohydrolase MERS1 and 3′ end-associated KPAF4 to enable mRNA circularization. This event appears to be critical for edited mRNA stability.In  Trypanosoma brucei, but the involved poly(A) binding protein is unknown. Here, Mesitov et al. show that a pentatricopeptide repeat factor KPAF4 binds to A-tail and prevents exonucleolytic degradation as well as translation of incompletely edited mRNAs.Polyadenylation stabilizes edited mitochondrial mRNAs in  Trypanosoma brucei (T. brucei) maintains a mitochondrial genome composed of catenated maxicircles and minicircles. A few 23-kb maxicircles encode 9S and 12S rRNAs, six protein-coding and 12 encrypted genes, a trans-acting MURF2-II and cis-acting CO2 guide RNAs (gRNA). Thousands of 1-kb minicircles produce gRNAs that direct U-insertion/deletion editing of cryptic maxicircle transcripts, thus giving rise to open reading frames3. Messenger and rRNA precursors are individually transcribed4 and processed by 3′–5′ exonucleolytic trimming, which is followed by adenylation or uridylation, respectively5. Trimming is accomplished by DSS1 3′–5′ exonuclease6 acting as subunit of the mitochondrial processome (MPsome), which also contains an RNA editing TUTase 1 (RET1) and several structural polypeptides7. Binding of the pentatricopeptide repeat (PPR) (35 amino acids) kinetoplast polyadenylation factor 3 (KPAF3) to purine-rich sequences near the encoded 3′ end recruits KPAP1 poly(A) polymerase and channels pre-mRNA into the adenylation/editing pathway8. Conversely, rRNAs lacking KPAF3-binding sites upstream of the MPsome-generated 3′ end are uridylated by RET1 TUTase5. The U-tails decorating ribosomal9 and guide RNAs10 reflect a mechanism in which antisense transcripts impede 3′–5′ degradation thereby creating a kinetic window for U-tail addition7. Thus, uridylation terminates rRNA and gRNA precursor trimming, but the resultant U-tails do not influence the stability of mature molecules12. In contrast, short A-tails (20–25 nt) exert profound and opposite effects on mRNA decay depending upon the molecule’s editing status. Knockdown of KPAP1 poly(A) polymerase leads to moderate upregulation of non-adenylated pre-edited mRNA but causes a rapid degradation of the same transcript edited beyond the initial few sites13. Remarkably, mRNAs containing functional coding sequence that do not require editing, referred to as unedited, also rely on KPAF3 binding and ensuing KPAP1-catalyzed A-tailing for stabilization. In massively edited (pan-edited) transcripts, sequence changes typically begin near the 3′ end and proceed in the 3′–5′ direction14. An unknown signaling mechanism monitors editing status and triggers short A-tail extension into a long (>200 nt) A/U-heteropolymer upon completion of the editing process at the 5′ region. The A/U-tailing is accomplished by KPAP1 poly(A) polymerase and RET1 TUTase and requires an accessory heterodimer of PPR proteins KPAF1 and KPAF2. The A/U-tail does not affect the stability, but rather activates mRNA for translation by enabling binding to the small ribosomal subunit15. Thus, the temporally separated pre-editing A-tailing and post-editing A/U-tailing processes are distinct in their factor requirements and functions.The hemoflagellate 5. It follows that KPAF3-bound pre-edited mRNA is protected against 3′–5′ degradation and remains stable while losing A-tail upon KPAP1 knockdown8. It has been suggested that KPAF3 displacement by the editing sequence changes would leave the partially edited transcript reliant on the short A-tail as an critical stability determinant5. This model, however, does not explain the resistance of adenylated RNA to either degradation by the MPsome, or uridylation by RET1 in vivo. Indeed, these features would be essential for partially edited mRNA stabilization and for blocking its A/U-tailing, hence premature translational activation. However, synthetic adenylated RNA represents a susceptible substrate for degradation by the MPsome7 and uridylation by RET116 in vitro.Selective KPAF3 binding to G-rich pre-edited, but not to U-rich-edited sequences, likely monitors initiation of mRNA editing at the 3′ end, which rationalizes the editing-dependent stability phenomenon4. The PPsome is comprised of three subunits: MERS1, a NUDIX (nucleoside diphosphates linked to x (any moiety)) hydrolase; MERS2 PPR factor; and MERS3, a subunit lacking any motifs. The PPsome binds the 5′ end of a primary transcript and converts the 5′ triphosphate moiety incorporated at transcription initiation into a monophosphate. Intriguingly, MERS1 knockdown severely compromises edited mRNA stability without affecting 3′ polyadenylation. To reconcile these observations, we hypothesized that poly(A)-binding protein (PABP) may inhibit mRNA 3′–5′ degradation and 3′ uridylation by sequestering the short A-tail. We further reasoned that PABP may interact with the PPsome at the 5′ end to stabilize mRNA during the editing process. Unable to identify a canonical RRM motif-containing PABP in mitochondria, we inquired whether a PPR factor capable of recognizing adenosine stretches may exist. A recognition code developed for PPRs from land plants suggests that each repeat binds a single nucleotide via amino acid situated in positions 5 and 35, or the last residue in helix-turn-helix motifs exceeding the canonical length17. For example, a combination of threonine and asparagine in positions 5 and 35, respectively, recognizes an adenosine base19.Recent identification of the 5′ pyrophosphohydrolase complex (PPsome) introduced another dimension to the mRNA processing and stabilization pathway20, we identified a polypeptide containing five adjacent repeats predicted to bind as many contiguous adenosines. Termed kinetoplast polyadenylation factor 4 (KPAF4), this protein interacts with established components of the polyadenylation and editing complexes and predominantly binds to short A-tails in vivo. KPAF4 knockdown downregulates A-tailed edited and unedited mRNAs, but not their A/U-tailed forms. Remarkably, KPAF4 repression also permits uridylation of A-tailed pre-edited mRNAs. Specific KPAF4 binding to adenylated substrate inhibits both 3′–5′ RNA degradation by the MPsome and uridylation by RET1 TUTase in vitro. Collectively, our data support a model in which KPAF4 binding to the short A-tail and interaction with the PPsome enable intramolecular circularization, thus inhibiting mRNA decay by the MPsome.By searching for repeats with T/N pattern among 38 predicted PPRs17, in annotated PPR-containing polypeptides from T. brucei20. We searched for threonine and asparagine residues in these positions, respectively, a combination that binds an adenosine19. By considering proteins with at least four adjacent repeats, we identified a candidate 31.8 kDa protein termed KPAF4 (Tb927.10.10160), which consists almost entirely of seven PPR repeats. The 6–7 repeat organization is conserved among orthologous proteins in Trypanosoma and Leishmania species, while repeats R1–R5 invariantly possess a T–N combination  form of T. brucei and demonstrated to have been enriched in the mitochondrial matrix by approximately eight-fold. Partial association with the inner membrane has also been detected , RNA editing core (RECC) (REL1/224) and substrate-binding (GRBC1/225) complexes, and KPAF1/2 polyadenylation factor15. Only a trace amount of RET1 TUTase26 was detected in the KPAF4 fraction.To place the candidate protein into a functional context, KPAF4 was isolated by tandem affinity chromatography Fig. . Purificors Fig. . KPAP1, 8, while KPAF4 was separated into particles of ~300 kDa (I) and ~600 kDa (II), and attached to an ~1 MDa complex . Notably, RNase pre-treatment of mitochondrial lysate mostly eliminated the 1 MDa KPAF4 complex III but left smaller particles unaffected. The PPsome and RNA editing substrate binding complex (RESC) co-fractionated as an ~1 MDa particle that closely resembles complex III, while the RECC migrated as a distinct ~800 kDa particle. Collectively, these results demonstrate that KPAF4 is a mitochondrial PPR factor engaged with at least three macromolecular complexes. The largest KPAF4-contanining complex III with an apparent molecular mass of ~1 MDa closely resembles a ribonucleoprotein assembly that encompasses PPsome, RESC, and polyadenylation complexes25.Co-purification with protein complexes responsible for mRNA 5′ end modification, editing, A-tailing, and A/U-tailing indicates that KPAF4 likely participates in mRNA processing, and that some interactions are RNA-dependent. To assess the heterogeneity and apparent molecular mass of KPAF4-containing particle(s) in relation to established mRNA processing complexes, mitochondrial lysates from parental and KPAF4-TAP cells were fractionated on glycerol gradients. Fractions were separated on a native gel and analyzed for polyadenylation, PPsome, RNA