OA is of particular concern for coral reefs, as the associated reduction in carbonate ion availability impairs biogenic calcification and promotes dissolution of carbonate substrata. While these trends ultimately affect ecosystem calcification, scaling experimental analyses of the response of organisms to OA to consider the response of ecosystems to OA has proved difficult. The benchmark of ecosystem-level experiments to study the effects of OA is provided through Free Ocean CO2 Enrichment (FOCE), which we use in the present analyses for a 21-d experiment on the back reef of Mo’orea, French Polynesia. Two natural coral reef communities were incubated in situ, with one exposed to ambient pCO2 (393 µatm), and one to high pCO2 (949 µatm). Our results show a decrease in 24-h net community calcification (NCC) under high pCO2, and a reduction in nighttime NCC that attenuated and eventually reversed over 21-d. This effect was not observed in daytime NCC, and it occurred without any effect of high pCO2 on net community production (NCP). These results contribute to previous studies on ecosystem-level responses of coral reefs to the OA conditions projected for the end of the century, and they highlight potential attenuation of high pCO2 effects on nighttime net community calcification.The Anthropocene climate has largely been defined by a rapid increase in atmospheric CO However, these goods and services are threatened by rising concentrations of atmospheric carbon dioxide (CO2) from anthropogenic sources2. A result of rising atmospheric CO2 concentrations is global climate change (GCC), which for tropical coral reefs is causing profound modifications in benthic community structure and function through thermally-driven mass bleaching events4. Ocean acidification (OA) is a process through which dissolution of CO2 into seawater lowers the calcium carbonate saturation state of seawater (Ω)5. In the marine environment, OA is expected to negatively affect organisms which produce calcareous shells and skeletons more than non-calcifying taxa8. These effects will be experienced acutely by coral reefs, where the physical structure of the calcareous substratum is built by scleractinian corals and calcified algae7. OA is predicted to severely impact the underlying biogenic substrata of coral reefs through increased dissolution in response to acidifying seawater conditions10. Field-based observations in natural CO2 seeps have shown decreased coral cover, diversity, and function in response to acidified conditions11. However, changes in carbonate chemistry across large spatial scales can interact with local effects to modulate processes that contribute to reef carbonate production 12. A common theme across papers addressing this topic is that OA is threatening the underlying biogenic physical structure of reefs and, therefore, understanding these effects has become a central objective of coral reef research in the 21st Century13.Tropical coral reefs provide coastal protection, livelihoods, and food to millions of people15. However, the complexities of understanding scaling effects16 as well as feedback loops between the inorganic substrata and living biota14 have resulted in difficulties attributing natural variation in NCC rates, and limited the ability to detect and attribute OA impacts on coral reefs. Laboratory-based studies with constructed communities have shown OA will increase dissolution of coral reefs18, as well as decrease their functionality through an alteration of the relationship between photosynthesis and calcification19.Depression of Net Ecosystem Calcification (NEC), the balance of gross calcification and gross dissolution, has been predicted for coral reefs in response to OA and warming7, in situ studies testing community-level responses are needed to predict how marine biota will respond to a changing climate20. Experiments conducted in situ, including those employing a Free Ocean CO2 Enrichment (FOCE) approach, are the benchmark for ecological relevance for determining OA effects on ecosystem function20 yet there are substantial challenges to implementing FOCE approaches on coral reefs. There are great benefits to addressing these challenges, however, as FOCE experiments embrace the natural complexity of ecosystems in terms of evaluating their response to OA, notably through emergent properties of multiple taxa interacting in a chemically and physically complex environment. To date, there have been two projects in which the effects of elevated CO2 have been empirically tested in situ on coral reefs. The first, by Albright et al. 2018, measured NEC of the back reef community at One Tree Islands, Australia, in response to short-term pulses of acidified seawater over multiple days15. A second project was conducted on Heron Island, Australia, in which deployment of technology described as a “coral proto-FOCE” (cp-FOCE) showed decreases in coral calcification21, and alteration in the boron isotopic composition of coral skeletons22. Here, we significantly expand on these previous experiments by conducting a FOCE experiment to test the effects of predicted end-of-century OA conditions on NCC and Net Community Production (NCP) of a natural back reef community in Mo’orea, French Polynesia. Our results show how the ecosystem function of coral reefs (NCC and NCP) will be altered by OA during the day and night, resulting in improved accuracy of predictions of the effects of OA on coral reef metabolism.While the impacts of OA on corals, other reef organisms, and reef communities are relatively well known2 under ecologically relevant environmental conditions23. Our experiment was conducted on two plots of coral reef (5.00 × 0.55 m) . Unidirectional flow speeds within the flumes were maintained at ~14 cm s−1 using motor-driven propellers, and were similar to average, long-term flow speeds recorded on the back reef  Fig.  that wer m) Fig. , and wer m) Fig. , and theef Table . Using a2  and elevated pCO2 targeted at ~1000 µatm  of human actions to control CO2 emissions24. NCC and NCP measurements were made over three consecutive days arranged into four sampling blocks equally spread over the 21-d experiment . Each day of measurements consisted of determinations in the morning, mid-day, afternoon, and once at night.Our experiment contrasted the effects of ambient pCOely Fig. ,B. The e2 treatment, and remained consistently depressed relative to 24-h NCC under ambient pCO2 15. In contrast, our study employed a greater decrease of pH , and a reduction in Ωaragonite of 1.89 between treatments, suggesting the decrease of NCC is not a linear function of Ω. On One Tree Reef, there was a higher cover of crustose coralline algae (CCA) (26%), and lower coral cover (12%) compared to the communities in our study  versus the back reef of Mo’orea could indicate that the coral reef community studied at One Tree Island was more sensitive to declining seawater pH than the present coral reef community studied in Mo’orea10.Following initiation of the experiment, 24-h NCC was depressed by 47% within the first day of the high