editing core (RECC), and substrate-binding (RESC) complexes Fig. . In agre27 were derived from LC–MS/MS analysis of tandem affinity purified complexes and used to build an interaction network  of the RESC25. KPAP1 and KPAF3 also featured prominently among KPAF4-associated proteins. Interestingly, relatively high levels of MRP1 and MRP2 were detected in KPAF4 preparation  polymerase, and KPAF1-2 and KPAF3 polyadenylation factors are sufficiently stable to withstand a two-step purification, but nonetheless depend on an RNA component ork Fig. . Polyade33), which has an estimated 10 nm labeling range34. KPAP1, GRBC2, MERS1, and KPAF4 were conditionally expressed as C-terminal fusions with BirA* biotin ligase and biotinylation was induced for 24 h. Labeled proteins were purified and analyzed by LC–MS/MS  form of RNA Fig.  and trigRNA Fig. . QuantitRNA Fig. . We nextRNA Fig. .Fig. 3KP14. Samples from KPAF3 knockdown were analyzed alongside to typify impeded mRNA adenylation and accelerated decay5. Northern blotting of pre-edited, partially-edited  and fully edited variants also distinguishes non-adenylated, A-tailed, and A/U-tailed mRNAs  stretches  of all unmapped KPAF4-CLAP reads while fragments ending with more than 30 As constituted 33% (1.5 × 106) of all reads mapped to mitochondrial mRNAs. Tail sequencing and KPAF4-CLAP statistics are provided in Supplementary Data To establish KPAF4 in vivo-binding sites, we applied UV-crosslinking of live cells, and two-step affinity purification of TAP-6His-tagged polypeptide followed by deep sequencing -binding specificity is conferred by amino acid residues occupying positions 5 and 35 or the last residue in KPAF4 repeats, we introduced T5N and N35/36D substitutions into all seven PPRs Fig. . The exp5. These observations may provide a mechanistic basis for the rapid decay of edited mRNA in MERS1 knockdown12. MERS1 pyrophosphohydrolase binds to the 5′ terminus and removes pyrophosphate from the first nucleotide incorporated by transcription, but the mechanism of mRNA stabilization by MERS1 remains unclear4. If circularization indeed takes place, we reasoned that MERS1 would also be expected to bind the 3′ end and/or A-tails. Mapping of MERS1-CLAP reads to the same transcripts exposed the KPAF4-like re-distribution of MERS1-binding sites from the 5′ end in pre-edited to both 5′ and 3′ termini including A-tails in edited mRNAs  polymerase activity is intrinsically limited to adding 20–25 adenosines38. In reactions containing a mixture of both enzymes, the extension patterns were dominated by RET1 activity. Uridylated RNA was efficiently utilized by RET1 but proved to be a poor substrate for KPAP1. In contrast to no-tail and U-tailed RNA, KPAP1 inhibited processive uridylation of the A-tailed substrate by RET1 TUTase. Unlike KPAF3, which dramatically stimulates KPAP1 activity on any tested RNA5, KPAF4 did not produce noticeable effects on either RET1 or KPAP1 activities with no-tail or U-tail RNA. However, KPAF4 inhibited processive uridylation of A-tailed RNA by RET1 TUTase, and this effect was further enhanced by KPAP1. Together, these results demonstrate that KPAF4 specifically recognizes adenylated RNAs and inhibits their uridylation by RET1 TUTase. Importantly, KPAF4’s inhibitory effect on uridylation is enhanced by KPAP1 poly(A) polymerase.In an electrophoretic mobility shift assay (EMSA), only adenylated RNA formed a single distinct ribonucleoprotein complex commensurate with increasing KPAF4-WT concentration Fig. . Convers7. While KPAF3 has been shown to protect any RNA against degradation by the MPsome in vitro5, KPAF4-binding properties and knockdown outcomes suggest that it may preferentially inhibit degradation of adenylated RNAs. To test this hypothesis, we reconstituted mRNA degradation with affinity-purified MPsome and the same 5′ radiolabeled substrates used in binding and 3′ extension assays. Reactions were performed for a fixed duration in the presence of increasing KPAF4 concentrations  polymerase is critical for edited and unedited mRNA stability13. Addition of 20–25 adenosines is stimulated by KPAF3 polyadenylation factor, which is recruited to pre-edited mRNA, but is then displaced by editing events5. Thus, transcripts edited beyond a few initial sites depend on the short A-tail for protection against destruction by the mitochondrial processome. Although 3′–5′ degradation is the main decay mechanism, mRNA stabilization also requires binding of PPsome subunit MERS1 to the 5′ end. Finally, post-editing A/U-tailing involving RET1 TUTase activates ribosome recruitment and translation, but this reaction is somehow blocked during the editing process to avoid synthesis of aberrant proteins from mRNA lacking an open-reading frame15. To reconcile these observations, we envisaged that a trans-acting factor may recognize a nascent A-tail to enable an interaction between protein complexes occupying 5′ and 3′ mRNA termini. Consequentially, this would increase resistance to degradation and uridylation. In this study, we identified the PPR-containing KPAF4 as a factor essential for normal parasite growth and demonstrated its role in recognizing 3′ A-tails, preventing mRNA uridylation by RET1, and inhibiting degradation of adenylated mRNAs.Studies of the unicellular parasite 39, each recognizing a single nucleotide via amino acid side chains occupying cardinal positions 5 and 3517. Bioinformatic analysis of trypanosomal PPRs identified KPAF4 as a factor potentially capable of binding five consecutive adenosines, and, therefore, a candidate for a mitochondrial PABP. Biochemical fractionation, immunochemical, and proteomics experiments demonstrate that KPAF4 interacts with polyadenylation and RESC complexes. In agreement with an established architecture of the RESC, KPAF4 contacts are mostly confined to the PAMC, which has been defined as a docking site for the polyadenylation complex25. A binding platform for RNA editing substrates and products40, RESC also recruits enzymatic RNA editing core complex and, importantly for mRNA stabilization, the 5′ end-bound PPsome4. Therefore, it seems plausible that RESC-mediated interaction network provides a physical basis for functional coupling between 5′ pyrophosphate removal by MERS1, KPAP1-catalyzed 3′ adenylation, and internal editing events. To that end, in vivo crosslinking identified 3′ termini and short A-tails as KPAF4 primary recognition sites, but also detected binding to the 5′ region. KPAF4 CLAP-Seq coverage displayed an instructive correlation with the editing status: The 3′ termini including A-tails were occupied in all mRNA types , while the 5′ regions were bound chiefly in pre-edited mRNAs. Remarkably, these patterns were mirrored by editing-dependent re-distribution of MERS1-binding sites. Collectively, interaction networks, proximity studies, and identification of in vivo-binding sites point toward circularization as the mRNA surveillance and stabilization event. In this scenario, only adenylated pre-mRNA proceeds through the editing while being protected by KPAF4-bound short A-tail from the MPsome assault, which degrades RNA5, and from A/U-tailing, which activates translation15.PPR proteins are defined by arrays of ~35-amino acid helix-turn-helix motifs20, but further studies are required to decipher a signaling mechanism. The KPAF4 stabilizing role is somewhat similar to PPR10 in maize chloroplasts, which defines mRNA 3′ end by binding to a specific site and impeding 3′–5′ degradation41. The distinction in lies in post-trimming addition of the KPAP4-binding platform, the A-tail. Conversely, upregulation of some pre-edited  and unedited (ND1) transcripts in KPAF4 knockdown also suggests a more nuanced transcript-specific functions of the PABP. These effects are reminiscent of the moderately destabilizing contribution of the A-tail structure13, an unresolved phenomenon that requires further investigation.Although circularization is likely to take place in vivo, KPAF4 in vitro properties are also consistent with short A-tail-dependent inhibition of 3′–5′ degradation by the MPsome and 3′ uridylation by RET1 TUTase. Accordingly, the outcomes of KPAF4 knockdown revealed specific loss of A-tailed molecules, but minimal impact on post-editing A/U-tailing reaction, which is accomplished by KPAP1, RET1, and KPAF1/2 polyadenylation factors. It seems plausible that the A/U-tailed mRNA no longer depends on KPAF4-mediated stabilization mechanism. The argument can be extended to suggest that completion of editing results in KPAF4 displacement from the short A-tail and/or loss of interaction with the 5′ end. These events would enable RET1 access and trigger A/U-tailing. The presence of a protein sensor that monitors RNA-editing completion has been suggested42. Although the recognition mechanisms are likely to be different, KPAF4 properties