pCOCO2 Fig. . This efCO2 Fig. , which c2 was depressed by 74% relative to ambient conditions. However, the magnitude of the pCO2-mediated depression of nighttime NCC attenuated over time, and after ~14 days the effect of pCO2 reversed, such that nighttime NCC in the elevated pCO2 treatment was ~1% higher than that under ambient conditions. After 21 days, nighttime NCC at elevated pCO2 was 27% greater than nighttime NCC at ambient pCO2 . The overall effect would result in a positive slope of NCC differences between ambient and high pCO2, shown as less reef dissolution over the 21-d experiment. While recent work has shown little acclimatization potential of reef calcifiers to OA25, the attenuation of the initially negative effect of high pCO2 on nighttime NCC of a shallow coral reef suggests these communities acquire some resistance to the negative effects of OA on nighttime NCC. The proximal mechanism(s) underlying this response are unknown, but potentially could include a rapid physiological acclimatization of calcification in organisms to high pCO2 as seen in polychaetes transplanted to CO2 vents26. While we did not observe a change in coral cover during the experiment, biological feedback loops caused by changes in relative composition of the microbial benthic community should be considered as a potential mechanism mediating the change in nighttime NCC27. Investigation into this mechanism might benefit from a further understanding of the relationship between biogenic processes that affect NCC and geochemical changes  within the pore water of the reef framework, as this interaction has been shown to be susceptible to OA28. Further work is required to explore these possibilities, and to determine whether the effects observed for nighttime NCC might also affect daytime NCC and/or 24-h NCC  was 41% higher in the community exposed to ambient pCO2 versus high pCO2 during the first 3-d incubation  for four months19. While CCA cover increased in both treatment groups in the present study, a greater proportional increase in algal turf cover and cyanobacteria occurred in the reef community maintained at high versus ambient CO2  comparing the response of mixed vs homogenous benthic community composition representing reefs found in Kaneohe Bay  to acidified conditions showed similar results to those reported here in which the NCP-NCC slope did not change significantly as a result of increased CO229. However, in the present study, we observed that the intercept was reduced by 58% for the community incubated under high CO2 (0.927 ± 0.402) versus ambient CO2 (2.129 ± 0.478) . While previous studies of reef community metabolism have not reported alteration of the intercept of the NCP-NCC relationship as a metric of change in metabolic function, the alterations of elevation between treatment groups while maintaining similar slopes seen in our study indicate that a shift in overall community function occurred in the community exposed to high CO2, where increased rates of NCP, are required to achieve similar rates of NCC19. For the reef communities in the present experiment, these changes likely are caused by a combination of decreases in the calcification rates of individual organisms and an increase in overall dissolution under high pCO210.Functional shifts in coral reef communities that alter the ratio of primary production to calcification are reflected in the slope of the NCP-NCC relationship, with a reduction in this relationship representing degradation in reef function, generally resulting from a shift in the dominant benthic community structure from calcifying organisms to algal-dominated communitiesime Fig. ,A. A pre3. Together with experimental analyses of the effects of global climate change and OA on corals, the rapid declines in coral cover have fueled concerns that coral reefs may not persist as calcified entities beyond the end of the current century15. The benthic community structure and ecological functions of coral reefs are tied intrinsically to the success of their ecosystem engineers, scleractinian corals and calcified algae, and these taxa already have been impacted by a wide variety of local disturbances that act in concert with global anthropogenic effects which cause a change in ecosystem functioning33. To date, predictions of how coral reefs will respond to OA have been based largely on species-level experiments testing the sensitivity of coral calcification to high pCO216. Our study highlights that the “organism approach” cannot capture the functional complexity arising from multiple organisms operating in concert in a natural environment subject to routine variation in select environmental conditions . An important implication of this outcome is that the whole coral reef community is more than the sum of its parts with regard to its response to OA. The present study highlights the importance of these emergent properties through in situ analysis of an undisturbed coral reef community. Our results suggest that these communities have the potential for adjustments to partially alleviate the negative consequences of OA on NCC. While this outcome might attenuate the risks of OA-related dissolution of the carbonate framework supporting most corals reefs10, the absence of a comparable effect on daytime NCC underscores that a further understanding of mechanisms that drive changes in NCC are needed.Global climate change is causing large declines in coral cover on reefs worldwide, particularly through the effects of mass bleaching2 Enrichment (SCoRe-FOCE) system23 was used to enrich pCO2 to levels projected for 2100 under representative concentration pathway (RCP) 8.5, which assumes a “business as usual” scenario with regards to anthropogenic emission of CO224. In February 2016, two plots were identified in the back reef, each 5.00 m long by 0.55 m wide, and they were selected to have a benthic community similar to this back reef in 200718, which consisted of 22% coral, 12% CCA, 23% turf algae, and 43% sand and rubble. Although this community structure differed from that occurring in 2018 when the experiment was conducted, our goal was to explore the response of an average back reef community, which is well represented by the state of the back reef in Mo’orea in 2007. When the study plots first were chosen, coral cover had declined compared to 2007 and, therefore, the community structure was augmented through transplantation of a few coral colonies from the adjacent reef to the study plots. Transplantations were completed >6 months prior to initiation of the incubations.Our study was conducted on the back reef of the north shore of Mo’orea, French Polynesia, which is a high volcanic island in the South Pacific. A custom-designed Shallow Coral Reef Free Ocean CO−1) into the closed flumes at the start and end of the experiment to evaluate the efficacy of the seal to the reef, and visual inspection was used to detect leaks; none were visible during ~2 h trials.Two-weeks before the start of the experiment (15 April 2018), two, 1.5-cm thick clear acrylic flumes (5.0 × 0.55 × 0.55 m (length × width × height) with UV-transparent tops were secured to fiberglass rails previously attached to the reef at the perimeter of each study plot. The internal volumes of each flume and the return-section was ~2,300 L. The flume was secured to the reef with stainless steel threaded rods drilled and epoxied into the carbonate substratum, and was sealed to the reef using rubber tubing inflated with water and placed between the supporting rails and the reef surface. Rubber matting extending 30 cm from the rails and weighted using 10-kg sand bags augmented the seal. An indicator dye  was injected (~1 mg L2 dosing to the flume was controlled through an Apex Aquacontroller (Neptune Systems), connected to an Atlas Scientific pH probe (ENV-40-pH), which controlled a solenoid (McMaster-Carr Model 5077T141) that injected CO2 gas into the flume from a 60-L gas cylinder on the floating platform. pH probes that controlled the autonomous dosing system were calibrated against pH values every 3 days using the m-cresol dye method (SOB 6 b Dickson34). Adjustments were made daily to maintain CO2 conditions within the flume  of 1000 µatm pCO2 , and the system was programed to decrease by 0.1 pH units , at night (18:00–06:00 h) to mimic in situ diel oscillations of pCO2 recorded on the backreef of Mo’orea35  within 15–30 min.During the experiment, each flume was flushed with ~200 L hent Fig. . Each daet al. [2007]) using an automatic titrator (Mettler-Toledo T50) fitted with a Rondolino-sample carousel (Mettler-Toledo). The titrator was fitted with a Mettler pH probe (DGi-115) that was operated with certified HCl (Batch A13 Dickson Laboratory). Certified reference material (Dickson CRM Batch #138) was used to evaluate the accuracy of the total alkalinity (AT) measurements (SOP 3b34).For each incubation, samples of seawater were collected at the beginning and end of each incubation to quantify seawater carbonate chemistry. Samples were drawn from the flume using a 60-mL syringe that was attached to a vinyl tube fitted with a shut off valve. Samples were transported immediately to the shore lab, where salinity was measured using a Thermo Scientific Orion Star A212 conductivity meter, then potentiometrically titrated following standard operating procedures (SOP 3b of Dickson 37 where −1) was calculated from the difference between AT in final and initial salinity-normalized water samples from each flume:t is time (h), SA is the planar surface area of the reef enclosed by the working section of the flumes (m2), ρ is the density of seawater , and V is the internal volume (L) of the flume (including the return sections).NCC was quantified using the alkalinity anomaly method2 concentration as a function of time, with dissolved O2 concentrations measured using MiniDOT O2 sensors  with one sensor in each flume. We chose to measure O2 flux as opposed to measuring DIC changes to maintain treatment conditions through CO2 dosing. Rates of change were determined using least squares linear regression of O2 concentration (mmol L−1) against time (h) . All O2 sensors were within factory calibration, which is stable for ~1 year.NCP was measured from the rate of change of O2 fluxes where DO is the change in O2 concentration (mg L−1), molar mass of O2 (32 g mol−1), SA is benthic surface area enclosed in the flumes (2.5 m2), t is incubation duration (h), and V is the volume of the flume (L).NCP was calculated from OTo calculate time-integrated values for NCC and NCP , hourly values were integrated over each 3–12-h incubation period and summed within each day to obtain 24-h values. For 12-hr daytime values, integrated 3 h incubation periods were summed from 06:00–18:00 h, and for 12-hr nighttime values, data were integrated from 18:00–06:00 h2 (ambient and high), and sampling period (incubations 1–4 throughout the whole experiment) were fixed factors, and 24-h NCC was the response variable. Each sampling period consisted of 3 consecutive days that were treated as statistical replicates. The effect of time (incubation day) on the difference of 24-h NCC between ambient and high pCO2 was tested using a Model I least squares linear regression, in which a significant slope indicated that the treatment effect  changed over time. Separate regressions were completed for daytime and nighttime NCC in order to distinguish between the effects of OA on NCC during the day and night. All analyses were performed in R.Twenty four hour NCC was analyzed using a two-way ANOVA in which pCO2 (ambient and high), and time (incubations 1–4 throughout each day) were fixed factors, and 12-h NCP or R was the response variable. Similar to NCC, each sampling period consisted of 3 consecutive days that were treated as statistical replicates.NCP and night community respiration (R) was analyzed in a similar way to NCC using a two-way ANOVA in which pCOsensu38) was evaluated from the relative scaling of calcification as a function of productivity, with decreases in this quotient generally representing a degradation of the reef, leading to increased primary producers, and decreased calcifiers. The relationship between NCP and NCC was calculated as the slope of the latter on the former (expressed as change in mmol CaCO3 per mmol O2) over the 21-d incubation, in which hourly NCP was regressed on hourly NCC using Model II regression31. The slope, elevation, and corresponding error (95% CI) for each of the flumes was calculated using a Major Axis (MA) approach, in which error on both x- and y-axes are accounted for39. Significance of differing slopes between the high and ambient CO2 treatment were evaluated using a Bartlett-corrected likelihood ratio statistic, and the p-value was calculated assuming a chi-squared distribution with 1 degree of freedom39. Difference in elevation (y-intercepts) between treatment groups was evaluated using a Wald statistic that tested for no difference among the y-axis intercepts, and similar to slope significance, this statistic was evaluated using a p-value assuming a chi-squared distribution with 1 degree of freedom39. This analysis was performed on an aggregation of all sampling points within the 21-d incubation , and analyzed using a model II linear regression with the package smatr in R.A change in community function (Supplementary Figures and Tables for Manuscript