are well aligned with a paradigm for PPR repeats as sequence-specific readers and modulators of diverse enzymatic activities. The latter effects can be stimulatory, as typified by KPAF1/215 and KPAF35, or inhibitory, like those conferred by KPAF4.In this example of convergent evolution, a PPR array in KPAF4 apparently carries a similar function to that of an RRM domain, a universal fold of canonical PABPs43. Linearized constructs were transfected into a procyclic 29-13 T. brucei strain22. For inducible protein expression, full-length genes were cloned into pLew-MHTAP vector44. For BioID experiments, full-length genes were cloned into the same vector with the C-terminal TAP tag replaced by a mutated BirA* ligase from E. coli33. DNA oligonucleotides are listed in Supplementary Data Plasmids for RNAi knockdowns were generated by cloning an ~500-bp gene fragment into p2T7-177 vector for tetracycline-inducible expression45. The change in relative abundance was calculated based on qRT-PCR, or northern blotting, data assuming the ratio between analyzed transcripts and control RNAs in mock-induced cells as 1% or 100%, respectively. Detailed protocol for BioID purification from crude mitochondrial fraction is provided in the Supplementary Methods.RNAi, mitochondrial isolation, glycerol gradient, native gel, total RNA isolation, northern, and western blotting, qRT-PCR, and tandem affinity purification were performed as describedm/z 350–1800, resolution of 60,000 at m/z 400) was followed by 10 MS/MS spectra acquired in the LTQ with normalized collision energy (setting of 35%). Following automated data extraction, resultant peak lists for each LC–MS/MS experiment were submitted to Protein Prospector (UCSF) for database searching46. Each project was searched against a normal form concatenated with the random form of the T. brucei database (http://tritrypdb.org/tritrypdb/).Affinity-purified complexes were sequentially digested with LysC peptidase and trypsin. LC–MS/MS was carried out by nanoflow reversed phase liquid chromatography (RPLC)  coupled on-line to a Linear Ion Trap (LTQ)-Orbitrap mass spectrometer (Thermo-Electron Corp). A cycle of full FT scan mass spectrum  to remove linear RNA, and termini were amplified with gene-specific primers listed in Supplementary Data 47.Total RNA (10 μg) was circularized with T4 RNA ligase 145, with modifications outlined in Supplementary Methods.UV-crosslinking, affinity purification, and RNA-Seq library preparation from KPAF4-bound and MERS1-bound RNA fragments have been performed as describedEdited RPS12 mRNA fragments were prepared by in vitro transcription and 5′ radiolabeled.No-tail: GGGTGGTGGTTTTGTTGATTTACCCGGTGTAAAGTATTATACACGTATTGUAAGUUAGAUUUAGAUAUAAGAUAUGUUUUUA-tail: GGGTGGTGGTTTTGTTGATTTACCCGGTGTAAAGTATTATACACGTATTGUAAGUUAGAUUUAGAUAUAAGAUAUGUUUUUAAAAAAAAAAAAAAAAAAAAU-tail: GGGTGGTGGTTTTGTTGATTTACCCGGTGTAAAGTATTATACACGTATTGUAAGUUAGAUUUAGAUAUAAGAUAUGUUUUUUUUUUUUUUUUUUUUUUUUU2, 20,000 cpm of 5′-labeled RNA, 2 μl of TAP-purified DSS1 fraction, and 50 nM of KPAF4. The reaction was pre-incubated at 30 °C for 20 min, and started with the addition of DSS1. Aliquots were separated on 10% polyacrylamide/8 M urea denaturing gel. Phosphor images were acquired with Typhoon FLA 7000 .MPsome assays were carried out in 20 µl reaction containing 50 mM Tris–HCl, pH 8.0, 1 mM DTT, 2 units/μl RNaseOut ribonuclease inhibitor (Life Technologies), 0.1 mM MgClFurther information on experimental design is available in the Supplementary InformationPeer Review FileDescription of Additional Supplementary FilesSupplementary Data 1Supplementary Data 2Supplementary Data 3Supplementary Data 4Supplementary Data 5Supplementary Data 6Source Data FileReporting Summary


Firre and three functionally distinct regions of the lncRNA Xist. incPRINT confirms previously known lncRNA-protein interactions and identifies additional interactions that had evaded detection with other approaches. Importantly, the majority of the incPRINT-defined interactions are specific to individual functional regions of the large Xist transcript. Thus, we present an RNA-centric method that enables reliable identification of RNA-region-specific RBPs and is applicable to any RNA of interest.Regulatory RNAs exert their cellular functions through RNA-binding proteins (RBPs). Identifying RNA-protein interactions is therefore key for a molecular understanding of regulatory RNAs. To date, RNA-bound proteins have been identified primarily through RNA purification followed by mass spectrometry. Here, we develop incPRINT (in cell protein-RNA interaction), a high-throughput method to identify in-cell RNA-protein interactions revealed by quantifiable luminescence. Applying incPRINT to long noncoding RNAs (lncRNAs), we identify RBPs specifically interacting with the lncRNA  RNA-interacting proteome can be identified by RNA affinity purification followed by mass spectrometry. Here the authors developed a different RNA-centric technology that combines high-throughput immunoprecipitation of RNA binding proteins and luciferase-based detection of their interaction with the RNA. There are several established protein-centric approaches such as cross-linking immunoprecipitation (CLIP) methods that reliably and systematically identify all transcripts bound by individual proteins of interest2. By contrast, when the aim is to determine the proteomes bound by individual transcripts, RNA-centric approaches are applied, which are mainly based on affinity capture of an RNA of interest followed by mass-spectrometry (MS) identification of the co-purified proteins3. These powerful methods have led to the identification of a number of functionally relevant RNA–protein interactions. Nevertheless, the unbiased identification of RNA-bound proteomes following probe-based RNA immunoprecipitation often remains challenging due to the generally low efficiency of RNA affinity purifications requiring scaling up of the starting cellular material. When applied to RNAs expressed at the low copy numbers characteristic of many lncRNAs4, generating sufficient input material for RNA affinity capture-MS is difficult and presents a risk of contamination by nonspecific RBPs. In addition, RBPs that are expressed at relatively low levels often do not reach the threshold required for their robust detection by mass spectrometry. Furthermore, nearly all currently available approaches enable the identification of proteins binding the full-length transcript of interest, whereas many lncRNAs have a modular organization with discrete RNA regions performing different functions5. While dChIRP based on RNA affinity capture-MS has allowed validation of already known RNA-region-specific protein interactions6, the de novo identification and assignment of RBPs bound to distinct RNA regions requires dedicated approaches including comparative RNA affinity capture-MS performed on a mutant variant of the RNA of interest7 or precise mapping of the individual RBP binding by protein-centric CLIP methods9.The identification of RNA–protein interactions is essential for deciphering the cellular functions and molecular mechanisms of regulatory RNAs that act through the formation of dynamic ribonucleoprotein (RNP) complexesGiven the growing repertoire of RNAs with important biological functions and the experimental challenges inherent in dissecting their molecular mechanisms of action, complementary strategies could prove helpful for defining the protein interactions of individual RNAs. Here, we present incPRINT, an alternative RNA-centric method to facilitate the in-cell identification and measurement of RNA–protein interactions. One of the distinguishing features of incPRINT is that it does not utilize direct RNA affinity capture, but is rather based on high-throughput immunoprecipitation of thousands of RBPs followed by luciferase-based detection of their interactions with the RNA of interest. Applying incPRINT to well-studied lncRNAs as a proof-of-principle demonstrates that incPRINT enables the identification of proteins interacting with RNAs expressed at low endogenous levels as well as proteins interacting with distinct regions of RNA, facilitating the assignment of RNA-interaction domains. We thus introduce incPRINT as a reliable method to identify in-cell RNA-interacting proteomes.MS2-MS2CP interaction; the RNA-luciferase complex is co-purified with the FLAG-tagged test protein immunoprecipitated from cell lysates by anti-FLAG antibody , ∼1300 transcription factors12, and ∼170 chromatin-associated proteins. The tagged protein content can be adapted to fit the desired experimental setup.To identify RNA–protein interactions systematically in living cells, we developed a method that measures the cellular interactions between any tagged test RNA and any tagged test protein. The principle of incPRINT is the transient dual expression of a MS2-tagged test RNA and a FLAG-tagged test protein in HEK293T cells stably expressing a luciferase detector fused to the MS2 coat protein (MS2CP) from a genomically integrated plasmid Fig. . The tesody Fig. . Indirecody Fig. . To contody Fig. . The incXist called the A repeat, hereafter referred to as Xist(A)13. Because several Xist(A)-protein interactions have been well established7, they served as controls in our initial incPRINT experiments. We engineered a construct to express Xist(A)-MS2 and assayed the ability of Xist(A)-MS2 RNA to interact with a selected set of previously identified Xist-binding proteins15. The non-discriminatory poly(A)-binding protein PABPC3 was used to control for RNA expression and EGFP (Enhanced Green Fluorescent Protein) was used as a negative control. incPRINT luminescence detected specific interactions of Xist(A)-MS2 with SPEN, RBM15, RBM15B, YTHDC1, HNRNPC, SRSF7, and RALY, whereas HNRNPU, reported to bind full-length Xist but not specifically Xist(A)7, and EGFP showed basal binding  was fused with two, four, six, ten or 24 MS2 stem loops, and their interactions with a set of control proteins were tested in a small-scale incPRINT experiment. An increase in the luminescence intensity directly correlated with an increased number of MS2 stem loops up to ten stem loops with no marked increase in binding to the EGFP control -MS2 RNA and FLAG-tagged test proteins were co-transfected in the same cell population as described above. In the second experiment, Xist(A)-MS2 RNA and FLAG-tagged test proteins were transfected separately in two different cell populations and pooled together only after the cell lysis step, permitting the formation of RNA–protein complexes exclusively in vitro -MS2 RNA and the FLAG-tagged proteins were co-transfected , with Xist(A)-MS2. To strengthen the confidence of the incPRINT-identified RNA–protein interactions, all interactions were assayed in biological duplicate, generating two luminescence (RNA–protein interaction intensity) and two ELISA (test protein expression level) values for each tested RNA–protein couple. After filtering out the proteins expressed at insufficient levels (see the ‘Methods’ section), interaction data were analyzed for 2405 proteins. incPRINT’s reproducibility was assessed by calculating the correlation scores of biological duplicates for both the luminescence  and ELISA signals . Notably, no correlation between luminescence and ELISA values was detected, indicating that the interaction intensities were not a mere reflection of the protein expression levels , we assessed its RBP-interactome with incPRINT. The full-length Firre transcript tagged by MS2 was expressed ∼40-fold higher than endogenous FIRRE in the HEK293T cells used for incPRINT  23, and its modular structure and function. The ∼17-kb long Xist transcript contains several conserved sequence regions  that carry out distinct functions during the XCI process, including initiation of gene silencing (the A repeat), maintenance of the X-inactive state (the F- and B-repeats) and proper chromosomal localization and focal accumulation of Xist (the C- and E-repeats)32 , Xist(F), and Xist(C)  to a near-endogenous expression level for Xist(C) , Xist(F), and Xist(C)) was interrogated with our library of ~3000 proteins. To compare signals across individual Xist regions expressed at different levels . Notably, MS2 data were not used as a binding specificity control, as many RBPs recognize low complexity RNA motifs36 present also within the MS2 tag, and since protein binding to MS2 does not exclude a potential functional interaction with a test RNA. Similarly to Firre, we found that the majority of proteins did not bind to any of the tested Xist fragments . For example, incPRINT identified SPEN as a Xist(A)-specific interactor  and Xist(F), but not Xist(C), confirming their reported binding to the 5′ end of Xist9 -specific interaction with HNRNPU  previously shown to be involved in Xist localization33  term enrichment analyses. Consistent with the differential functions reported for the individual Xist regions, Xist(A)- and Xist(F)-associated proteins were enriched for RBPs involved in RNA processing, while the C-repeat region preferentially interacted with DNA-binding proteins involved in transcriptional regulation -interacting proteins were enriched for the SPOC  and RRM protein domains, Xist(F)-interacting proteins were enriched for the RRM domain and Xist(C)-interacting proteins showed no particular enrichment 32 Fig. . Moreove(C) Fig. . When exnts Fig. , whereasion Fig. . Importaons Fig. . Thus, ator Fig. , confirmst9 Fig. . MoreoveXist has a well-characterized cellular function in gene silencing during XCI, we sought to test if some of the Xist-protein interactions uncovered using incPRINT are functionally relevant. The focus was on the ZZZ3 protein, which interacts with all three tested Xist regions, and RBM6, which exhibits more specific binding to the Xist(A) and Xist(F) regions  cells. The proteins were HA-tagged in the polymorphic TX1072 ES cell line that enables doxycycline-induced Xist expression, triggering XCI in the absence of differentiation39. RNA immunoprecipitation (RIP) after doxycycline induction of Xist and UV-cross-linking of cells followed by qRT-PCR analyses identified a significant enrichment of the Xist transcript with RBM6 and ZZZ3 proteins, confirming their interaction in vivo , Lamp2 expression remained unchanged upon depletion of the proteins above  to assess the expression of endogenous 45. To identify the protein interactomes of individual RNAs of interest, we report here incPRINT, a reliable RNA-centric technology which we show uncovers both known and previously unknown RNA–protein interactions. incPRINT successfully overcomes two major obstacles in determining the proteomes of specific RNAs: first, incPRINT enables the identification of proteins associated with RNAs expressed at low endogenous levels, common for many lncRNAs and, second, incPRINT allows the detection of RNA-region-specific protein interactions and the assignment of protein binding to defined regions of a full-length transcript. This is particularly advantageous for defining the RNA-region-specific binding proteome of large modular RNAs with multiple functional regions, such as the ∼17-kb-long Xist transcript. While other RNA-centric methods such as dChIRP have allowed the validation of already known RNA-region-specific protein interactions6, incPRINT enables their de novo identification. Considering this feature of incPRINT, one could envision a higher-resolution mapping of RBP binding along a transcript of interest by applying a low- or medium-throughput-scale incPRINT to shorter RNA fragments or their mutated variants.Several methods have been developed to comprehensively define the RNA-binding protein repertoire of all cellular RNAsXist throughout different stages of XCI7. Finally, incPRINT is flexible in its throughput: we employed here a customized library of ~ 3000 human test proteins comprising together ~1/7 of the human proteome and representing the majority of all known RBPs, transcription factors and chromatin modifiers. Using this protein library, we discovered functional interactors that had evaded detection with other methods; however, the library can conveniently be expanded to include a range of additional proteins, or reduced and customized to fit the experimental design and need.Several key aspects of incPRINT distinguish it from the RNA-centric methods currently employed for the identification of RNA–protein interactions. First, incPRINT does not rely on probe-based capture and direct RNA affinity purification, which generally exhibits a low efficiency and thus requires large amounts of material. Because it uses a sensitive luciferase detector, incPRINT is not limited by RNA low copy number and can be applied to any RNA of interest. Second, the quantitative nature of the luciferase detector used in incPRINT makes it suitable for various custom applications and offers the possibility of structure/function analyses of mutated or disordered RNA–protein interactions and their relative effects on binding affinities. Third, the ability of incPRINT to measure cellular RNA–protein interactions one by one, independently of the cell’s physiological state, is particularly relevant for defining RNA-bound proteomes of transcripts that display a dynamic expression range during development and/or cellular differentiation, as observed with Xist-protein interactions in a heterologous cell line. Because incPRINT is based on ectopic expression of the test protein and RNA components, it is possible that a fraction of the identified interactions will be false positives. While such interactions cannot be excluded, the number of highly specific interactors for each interrogated RNA corresponding