C. elegans. Applying vIPR to the germline-specific transcript gld-1 led to enrichment of known and novel interactors. By comparing enrichment upon gld-1 and lin-41 pulldown, we demonstrate that vIPR recovers both common and specific RNA-binding proteins, and we validate DAZ-1 as a specific gld-1 regulator. Finally, combining vIPR with small RNA sequencing, we recover known and biologically important transcript-specific miRNA interactions, and we identify miR-84 as a specific interactor of the gld-1 transcript. We envision that vIPR will provide a platform for investigating RNA in vivo regulation in diverse biological systems.Understanding regulation of an mRNA requires knowledge of its regulators. However, methods for reliable de-novo identification of proteins binding to a particular RNA are scarce and were thus far only successfully applied to abundant noncoding RNAs in cell culture. Here, we present vIPR, an RNA-protein crosslink, RNA pulldown, and shotgun proteomics approach to identify proteins bound to selected mRNAs in  C. elegans gld-1 mRNA, including DAZ-1 and miR-84.The entire mRNA interactome capture has been established but identification of specific mRNA-binding proteins has been challenging. Here, the authors developed an in vivo RNA–protein crosslinking and RNA pulldown approach and characterized novel interactors of the  While the pool of RNAs bound by one RBP can be assessed by RBP immunoprecipitation (RIP) and crosslinking and immunoprecipitation (CLIP) approaches7, identifying all proteins bound to one particular RNA remains challenging. Conventional methods rely on genetic tagging of the RNA of interest or retrieval of in vitro-formed RNA-protein interactions (reviewed in ref. 8). Recently, also CRISPR-based approaches have been introduced to target native RNA-protein complexes10. These strategies have proven successful for several selected RNAs, but pose many problems impeding broader application. Constraints applying to one or all three approaches include: (1) not reflecting in vivo regulation, (2) laborious engineering, (3) high non-specific background, (4) non-physiological post-lysis associations of RNAs and proteins, and (5) inefficiency (reviewed in ref. 8). To overcome these limitations, additional methods have been developed in the last years. These employ crosslinking of native RNA-protein interactions in intact cells and pulldown of the RNA of interest by oligonucleotide probes under denaturing conditions16. This strategy enables efficient capture of endogenous transcripts with their native interactors while reducing background. However, these methods were not tested with mRNAs and key challenges remain. The lower the RNA of interest is expressed, the more input material is needed and the higher is the non-specific background from other abundant RNA-protein complexes. Consequently, most studies have focused on highly expressed transcripts thus far. Importantly, all these studies were performed in cell culture, not reflecting temporal and spatial regulation, e.g., interactions occurring in a specific developmental stage or in a specific tissue, altogether not recapitulating in vivo regulation.Throughout their lives, mRNAs are bound by proteins which regulate their biogenesis, transport, stability, localization, and translation. Understanding how an mRNA is regulated requires knowledge of its complement of protein binders. Techniques for mRNA interactome capture allow identification of hundreds of RNA-binding proteins (RBPs) associated with poly(A)+ RNA in diverse biological systems  light21, and its germline is a well-established model for the study of post-transcriptional regulation (reviewed in ref. 22), with 3′ UTRs rather than promoters determining expression of most genes23. Applying vIPR, we not only identify protein binders of two mRNAs expressed in the C. elegans germline, but also recover microRNAs (miRNAs) binding differentially to them. Our method is not restricted to mRNAs, but can be applied to any similarly-expressed RNA molecule in C. elegans. We anticipate that our method can be extended to application in any organism amenable to in vivo crosslinking, thus shedding light on a multitude of in vivo regulatory mechanisms in diverse biological contexts.Here, we describe vIPR (in vivo Interactions by ulldown of RNA), a method to identify factors binding to selected mRNAs in the complex context of an entire animal. We performed experiments in C. elegans with subsequent retrieval of the RNA of interest by an array of complementary oligonucleotides . Compared to these studies, challenges were (1) the reduced crosslinking efficiency in vivo, (2) isolation of the mRNA from a complex mixture of tissues, and (3) the expected lower number of bound proteins due to comparatively short 3′ UTRs and thus limited space for specific protein binding.To enable identification of proteins that interact with specific mRNAs in vivo, we developed vIPR (in vivo Interactions by Pulldown of RNA). vIPR relies on crosslinking of native protein–RNA interactions in live des Fig. . To achiC. elegans strain expressing a GFP fusion protein of the RBP GLD-1 at endogenous levels24. GLD-1 binds its own transcript at five reproducible binding sites in the 3′ UTR19 and thus can serve as a positive control. Additionally, the nearly identical RBPs FBF-1 and FBF-2  have been described to directly bind to gld-126. To retrieve the gld-1::gfp transgenic transcript, we designed probes exclusively tiling the gfp coding sequence, reasoning that this would permit straightforward application of the method to any other gfp strain. After transcript capture, proteins and RNAs were selectively eluted by nuclease or protease treatment, respectively. Eluted proteins were then identified by quantitative mass spectrometry and RNA was assessed by RT-qPCR or RNA sequencing . PFA-XL leads to both nucleic acid–protein and protein–protein linkages. Second, we used UV light at 254 nm (cXL) which results in direct crosslinks between RNA and protein only. Third, we employed PAR-XL, labeling of nascent RNA with 4-thiouridine (4SU) and subsequent activation of these modified nucleotides by UV irradiation at 365 nmgld-1::gfp pulldown efficiency and specificity, we measured RNA levels in input, supernatant, and elution samples by RT-qPCR. Irrespective of the crosslinking method, we retrieved ~60% of the input RNA  was barely detected in the elution and consistently found not to be depleted from the supernatant. Compared to tbb-2, the gld-1::gfp transcript was enriched ~20,000-fold in elution samples  polymerases GLD-2 and GLD-4 have been reported to activate and/or stabilize the gld-1 mRNA, likely by poly(A) tail extension28. These enzymes lack RNA-binding domains and are thought to be recruited to their targets by accessory proteins29. While GLD-3, RNP-8 and FBF have been found to co-precipitate with one or both enzymes in a complex with gld-1 mRNA31, it is currently unclear which protein(s) mediate the specific RNA contact, and apart from FBF, none of them was detected in our pilot experiment.While with all crosslinking methods, many proteins were detected non-specifically in both own Fig. . To testown Fig.  and alsoown Fig. . Importaach Fig. . A numbe33. Since many of these are known or predicted to bind RNA, it is possible that GLD-1 interacts with them via jointly bound RNA. We thus considered these proteins as additional candidate interactors of the gld-1 transcript. Interestingly, all proteins (7/7) consistently identified in both studies were enriched in the gld-1::gfp cXL pulldown 34 to accurately determine peptide intensities and only considered proteins quantified in all three gld-1::gfp pulldowns. Of the 273 reproducibly detected proteins, 29 proteins were found >4-fold enriched in all replicates  correction), comparing all target pulldowns with all no-target controls  and additionally identified further candidates (blue dots).In our pilot experiment, many of the enriched proteins were detected with few peptides only Fig. . To asseols Fig. . We coul35 and found that the set of candidate binders contains many more interaction edges than expected by chance  were annotated to function in or associate with RNA granules, suggesting a role for these proteins in RNA storage. Crosslinking with cXL is assumed only to capture direct RNA-protein contacts. Consistently, most candidates (21/24) are described or predicted to have RNA-binding activity, with motifs belonging to diverse RNA-binding domain classes  terms of the candidates, using all reproducibly detected proteins as the background set. GO terms related to RNA metabolism were highly over-represented in the candidate set Fig. . Interesses Fig. .gld-1. To this end, we performed additional vIPR experiments with another mRNA, gfp::lin-41. In adult C. elegans, lin-41 is predominantly expressed in the germline, as is gld-1. Using the same gfp-complementary probes as for gld-1::gfp, we performed triplicate pulldowns and reproducibly detected 278 proteins. We found 15 proteins consistently enriched >4-fold , the gld-1::gfp transcript consistently enriched for the proteins GLD-1 and DAZ-1 , (2) RNA retrieval specificity and efficiency, (3) the signal-to-noise ratio, (4) sensitivity in mass spectrometry, and (5) stringent controls. We successfully identified interactomes of the gld-1 and lin-41 transcripts which are far less abundant than most previously analyzed RNAs and are exclusively or predominantly expressed in one tissue of C. elegans, the gonad. We sought to establish a method to find direct RNA binders. Both PAR-XL and cXL have been commonly used to generate covalent linkages between RNAs and directly bound proteins, while PFA-XL also crosslinks indirectly bound proteins. PAR-XL has successfully been applied to recover protein-RNA interactions for three different RBPs in C. elegans20, and enables mapping of RBP binding sites at nucleotide resolution. However, since PAR-XL depends on labeling efficiency and has a stronger nucleotide bias than cXL, and since crosslinking with cXL recovered known and anticipated binders of the gld-1 transcript in our pilot experiment, we used cXL for all further experiments.Identification of proteins that bind to mRNAs in the context of an entire living animal is impeded by tissue complexity and a higher background of non-specific interactions. Generally, the success of an RNA pulldown experiment crucially depends on (1) the crosslinking efficiency . For ChIRP-MS and RAP-MS, several controls were tested that performed equally well in discriminating specific from non-specific binders12. In our experiments, choice of the control was crucial. Pulldown without prior crosslinking or pulldown after RNase treatment proved as inappropriate controls, presumably since these do not account for proteins crosslinked to background RNA. RAP-MS of the lncRNA Xist features transcript capture by ~140 90 nt long probes, with subsequent washes at elevated temperature under highly denaturing conditions12. The lower stringency of our washes, which is necessary to maintain transcript association of our 10–12 20 nt long probes, may contribute to the observed protein background in our no-target control. However, target RNA enrichment in vIPR experiments was comparable to the enrichment observed with RAP-MS12, suggesting that RNA background, and thus RNA-associated protein background, is similar with both approaches.To control for non-specific binding, we performed vIPR with gld-1::gfp and gfp::lin-41, we indeed found that many proteins bind to both transcripts . Consistently, gld-1 mRNA was shown to partly reside in P granules and to depend on P granule localization for interaction with the protein FBF-252.Both gld-1 and lin-41 pulldowns suggest that most enriched binders represent authentic in vivo interactors. However, we likely did not identify all transcript binders. PAR-XL and cXL were shown to recover different RBPs with different efficiencies3. Furthermore, some RBP classes, e.g., double-strand RBPs, are under-represented in UV-crosslink-based studies, and the sequence context of an RBP binding site can disfavor crosslinking53. The fact that we did not identify the known binder FBF in the gld-1::gfp triplicate pulldowns . GLD-1 acts as a translational repressor and stabilizes a subset of targets55. We identified DAZ-1 as an additional regulator of the gld-1 transcript. DAZ-1 is a germline-specific RBP important for oogenesis, and has been suggested to act as a translational activator56. Using a gld-1 3′ UTR reporter, we showed that protein expression was strongly downregulated upon daz-1 knockdown  the seed-complement and (2) bases pairing outside of the seed, both resulting in loss of miR-84 enrichment . Of note, ALG-1 has neither been identified in a recent study assessing the entire C. elegans mRNA interactome4, although ALG-1 CLIP has been performed successfully, and a large proportion of mRNAs are targeted by miRNAs21. This may be explained by a poor UV-crosslinkability of ALG-1. While a low crosslinking efficiency may suffice to detect crosslinked mRNAs in CLIP experiments, crosslinked RBPs may be missed in RNA pulldown experiments due to the lack of protein amplification methods and hence the lower sensitivity.The ent Fig. . FurtherC. elegans. We propose that mutation of an RBP binding site with subsequent vIPR may reveal changes in the composition of interacting proteins, thus providing insights into regulatory cascades. Furthermore, we anticipate application of vIPR to unravel differential regulation during development or in different cell types and of distinct transcript isoforms. We focused here on gfp transcripts for the ease of probe design and control. However, we demonstrated that endogenous transcripts can be retrieved similarly. vIPR should thus be readily applicable to any other similarly expressed C. elegans transcripts and we believe that it can be extended to discover interactions in any other animal or tissue amenable to UV crosslinking.In conclusion, vIPR allows identification of both proteins and miRNAs binding to RNAs of interest in live C. elegans strains (Supplementary Table Escherichia coli OP50-seeded NGM plates59. The MosSCI injection strain EG6699 was kept at 16 °C for maintenance. The BS1080 gld-1 transgenic strain24 was a gift from Tim Schedl. All other strains were obtained from the Caenorhabditis Genetics Center (CGC) or generated in the course of this study.gfp CDS or entire endogenous transcripts were designed essentially as described previously60. Probes were mixed to obtain a final concentration of 100 μM . Sequences are listed in Supplementary Table Probe design: Probes  tiling the 2PO4, 42 mM Na2HPO4, 86 mM NaCl, 1 mM MgSO4) without food source59. Experiments were typically performed with ~500,000 worms. Labeling of worms with the photoreactive nucleoside 4SU  was done in liquid culture  at 24 °C, as described before19. Worms were harvested