to 2–3% of all screened proteins and the validation of a set of interactions by orthogonal methods suggest that the rate of false positive binding identified by incPRINT is rather low. Furthermore, whereas all interactions validated in this study were direct, the design of the method does not make it possible to distinguish between direct and indirect RNA–protein interactions and additional assays are required to stipulate direct RNA binding. In summary, applying the incPRINT method to other noncoding or coding RNAs of various structural complexities will facilitate the discovery of the precise mechanisms by which they exert their cellular functions through the identification of their binding proteins.While incPRINT enables fast, systematic and quantitative identification of RNA–protein interactions, it is a binding assay that does not consider the developmental context or timing of these interactions. However, this potential limitation did not hinder the identification of NanoLuc-MS2CP expression plasmid was transfected using polyethylenimine (PEI) followed by puromycin selection. The expression of NanoLuc-MS2CP was verified by measuring luciferase activity. The cells were maintained in DMEM (Gibco 12007559) containing 10% fetal bovine serum (Gibco 11573397) and 1% penicillin/streptomycin (Gibco 15140122).To generate the stable monoclonal HEK293T cell line  expressing NanoLuc-MS2CP fusion protein, the 37  that harbors a doxycycline-responsive promoter controlling Xist expression on the B6 X chromosome and was derived from a cross of a TX/TXR26rtTA/rtTA female38 with a male Mus musculus castaneus was used. ES cells were cultured in high-glucose DMEM (Sigma) supplemented with 15% fetal calf serum , 0.1 mM β-mercaptoethanol, 1000 U/mL leukemia inhibitory factor , and 2i . Xist expression was induced with doxycycline (Sigma) for 24 h for RNA FISH analysis.The female hybrid mouse TX1072 ES cell line37 was used to generate HA-tagged RBM6 and ZZZ3 cell lines. The HA tag was introduced at the C-terminus of RBM6 and ZZZ3. The tag flanked by two ∼600 nt homology arms was cloned into a pBR322 vector. TX1072 ES cells were transfected using Lipofectamine 2000, with the RBM6 or ZZZ3 HA-targeting vectors together with the pX459 vector containing a gRNA sequence targeting each of the proteins, promoting homologous recombination. Transfected clones were selected with 0.4 μg/mL of puromycin. Clones were genotyped for integration at the correct genomic location by PCR and DNA sequencing to ensure no deletions or mutations in the protein-coding sequence. All relevant oligonucleotide sequences are listed in Supplementary Data The TX1072 ES cell line46 (Eupheria Biotech) directed against Rbm6 and Zzz3, luciferase and Thap7 as negative controls and Spen as a positive control. esiRNA transfections into TX1072 ES cells were conducted using 0.1 μg esiRNAs and Lipofectamine 2000 Transfection Reagent (Thermo Fisher Scientific), in Optimem I reduced serum medium (Thermo Ficher Scientific). Doxycycline-mediated Xist expression was initiated 48 h post-transfection, and cells were harvested 24 h post-doxycycline induction of Xist. The same experimental setup was used for RNA-FISH, RT-qPCR and RIP experiments. To assess esiRNA knockdown efficiencies, 50 ng of total RNA were reverse-transcribed using the SuperScript IV kit (Invitrogen) followed by qPCR using SYBRgreen (Applied biosystems). Arppo mRNA was used to normalize RNA levels between samples. All relevant oligonucleotides are listed in Supplementary Data Gene knockdown was achieved using esiRNAs47 using PstI and BamHI restriction sites. A puromycin resistance gene was added to the plasmid using PvtI restriction site. To remove the FLAG tag present in the original plasmid, a stop codon was introduced between the MS2CP and the FLAG tag by site-directed mutagenesis (Agilent).To generate the luciferase-MS2CP expression vector, NanoLuc luciferase was amplified from the pNL1.1 plasmid (Promega) and cloned into the pCi-MS2 vectorXist(A), (F), and (C) fragments were PCR-amplified from BAC 399K20 , and cloned upstream of the MS2 stem loops by Gibson assembly (New England Biolabs). Primers used for Xist fragments amplification and cloning are indicated in Supplementary Data To generate customized RNA-10xMS2 constructs, the pCDNA3.1 plasmid (Thermo Fisher) was modified. First, additional restriction sites  were incorporated between the KpnI and BamHI sites. Second, MS2 stem loops were inserted between the BamHI and BglII sites. 12. The collection of human RBPs and chromatin-associated proteins were cloned with Gateway recombination from the human ORFeome 5.1 (http://horfdb.dfci.harvard.edu/hv5/index.php) into a mammalian expression vector with a C-terminal 3xFLAG-V5 tag. The expression clones were verified by restriction enzyme digestion.The transcription factor collection has been previously describedFirre-MS2, 30 ng (1:10), and 6 ng (1:50) of the Firre-MS2 encoding vector were used for transfections. Two days after transfection, cells were washed twice with ice-cold 1x PBS and lysed in ice-cold RQ1-HENG buffer  containing 30 U/ml of RQ1 DNase (Promega M6101). After lysis  and DNase incubation , the lysates were transferred to 384-well plates coated with anti-FLAG M2 antibody . Following a three-hour incubation at 4 °C, plates were washed seven times with RQ1-HENG buffer. Furimazine substrate (Promega) was added to the plates and luminescence in each well was measured with a plate reader (Biotek Synergy Neo). Following luminescence measurement, HRP-conjugated anti-FLAG antibody  in ELISA buffer  was added to each well. After 30 min of incubation at room temperature, plates were washed with 1x PBS, 0.05% Tween 20, HRP substrates (Pierce) were added and ELISA signals were detected with the plate reader. For the RNase treatment experiment, RNA–protein interactions were first measured as described above. Fifty microliters of RNase buffer containing 50 mM Tris-HCl pH 8.0, 10 mM EDTA, +/−100 μg/ml RNase A (Qiagen) were added to the samples and incubated for 30 min at 37 °C, followed by three washes with RQ1-HENG buffer, before proceeding with luminescence measurements. All 96- and 384-plate washes were performed with a plate washer (Biotek EL406).384-well plates (Greiner Bio-One 781074) were coated overnight with the anti-FLAG M2 antibody  and blocked for two hours at room temperature in 1% BSA, 5% sucrose, 0.5% Tween 20 in 1x PBS. Plasmids encoding RNA-MS2 and 3xFLAG-tagged test proteins were co-transfected into a 293T stable cell line expressing the NanoLuc luciferase fused to MS2CP. The day before transfection, cells were seeded in 96-well plates . Co-transfections were performed using 12 µg/ml (300 ng of each plasmid per well). To achieve different expression levels of Firre-MS2, Xist(A)-MS2, Xist(F)-MS2, Xist(C)-MS2), all FLAG-tagged proteins were tested in duplicates achieved by independent transfections. After measuring the NanoLuc and ELISA luminescence, log2-transformed ELISA values were binned to assess the distribution of the FLAG-tagged protein expression. Proteins with the lowest ELISA signals were filtered out  were removed from the dataset -MS2, Xist(F)-MS2, and Xist(C)-MS2 interactors, includes all test proteins showing interaction with at least one test RNA, and expressed in all three assays.To compare the interaction intensities among individual A list of the Pfam domains of each protein was downloaded from the Ensembl site. Enrichment of each domain in the set of interactors of a particular RNA vs. the protein library was calculated using a proportion test.Xist expression was induced in undifferentiated ES cells by addition of 1 μg/mL of doxycycline to the culture medium 48 h post-transfection. Twenty-four hours upon Xist induction, cells were harvested using TrypLE Express Enzyme (Thermo Fisher Scientific), washed with 1x PBS, and adsorbed onto Poly-L-Lysine (Sigma)-coated glass coverslips for 10 min. Fixation was performed with 3% paraformaldehyde for 10 min at room temperature, followed by permeabilization in 1x PBS containing 0.5% of Triton X-100 and 2 mM of vanadyl-ribonucleoside complex (New England Biolabs), for 5 min at 4 °C. Coverslips were washed three times in 70% ethanol, and preserved at −20 °C in 70% ethanol. Prior to hybridization, coverslips were dehydrated with increasing concentrations of ethanol , and air-dried. Transcription of the X-linked gene Lamp2 was detected with a BAC spanning its genomic region (RP24-173A8). The Lamp2 probe was labeled with dUTP-Spectrum Green  by nick translation (Abbot). The probes were precipitated with ethanol, resuspended in formamide at 37 °C, denatured at 75 °C for 10 min, and competed with mouse Cot1 DNA (Thermo Fisher) for 1–2 h at 37 °C. Xist was detected with a dUTP-Spectrum Red  nick translation probe from a plasmid spanning its genomic region48. The Xist probe was prepared as described above for the Lamp2 probe, except for the competition step, which was not performed. Probes were mixed and co-hybridized in FISH hybridization buffer , 10 mM Vanadyl-ribonucleoside) overnight at 37 °C. Coverslips were washed 3 × 6 min in 50% formamide in 2x SSC, pH 7.2, at 42 °C, followed by two 5-min washes in 2x SSC at 42 °C. Nuclei were counterstained with 0.2 mg/mL of 4′,6-Diamidine-2′-phenylindole dihydrochloride (DAPI) in 2x SSC for 3 min at room temperature, and mounted onto glass slides using VectaShield mounting medium. Images were acquired using the wide-field DeltaVision Core microscope (Applied Precision) and the inverted confocal Spinning Disk Roper/Nikon-FRAP microscope. 