as young adults , washed 3× in 0.1 M NaCl, transferred to non-seeded NGM plates and crosslinked in a custom UV Stratalinker 2400 crosslinker at a wavelength of 365 nm . For crosslinking methods not requiring labeling, worms were grown on OP50-seeded plates  until reaching young adulthood . After three washes in 0.1 M NaCl, worms were either transferred to non-seeded NGM plates and crosslinked at a wavelength of 254 nm  or incubated for 30 min with 2% PFA in M9, with subsequent quenching by 0.1 M Tris–HCl, pH 7.6, and two more washes in 0.1 M NaCl. Worms were pelleted, buffer was removed, and crosslinked worm pellets were frozen in liquid nitrogen. Pellets were kept at −80 °C until processed further.Preparation of worm pellets: Arrested synchronized L1 larvae were generated by bleaching of gravid adult worms and o/n hatching of larvae in M9 , 0.1 U μL−1 RiboLock RNase Inhibitor (Thermo Fisher Scientific)) and incubated for 30 min on ice. During incubation, lysates were passed 4× through a 20 gauge needle and 3× through a 25 gauge needle. For PFA-crosslinked samples, lysates were additionally sonicated  with 7 W for 2 min . All lysates were cleared by centrifugation , pellets were discarded, and supernatants filtered . Protein concentrations were determined using the Pierce BCA Protein Assay Kit (Thermo Fisher Scientific), and adjusted with lysis buffer, so that all samples processed simultaneously had the same final concentration . Lysates were diluted by adding 2× volume of pulldown hybridization buffer , and input samples were taken for RNA and protein analysis. The pulldown procedure was adapted from the protocol of Chu and colleagues60, with modifications.Worm lysis: After grinding worm pellets with mortar and pestle in liquid nitrogen, worm powder was resuspended in ~7× volume pulldown lysis buffer  were washed 3× in 1× original volume lysis buffer, resuspended in 0.5× volume lysis buffer, and added to the lysate. Preclearing was done at 37 °C for 1–2 h under constant rotation. To ensure complete removal of beads from lysates, tubes were placed on a magnet  and lysates were transferred to new tubes twice.Preparation of beads and preclearing: For pulldown with subsequent mass spectrometry, lysates were precleared prior to pulldown. MyOne Streptavidin C1 magnetic beads  were added to lysates and samples were incubated at 37 °C for 2 h or o/n. For probe capture, MyOne C1 beads, prepared as above, were added (100 μL per 50 pmol probes) and samples were incubated for an additional hour at 37 °C. Beads were separated from lysate and supernatant samples were taken. Beads were washed 5× with ~13 mL wash buffer . Finally, beads were resuspended in 1 mL wash buffer and transferred to Protein LoBind tubes (Thermo Fisher Scientific). A 50–100 μL aliquot was used for RNA isolation . Wash buffer was removed via magnet , and beads resuspended in Benzonase elution buffer ) for protein elution and Proteinase K buffer ) for RNA isolation, respectively.Probe hybridization and capture: Probes  was added, followed by Trizol RNA isolation (Thermo Fisher Scientific).Protein elution: Input and supernatant samples (10 μL) were resuspended in 100 μL and beads in 200 μL Benzonase elution buffer. Crosslinked proteins were eluted by incubation at 37 °C for 3 h, shaking at 1300 rpm. Beads were separated from elution via magnet, and elution was transferred to new Protein LoBind tubes. Samples were snap-frozen in liquid nitrogen and kept at −80 °C until processed further.61 prior to nanoLC–MS/MS analysis.For input and supernatant samples, 6–13 μg of protein were used for analysis by mass spectrometry. For pulldown elution samples, we roughly estimated the protein amount to be ~250 ng, based on comparison of peptide intensities of Trypsin and Benzonase in input and elution samples with corresponding total peptide intensities. Proteins were precipitated with ethanol and resuspended in 50 μL of 8 M urea and 0.1 M Tris–HCl, pH 8. Proteins were reduced with 10 mM DTT at room temperature for 30 min and alkylated with 50 mM iodoacetamide at room temperature for 30 min in the dark. Proteins were first digested by LysC (Wako) at a LysC-to-protein ratio of 100:1 at room temperature for 3 h. Then, the sample solution was diluted to a final concentration of 2 M urea with 50 mM ammonium bicarbonate. Trypsin (Promega) digestion was performed at a trypsin-to-protein ratio of 100:1 under constant agitation at room temperature for 16 h. Digestion was stopped and pH adjusted to <3.0 with TFA. Peptides were desalted with SCX (strong cation chromatography) and C18 Stage Tips62; 75 μm ID packed with ReproSil-Pur C18-AQ 3-μm or 1.9-μm resin, Dr. Maisch) connected on-line to the electrospray ion source (Proxeon) of a Q Exactive plus or a Q Exactive HF-X mass spectrometer (Thermo Fisher Scientific). Peptide samples were eluted at a flow rate of 250 nL min−1 with a 5–48% acetonitrile gradient in 0.1% formic acid over 2 h. Settings for MS analysis for Q Exactive Plus were as follows: one full scan  followed by top 10 MS/MS scans using higher-energy collisional dissociation (HCD) . Settings for MS analysis for Q Exactive HF-X were: one full scan  followed by top 20 MS/MS scans using HCD . Ions with an unassigned charge state and singly charged ions were rejected. Former target ions selected for MS/MS were dynamically excluded for 30 s.Reversed-phase liquid chromatography (rpHPLC) was performed employing an EASY nLC II (Thermo Fisher Scientific) using self-made fritless C18 microcolumns . Worms  were synchronized and grown until reaching young adulthood. After harvesting, worms were washed 3× with 0.1 M NaCl. Worms were transferred to non-seeded NGM plates, crosslinked at 254 nm (energy: 1 J cm−2), pelleted, and snap-frozen in liquid nitrogen. Pellets were kept at −80 °C until processed further. For the CLIP experiment, worm pellets were resuspended in 1 mL NP-40 lysis buffer  NP-40, 2 mM EDTA, 0.5 mM DTT, Complete mini EDTA-free Protease Inhibitor (Roche), 1 µg mL−1 Pepstatin, 1 mM PMSF, 0.1 U µL−1 RiboLock RNase Inhibitor (Thermo Fisher Scientific)) and homogenized with SiLibeads (Sigmund Lindner) in a tissue lyzer . Lysates were incubated on ice for 20–30 min, passed 10× through a 20 gauge needle, cleared by centrifugation , and filtered . After taking input samples, lysates were added to previously equilibrated α-GFP Trap_A beads (25 µL bead slurry per sample) for immunoprecipitation (90 min at 4 °C). After collection by centrifugation, beads were washed 4× with NP-40 lysis buffer and 2× with high-salt IP wash buffer  NP-40, 2 mM EDTA, 0.5 mM DTT, Complete mini EDTA-free Protease Inhibitor, 1 µg mL−1 Pepstatin, 1 mM PMSF). Beads and input samples were resuspended in 100 μL Proteinase K buffer ), and incubated at 50 °C for 45 min, shaking at 1300 rpm. Proteinase K was inactivated by boiling at 95 °C for 10 min. Samples were chilled on ice, and 1 μL GlycoBlue (ThermoFisher Scientific) was added, followed by Trizol RNA isolation (Thermo Fisher Scientific).CLIP-qPCR was performed to validate candidate interactors identified with vIPR. For reproducible and efficient immunoprecipitation, we used transgenic For quantification of RNA levels by qPCR, 1 µg of HEK RNA was added to samples prior to RNA isolation. Extracted RNA was RQ1 DNase-treated (Promega), and cDNA was generated with random hexamer primers, dNTPs and Maxima H Minus Reverse Transcriptase (Thermo Fisher Scientific). qPCRs were performed with Maxima SYBR Green/ROX qPCR Master Mix (Thermo Fisher Scientific) and 1/5 dilutions of cDNA as template. Oligonucleotides used for qPCRs are listed in Supplementary Table To extract total RNA from worms, worms were washed in 0.1 M NaCl, resuspended in 1 mL Trizol reagent (Thermo Fisher Scientific), and homogenized with SiLibeads (Sigmund Lindner) in a tissue lyzer , followed by RNA extraction according to the Trizol protocol. Pulldown elution samples for RNA sequencing were supplemented with 1 μg HEK RNA before RNA isolation to enhance the efficiency and reproducibility of RNA precipitation. Additionally, samples from PAR-XL and PFA-XL pulldowns contained ERCC RNA Spike-In Mix 1 . After Trizol RNA isolation, ribosomal RNAs were depleted by one of the methods described below.63. Briefly, 500 ng RNA were mixed with 500 ng ribosomal RNA-complementary DNA oligonucleotides  in hybridization buffer , heated at 95 °C for 2 min and cooled to 45 °C at a rate of −0.1 °C s−1 for specific annealing of oligonucleotides to ribosomal RNA. Samples were supplied with an equal volume of RNase H buffer (1 μL Hybridase RNase H (Lucigen) per 500 ng RNA, 0.2 M NaCl, 0.1 M Tris–HCl, pH 7.6, 40 mM MgCl2), mixed and incubated for additional 30 min at 45 °C for ribosomal RNA depletion. RNA was extracted with Agencourt RNAClean XP beads (Beckman Coulter) and TURBO DNase-treated (Thermo Fisher Scientific) for 30 min at 37 °C to remove DNA oligonucleotides. Reaction was stopped by addition of EDTA to a final concentration of 15 mM and incubation at 75 °C for 10 min. RNA was purified a second time by RNAClean XP beads and resuspended in 19.5 μL of Elute, Prime, Fragment Mix from the TruSeq RNA library prep kit v2 (Illumina).rRNA depletion by RNase H treatment: Ribosomal RNA depletion with RNase H was essentially performed as described by Adiconis et al.rRNA depletion with RiboMinus kit: Elution RNA of PAR-XL and PFA-XL pulldowns was TURBO DNase-treated for 30 min at 37 °C. Reaction was stopped by addition of EDTA to a final concentration of 15 mM and incubation at 75 °C for 10 min. RNA was purified by RNAClean XP beads, and ribosomal RNA depletion performed with the RiboMinus eukaryote probe mix v2 (Thermo Fisher Scientific), according to the manufacturer’s protocol. After depletion, RNA was ethanol-precipitated (o/n), and resuspended in 19.5 μL Elute, Prime, Fragment Mix from the TruSeq RNA library prep kit v2.Library preparation and sequencing: All samples were subjected to fragmentation, reverse transcription and adapter ligation, as described in the TruSeq RNA sample preparation v2 guide. By means of a pilot qPCR with 1/10th of generated libraries and library- and adapter-specific primers, the optimal Ct for library amplification was determined. Library quality, quantity and average fragment sizes were assessed by Bioanalyzer 2100  and Qubit (Thermo Fisher Scientific). Samples were sequenced in 1 × 76 run mode on a NextSeq 500 system (Illumina) or 1 × 51 run mode on a HiSeq 4000 system (Illumina).2O, and 7 µL were used for library preparation with the SMARTer smRNA-Seq Kit for Illumina (Clontech). After PCR purification, the library concentration was measured with Qubit (Thermo Fisher Scientific) and size distribution was checked by Bioanalyzer . The libraries were size-separated via BluePippin , according to the SMARTer smRNA-Seq Kit protocol. Size-selected libraries were again analyzed by Bioanalyzer (HS DNA chip). Equimolar amounts of libraries were pooled and loaded onto a NextSeq 500 (Illumina) in 1 × 76 run mode or a HiSeq 4000 (Illumina) in 1 × 51 run mode.RNA from target and no-target pulldowns was extracted with Trizol reagent, after addition of 1 µL GlycoBlue (Thermo Fisher Scientific). RNA was RQ1 DNase-treated (Promega), and purified with Roti-Aqua-PCI (Carl Roth), followed by ethanol precipitation. RNA was resuspended in 8.5 µL H64. The final construct contained the gld-1 promoter and 5′ UTR, the GFP::H2B CDS, and the gld-1 3′ UTR within the backbone of the vector pCFJ151. The transgenic strains were generated by the MosSCI technique66, resulting in single-copy insertion of the transgene at a defined locus on chromosome II of the C. elegans genome. Oligonucleotides used for construct cloning are listed in Supplementary Table The reporter construct was generated by conventional restriction-ligation cloning, as described before59. Mock (L4440 empty vector) and daz-1 (sequence name in library: F56D1.5) RNAi clones were obtained from the Ahringer library. Per well in a 6-well plate, 400 L1 animals were seeded. For analysis of RNA levels, young adult worms were harvested after growth at 24 °C for 45 h, washed in 0.1 M NaCl, and resuspended in 500 µL Trizol reagent (Thermo Fisher Scientific). After homogenization with SiLibeads (Sigmund Lindner) in a tissue lyzer , insoluble material was pelleted by centrifugation for 5 min at 16,100 × g. From the supernatant, 350 µL were used for RNA isolation by means of the Direct-zol kit (Zymo Research), which includes a DNase digestion step. Of the purified RNA, 100 ng were used as input for amplification-free RNA quantification by the Nanostring nCounter gene expression assay using a 72-multiplex custom Nanostring Gene Expression code set on a nCounter SPRINT profiler (Nanostring Technologies) following the manufacturer’s instructions. Raw counts were normalized to the internal positive controls and to two reference genes , using the nSolver 4.0 software  on a 2% agarose pad. Pictures of GFP expression in gonads were taken on an inverted fluorescence microscope  with a 40× objective . All gonads were monitored using the same exposure time (1/7.5 s). Pictures were processed and analyzed using ImageJ in an identical manner. Germline GFP fluorescence was quantified by measuring pixel intensity profiles with ImageJ along a segmented line (thickness: 35) from the distal tip to the bend of each gonad arm as described previously64. Background mean gray values were subtracted, and values were binned into 30 bins. Averages and standard error of the mean (SEM) were calculated for all gonads analyzed per condition and independent reporter line. Quantification was restricted to distal gonads as absence of DAZ-1 leads to meiosis arrest of oogenic germ cells at the pachytene stage56.Young adult worms were mounted by picking clean worms into a drop of 1 mM levamisole in M9  was pre-annealed with the target crRNA  and the dpy-10 crRNA  by incubation at 95 °C for 5 min and cooling to 25 °C at a rate of −0.1 °C s−1. Recombinant Cas9 protein (9 µg) was mixed with the pre-annealed RNA, the target repair ssODN  and the dpy-10 ssODN  in a solution containing 150 mM KCl and 7.5 mM HEPES, pH 7.4 . The mix was injected into