3D image stacks were analyzed with ImageJ.RLuc (as a negative control), Rbm6 and Zzz3. Whole cell extracts were prepared with RIPA buffer  with freshly added 1x protease inhibitor (Roche). Western blot analysis was performed with antibodies detecting RBM6 , ZZZ3  and Tubulin , Merck Millipore; 1:3000). Full blots are provided as a Source Data file.To assess knockdown efficiency at the protein level, TX1072 ES cells were transfected with 2.5 μg of esiRNAs for 2, at 254 nm . Cells were then collected, pelleted and lysed for 10 min on ice in 200 µl of RIPA buffer containing protease inhibitors (Roche) and RNAsin (Invitrogen). Cell lysates were sonicated using a Bioruptor (Diagenode) , and centrifuged for 10 min at 15,000 × g. Supernatant were collected and diluted in RQ1-HENG buffer (to 1.2 ml). Fifty microliters of anti-HA bead slurry  was washed twice with 1 ml of RQ1-HENG buffer and incubated with the cell lysate for 2 h at 4 °C. Following the incubation, beads were washed twice with TNM600 buffer , and twice with TNM100 buffer . Elution was performed in 50 µl Laemmli buffer , RNA was extracted (Trizol-Chloroform) and treated with TURBO DNase (Ambion). Two hundred and fifty nanograms of RNA were reverse-transcribed using the SuperScript IV kit (Invitrogen) followed by qPCR using SYBRgreen (Applied biosystems). Gapdh was used as a reference gene to normalize RNA levels between samples. All relevant oligonucleotides are listed in Supplementary Data RIP experiments were performed using corresponding HA-tagged RBM6 and ZZZ3 TX1072 cell lines. Approximately 15 million cells were treated with doxycycline (1 µg/ml) for 16 h. Cells were washed once with ice-cold 1x PBS and UV-crosslinked using 800 mJ/cmg, 4 °C), washed with 1x PBS, resuspended in 100 μl of buffer A , incubated for 10 min on ice, and centrifuged again . The supernatant was collected (cytosolic fraction), and the pellet was resuspended in 100 μl of buffer B . After sonication  and centrifugation , the supernatant was collected (nuclear fraction). RNA was extracted from both fractions, and treated with Turbo DNase (Ambion). Hundred nanograms of RNA were reverse-transcribed using the SuperScript IV kit (Invitrogen) followed by qPCR analysis using SYBRgreen (Applied Biosystems). β-actin was used as a reference gene. Nuclear/cytoplasmic ratios were calculated for every tested transcript. All relevant oligonucleotides are listed in Supplementary Data Two days after the RNA-MS2 transfections, HEK293T cells were centrifuged (5 min, 100 × Further information on research design is available in the Supplementary InformationReporting SummaryDescription of Additional Supplementary FilesSupplementary Data 1Supplementary Data 2Supplementary Data 3Supplementary Data 4Supplementary Data 5


Bioerosion was estimated at 3.4 ± 0.4 G, resulting in an average net carbonate production rate of 2.5 ± 2.4 G. Comparison of results with a study of the fore-reef slope highlights major differences in post-bleaching carbonate budget state between the fore-reef slope and the reef platform surface. The positive reef flat carbonate budget is attributed to the persistence of massive corals (Porites spp. and Heliopora spp.) through the bleaching event.Coral reefs experienced the third global bleaching event in 2015–2016 due to high sea-surface temperature (SST) anomalies. Declines in net carbonate production associated with coral bleaching are implicated in reef structural collapse and cascading impacts for adjacent coral reef islands. We present the first carbonate budget study of a reef platform surface (reef crest and reef flat) in the southern Maldives and the first record of upper reef flat condition in the central Indian Ocean post the 2015–2016 coral bleaching event. Scleractinian corals were the primary carbonate producers, with live coral cover averaging between 11.1 ± 6.5 and 31.2 ± 21.8% and dominated by massive corals. Gross carbonate production rates averaged 5.9 ± 2.5 G (kg CaCO The ability of reefs to maintain these functions in the future is uncertain, as coral reefs globally face multiple natural stressors, such as tropical cyclones/hurricanes4 and changes in sea-surface temperature (SST)5, which threaten the health and building capacity of reef systems. These natural stressors are compounded by anthropogenic factors including global climate change and pollution7. Indeed, periods of elevated SST just 1–2 °C above average summer SSTs can lead to coral bleaching, whereby temperature stressed scleractinian (reef-building) corals expel their symbiotic algae (zooxanthellae) from their coral tissue8, leaving the corals with a bleached appearance, with only the aragonite skeleton visible through translucent tissue. Bleaching may lead to coral mortality and bleaching/mortality can occasionally be widespread globally, often referred to as a mass global bleaching event10. Reefs affected by coral bleaching can experience reduced live coral cover5, reduced reef rugosity and consequent decreases in reef biodiversity11, carbonate production and reef accretion12. The third recorded mass global bleaching event occurred in 2015–2016 when El Niño Southern Oscillation (ENSO)-induced warm SST anomalies caused extensive coral bleaching and mortality in tropical waters throughout the Indo-Pacific and the western Atlantic Ocean14. The 2015–2016 coral bleaching event has been associated with declines in reef ecosystem functionality, reef productivity and reef carbonate budgets16. In the southern Republic of Maldives, SST persisted above the regional bleaching threshold of ~30.9 degrees Celsius for around three months from late March to mid-May 201616, leading to significant declines in live coral cover on fore-reef slopes. Observed reduction in live coral cover resulted in a collapse in reef carbonate budget state, with calcium carbonate production on these Maldivian fore-reefs switching from net positive (pre-bleaching) to net negative (post-bleaching), ultimately resulting in an inferred negative reef accretionary status16.Tropical coral reefs provide important biological functions and ecosystem goods and services, including dissipation of ocean wave energy, food and building resources, and income through tourism19. The primary carbonate producers on coral reefs are corals, with calcifying algae  contributing to secondary carbonate production, alongside other calcifying organisms (such as foraminifera and molluscs) and abiotic/microbial cementation. Organisms contributing to bioerosion include endolithic micro- and macro-boring organisms, parrotfish and echinoderms. Temporal shifts in carbonate budget state can be used as an indicator of trends in reef health and provide extended insights into biological erosion20, reef framework production and reef accretion potential22, compared with standard ecological abundance surveys, which do not consider the effects of net carbonate production or removal25.Carbonate budgets are one measure of reef condition that reflects the net amount of calcium carbonate produced within a reef system, and balances the biological accretion and erosion processes operating on an individual reef12. The temporal dynamics of carbonate budgets can have broader implications for reef systems and adjacent landforms. For example, recent collapses in reef carbonate budgets observed in the southern Maldives following the 2016 bleaching event16 could have cascading impacts for the geomorphic maintenance and stability of adjacent low-lying reef sedimentary landforms (beaches and islands). In the short-term (seasons to years), impacts may include reef or island accretion due to elevated carbonate sediment production associated with increased physical and biological erosion of dead coral. Sediment produced may be supplied to islands or contained within the reef framework and cemented, thus contributing to accretion. However, additional sediment production may also be transported off the reef system. In the longer-term (decades to centuries) reef island stability may be compromised due to a reduction in coral growth and the reef’s capacity to provide a physical barrier to incident wave processes that modulate beach and island erosion. Impacts may be compounded under future scenarios of sea-level rise, promoting widespread erosion and frequent island overtopping27. Such outcomes are of global concern, given recent suggestions that many coral reefs will be unable to continue accreting vertically at a sufficient pace to keep up with rates of projected global sea-level rise12. However, there are multiple interacting factors that will contribute to future reef accretion rates, which remain poorly resolved. These factors include: hydrodynamics and the rate of sea-level rise; the contribution of reef flat carbonate production to overall reef accretion; sediment delivery, retention and removal29 (including storm deposits); biological reef-binding processes ; microbial cementation; algal rim accretion rates; the role of carbonate production on sand flats, patch reefs and seagrass meadows; and biochemical dissolution of carbonate framework and sediment30.Understanding the effects of mass coral bleaching and mortality on reef carbonate budgets can also provide useful insights into a reef’s accretion potential under current and future sea-level rise scenarios32. However, recent work in the past couple of decades has expanded the growing knowledge of reef carbonate budgets to the central Indian Ocean35 and elsewhere in the Indo-Pacific37. A recent global analysis of carbonate budget data12 indicates that net carbonate budgets of reefs in the Indo-Pacific and tropical western Atlantic are currently low and are sub-optimal for reef accretion potential. Despite the growing number of studies of reef carbonate budgets, the overwhelming majority of data are derived from fore-reef slopes between 2–10 m depth38, where conditions for data collection are optimal . The contribution of the reef crest and reef platform surface (upper surface at ~0–1 m depth) carbonate producers to the overall reef budget is often not quantified, with few studies reporting reef flat carbonate budget rates37. This disparity in focus of studies is at odds with the fact that the reef crest and upper reef flat are critical environments when relating reef carbonate budgets to reef accretionary potential under sea-level rise and associated wave impacts at adjacent shorelines. The critical nature of these geomorphic sub-components rests on the importance of the reef crest and reef flat as the key wave energy dissipation zone on a reef39. Studies have shown that 94–98% of incident incoming wave energy is dissipated as waves interact with the reef edge and propagate across the reef flat1. Consequently, it is the residual energy from these interactions that controls the available energy to stimulate sediment deposition/removal at island shorelines41. Although the fore-reef slope environments play a key functional role in coral reef ecosystems, the potential for vertical reef accretion and the associated impact of theoretically higher water levels on reef flats under sea-level rise scenarios is largely dependent on carbonate production  and removal processes  operating at the reef crest and upper reef flat. Furthermore, the reef crest and reef flat are critical components of the carbonate factory for sediment supply to reef islands43, while sediment produced on the fore-reef slope is largely exported from the reef sytem45.Despite their recognized importance, carbonate budgets have been estimated for few reefs globally, most of which are located in the Caribbean3 m2 yr−1). We examine reef flat-wide net biological carbonate production with high spatial coverage of observations and evaluate the effect of the 2016 warm SST anomaly event on the reef flat carbonate budget. We highlight the differential roles of coral genera and growth morphologies in the carbonate budget of the reef platform and discuss the important contribution of bleaching-resilient massive corals (such as Porites spp.) to the reef flat live coral assemblage following major stress events.Here we present the first ecological survey of a coral reef flat in the Indian Ocean following the 2015–2016 global mass bleaching event. We quantify the carbonate budget of Mahutigala reef platform, a low-energy, sub-tidal, lagoonal platform reef located in Huvadhoo Atoll, Maldives  colonized areas of available substrate in the inner and outer reef zones. Peyssonnelia sp. was often found colonizing the available dead substrate and crevices of massive living coral colonies (Porites spp. and Heliopora spp. in particular).Five eco-geomorphological zones were differentiated on the Mahutigala reef surface based on 6,300 observations along seven radial transects around the reef platform. Zones were identified based on variations in substrate type including: sand and rubble cover, live and dead coral cover and coral morphology Fig. . An innePorites spp., but also Goniopora spp., Platygyra spp., Goniastrea spp. and Favia spp.) were the dominant contributor to live coral cover, comprising 66.3% of all live coral recorded on the reef flat  and the Octocoral Heliopora coerulea comprised 13.0% and 12.6% of all reef flat live coral, respectively, with the remaining 8.1% comprised of small colonies of encrusting, digitate, free-living and tabular corals of varying genera  , ranging from 11.1 ± 6.5% at the W transect to 31.2 ± 21.8% at the E transect Figs . Typicallat Fig. . Branchira) Fig. .H. coeru36. The average reef rugosity coefficient across the carbonate-producing zones of Mahutigala reef flat (zones 2–5) was 2.6, with averages ranging from 1.9 at the NE and SE transects and 2.9 at the N transect  were uncommon across the reef platform . All observed D. antillarum were within the 21 to 40 mm test size range, while P. imperialis were all within the 81 to 100 mm test size range. On average, echinoderms contributed only 0.1% to bioerosion.The total averaged biological erosion rate on the Mahutigala reef flat was estimated as 3.3 ± 0.4 G, with rates varying from 2.8 ± 0.7 to 3.9 ± 0.2 G across different radial transects Fig. . ParrotfNet carbonate production thus averaged 2.5 ± 2.4 G across the Mahutigala reef platform surface. While the carbonate production state was positive, on average reef erosion and accretion were balanced, given the considerable spatial variation in net G values on the platform, ranging from a maximum of 7.3 ± 5.3 G in the north to a minimum of −0.9 ± 2.3 G (negative budget) at the SE transect . Upper reef platform surfaces (reef crest and reef flat) are often neglected in studies that explore reef carbonate budgets, despite the upper reef surface acting as the carbonate factory for islands, and arguably being more important than the fore-reef slope in acting as a dissipative barrier to incoming wave energy1 and thus exerting strong control on the wave energy that reaches adjacent shorelines. Thus, the effects of the 2015–2016 bleaching event on reef crest and reef flat carbonate production are essential to consider, particularly in light of recent suggestions that declines in reef productivity and carbonate budgets could lead to a reduction in reef accretion potential. This could exacerbate reef island wave overtopping and associated erosion due to enhanced wave access to shorelines and sea-level rise16 and also affect the sediment supply to island shorelines.Our results present new insights of the effects of bleaching on the carbonate budget of a reef platform surface, expanding the limited number of budget studies in upper reef flat environments. Coral reefs experienced the third mass coral bleaching event in 2015–2016, causing significant declines in live coral cover, reef productivity and reef carbonate budgets in different regions16. This inferred coral collapse is supported by field observations prior to the bleaching event and a recent study of several fore-reefs on lagoonal platforms in Huvadhoo Atoll  that records a significant decline of 91% in branching and tabular Acropora spp. corals after the high SST anomaly event16. Despite high mortality of branching corals on the reef surface, observations indicate that the structural complexity of the reef platform surface remained comparable to pre-bleaching levels  and March 2017 (rugosity index of 2.0). However, expected reductions in structural complexity on the reef platform surface may be of a reduced magnitude than declines recorded on the reef slope, due to the predominance of massive corals on the reef flat, with lower surface-area ratios compared to branching and digitate corals, which dominated the fore-reef environment prior to the bleaching event16.Results of ecological surveys conducted four months after the 2016 bleaching event in the southern Maldives showed the Mahutigala reef platform surface had an average live coral cover of 17.9 ± 7.3%. Erosion and accretion were essentially balanced at Mahutigala reef platform, where an average net positive carbonate budget of 2.5 ± 2.4 G was recorded  used in this study across all orientations of the reef provides confidence to conclude that the majority of the Mahutigala upper reef platform is characterized by a positive carbonate budget. In contrast, Perry and Morgan’s16 survey was focused on only one sector of Mahutigala (the southwest) and interestingly, their recorded net negative G values are in accord with the nearby SE transect of the reef surface in this study, where net negative G (−0.9 ± 2.3 G) was also observed  recorded for Mahutigala reef platform surface of 2.5 ± 2.4 G, is up to 180% greater than the negative post-bleaching carbonate budget recorded for the Mahutigala reef slope . The higher net G values on the reef upper surface are directly attributable to the higher live coral cover (up to 37.3% and 19.7%) at the southeastern and southwestern reef platform, respectively  to the overall reef platform surface live coral cover is crucial to maintaining a positive carbonate budget following the bleaching event. The massive corals displayed strong resilience to the high SST anomaly event and did not experience mass mortality due to coral bleaching. However, it remains unclear whether these massive colonies temporarily bleached and recovered, or did not bleach at all.Net G values recorded on the southeastern and southwestern portions of the reef platform surface  were notably higher than that of the southwestern fore-reef slope46 and can reach the southeastern sector of Mahutigala reef by penetrating through the reef rim passage located 11 km to the southeast of Mahutigala. The northern sector of Mahutigala has a 50 m lagoonal fetch that likely generates significant internal wind-driven waves that impact the reef. Consequently, we infer exposure and hydrodynamic setting as a potential reason for the discrepancy in net G and massive coral dominance around Mahutigala. This highlights an important research gap that could be the focus of future investigations.The northern and eastern sectors of the reef platform displayed the highest average net G values  and several hypotheses have been proposed for explaining this preferential resilience including: greater tissue thickness of massive corals providing protection for the symbiotic alga within the coral tissue; and greater opportunity for mass transfer of toxins that accumulate during bleaching in massive, flat colonies compared to branching colonies51. Despite the shift in reef platform coral composition that has occurred at Mahutigala, from branching/digitate dominance to massive dominance, the platform surface still boasts a positive net carbonate budget, which has important implications for considering reef accretion potential.The differential survival of coral genera on the reef platform surface, with massive corals persisting and high mortality of branching, digitate and tabular corals, has led to marked changes in the reef community composition and structure, comparable to observations in the Maldives16 with recent studies suggesting that many coral reefs do not currently display vertical accretion rates sufficient to keep pace with projected rates of global sea-level rise12. However, the contribution of reef platform surface accretion is not considered in such studies, despite the reef flat acting as the critical geomorphic sub-component of a reef for wave energy dissipation40. Perry and Morgan16 identify a live coral cover threshold of around 12.0% for maintaining a net positive carbonate budget and a live coral cover threshold of around 8.0% for maintaining net positive reef accretion potential. Their threshold value for maintaining a net positive carbonate budget (12.0%) is comparable to that based on our data from Mahutigala reef flat of around 10.0% ; the morphologic response of the reef island to future sea-level rise and associated changes in hydrodynamic regimes; and the impacts of various anthropogenic activities on these natural processes.Carbonate budgets have been increasingly used to approximate reef accretion potential53, ultimately increasing reef biodiversity. At Mahutigala branching and digitate corals together comprised 14.5% of the total reef flat live coral composition in September 2016 , surrounded by an island beach, lagoon and elliptical shaped reef platform. Reef flat eco-geomorphological zones were visually differentiated across seven shore-perpendicular radial transect lines . To characterize benthic cover, observations were made at 10 cm intervals along the 10 m survey lines (100 observations per survey line) using the Point Intercept Transect method55, whereby the nature of the benthic cover directly below the tape measure at each 10 cm increment was recorded . Scleractinian corals were recorded to growth morphology  and genus level where feasible. Pre-bleaching average coral cover (%) is estimated based on unpublished data collected by postgraduate students from the University of Auckland in February 2016. Coral cover data were collected using the same Point Intercept Transect method, where observations of substrate cover were made at 10 cm intervals. Substrate rugosity was quantified along three randomly selected 1 m sections of each 10 m survey line using the chaining technique, whereby the total substrate surface length is divided by 1 m linear length of chain36. The three rugosity values were averaged to determine a rugosity coefficient (R) for each 10 m survey line. Gross carbonate production rates  for corals and coralline algae were estimated for each 10 m survey line using genera and coral morphology specific skeletal density and linear growth rates  using the following equation after33 and18:Xi is the mean percent cover of the ith genera. The calcification rate (Ci) of the ith genera was estimated in g cm−2 y−1 from mean published coral and coralline algae Ci rates from the Maldives .Ecological surveys were conducted at Mahutigala reef  by endolithic organisms, regional Indo-Pacific published rates were used  using equation:Biological erosion rates were conservatively estimated using a combination of published data and 16 based on belt transect surveys conducted at the Mahutigala fore-reef slope (2 m depth) in September 2016. Perry and Morgan16 estimated parrotfish bioerosion at Mahutigala as 3.81 ± 4.17 G and this rate was applied to all available substrate at Mahutigala reef flat. Uncertainties must be considered around using this published parrotfish erosion rate for the fore-reef slope and applying it to available substrate at Mahutigala reef flat. These uncertainties include disparities in parrotfish abundance, species and size class on the fore-reef slope compared with the reef flat, associated variability in bioerosion rates by fish of different species and size classes57, and unknown spatial differences in parrotfish abundance or potential feeding location preferences on the Mahutigala reef flat.To calculate bioerosion by parrotfish (G) we used published ratesDiadema spp. and Phyllocanthus spp. were calculated as a function of urchin genera and size, following18 using the equations:X is the average echinoid test size (mm) and e is the echinoid density in the 30 m2 belt transect. The average echinoid test size (mm) was 30 mm and 90 mm for Diadema spp. and Phyllocanthus spp., respectively. Macro- and micro-borer, parrotfish, and echinoid bioerosion rates were all extrapolated to the available reef substrate for erosion at Mahutigala and summed to provide an estimate rate (G) for total annual bioerosion. Net carbonate production rates (G) were determined by subtracting total bioerosion from gross production.Echinoid bioerosion rates were quantified within each of the productive eco-geomorphic zones on transects SW, NW, NE and SE at Mahutigala reef flat along two replicate 30 m-long by 2 m-wide belt transects. Urchin abundance and genera were recorded along the belt transects. Echinoid bioerosion rates (G) for Supplementary Information


The development of techniques allowing the systematic capture of the RNA-bound proteome has yielded many surprises. Among these, metabolic enzymes have been frequently detected as RNA-binding proteins (RBPs) by different profiling methodologies in various cell types . Compare+ into NADPH. Of note, α-KG is not only the substrate for subsequent TCA reactions, ultimately aiming to produce ATP, but also a cofactor for dioxygenases including DNA, histone, and RNA demethylases (Isocitrate dehydrogenases (IDHs) compose a family of enzymes (IDH1–IDH3) catalyzing the reversible conversion of the TCA product isocitrate into α-ketoglutarate (α-KG) , left, athylases . Moreovethylases . Some IDthylases , supportthylases . These oJournal of Molecular Cell Biology, + binding significantly impaired the ability to promote protein translation compared to wild-type IDH1 .]