gonads of adult wild-type N2 worms. Sequences of oligonucleotides used for CRISPR are listed in Supplementary Table Genome editing for destruction and conversion of the 68; version: 0.43.1; parameters: reads = single end, bootstraps = 100) to the C. elegans transcriptome , and if applicable to the gld-1::gfp or gfp::lin-41 transgene sequence (determined by Sanger sequencing), and the ERCC RNA Spike-Ins. For pulldown elution samples that contained HEK RNA, the human transcriptome (GENCODE Release 24) was added to the index. Average fragment length and corresponding standard deviation were set according to the respective library Bioanalyzer profiles. Ribosomal RNAs (and overlapping pseudogenes) and, if applicable, human RNAs and ERCC Spike-Ins, were removed from output, and TPMs were re-normalized to the remaining transcripts. TPMs were summed up per gene, and plotted with the R software package.After de-multiplexing , sequencing reads were pseudo-aligned with kallisto  for removal of E. coli RNA contamination. miRNAs were then identified by miRDeep2  with the miRBase21 reference. For miRNA enrichment analysis, raw read counts were plotted with the R software package and enrichment was determined by comparing regression-normalized miRNA counts.After de-multiplexing , adaptors and poly(A)-tails were removed using cutadapt (version 1.14) and the following parameters --format = fastq --adapter = AAAAAA --error-rate = 0.2 --times = 3 --overlap = 3 --cut = 3 --quality-cutoff = 20 --quality-base = 33 --minimum-length = 17 --max-n = 0 --discard-untrimmed. The adaptor- and poly(A)-trimmed libraries were subsequently mapped with bowtie2 (version 2.3.3.1) using the parameters --very-fast-local --phred33 --local to the 70; v1.5.1.2, 1.5.7.4, or 1.6.0.16) with standard settings, unless stated otherwise in the following. Search parameters included two missed cleavage sites, fixed cysteine carbamidomethyl modification, and variable modifications including methionine oxidation, N-terminal protein acetylation, and asparagine/glutamine deamidation. The peptide mass tolerance was 6 ppm for MS scans and 20 ppm for MS/MS scans. The match between runs option was enabled. Database search was performed using Andromeda71 against the UniProt/Swiss-Prot worm database  with common contaminants. The false discovery rate (FDR) was set to 1% at both peptide and protein level. Protein quantification was done based on razor and unique peptides. The label-free algorithm34 based on peptide extracted ion chromatograms (XICs) was used. The minimum LFQ ratio was set to 1 (vIPR triplicates of transgenic transcripts) or 2 (vIPR of endogenous transcripts). For analysis of identified proteins, known contaminants, proteins only identified by site, and reverse mappings were filtered out from MaxQuant output. For pulldown samples, we additionally removed proteins with gene ontology (GO) terms related to “biotin” , which are expected to enrich unspecifically due to direct binding to the streptavidin-coated beads. For the initial performance test of crosslinking methods, we compared raw peptide intensities. For more accurate quantification of the cXL vIPR triplicate experiments, we compared LFQ intensities. Here, imputation of missing intensities was done with the Perseus software package , after log2-transformation of LFQ values . To determine significance of proteins identified in triplicate, we calculated p-values with a moderated t-test, implemented in the Bioconductor LIMMA package73, and corrected for multiple comparisons by the Benjamini-Hochberg procedure74.Raw data were analyzed and processed using MaxQuant (http://www.uniprot.org/uploadlists/) and curated manually. Wormbase IDs were then used to retrieve gene ontology (GO) terms. Test for over-representation  was performed by the online tool at http://pantherdb.org/ (75; results in Supplementary Data gld-1::gfp pulldowns. Of 271 genes, 269 were retrieved and had annotated GO terms.Majority protein IDs from MaxQuant output were mapped to Wormbase IDs by Uniprot ID mapping (Further information on research design is available in the Supplementary InformationPeer ReviewReporting SummaryDescription of Additional Supplementary FilesSupplementary Data 1Supplementary Data 2Supplementary Data 3Supplementary Data 4Supplementary Data 5Supplementary Data 6Source Data


Nuclear RNAs are subject to a number of RNA decay pathways that serve quality control and regulatory functions. As a result, any virus that expresses its genes in the nucleus must have evolved mechanisms that avoid these pathways, but the how viruses evade nuclear RNA decay remains largely unknown. The multifunctional Kaposi’s sarcoma-associated herpesvirus (KSHV) ORF57 (Mta) protein is required for the nuclear stability of viral transcripts. In the absence of ORF57, we show that viral transcripts are subject to degradation by two specific nuclear RNA decay pathways, PABPN1 and PAPα/γ-mediated RNA decay (PPD) in which decay factors are recruited through poly(A) tails, and an ARS2-mediated RNA decay pathway dependent on the 5ʹ RNA cap. In transcription pulse chase assays, ORF57 appears to act primarily by inhibiting the ARS2-mediated RNA decay pathway. In the context of viral infection in cultured cells, inactivation of both decay pathways by RNAi is necessary for the restoration of ORF57-dependent viral genes produced from an ORF57-null bacmid. Mechanistically, we demonstrate that ORF57 protects viral transcripts by preventing the recruitment of the exosome co-factor hMTR4. In addition, our data suggest that ORF57 recruitment of ALYREF inhibits hMTR4 association with some viral RNAs, whereas other KSHV transcripts are stabilized by ORF57 in an ALYREF-independent fashion. In conclusion, our studies show that KSHV RNAs are subject to nuclear degradation by two specific host pathways, PPD and ARS2-mediated decay, and ORF57 protects viral transcripts from decay by inhibiting hMTR4 recruitment. Eukaryotic cells contain numerous nuclear RNA quality control (QC) systems that ensure transcriptome fidelity by detecting and degrading aberrant RNAs. Some viral RNAs are also predicted to be degraded by these RNA QC systems, so viruses have evolved mechanisms that counter host RNA QC pathways. Previous studies showed that the Kaposi’s sarcoma-associated herpesvirus (KSHV) expresses the ORF57 protein to protect its RNAs from nuclear decay. However, neither the specific host pathways that degrade KSHV RNAs nor the mechanisms describing ORF57 protection of viral RNAs were known. Our data suggest that ORF57 protects viral RNAs from two different nuclear RNA QC pathways, PABPN1 and PAPα/γ-mediated RNA decay (PPD) and an ARS2-mediated RNA decay pathway. Mechanistically, we show that ORF57 binds directly to viral RNAs and prevents the recruitment of hMTR4, a cellular factor whose function is to recruit the exosome, the complex responsible for RNA decay, to the transcript. We conclude that by preventing hMTR4 recruitment, ORF57 protects viral RNAs from degradation resulting in robust expression of viral genes. The Kaposi’s sarcoma-associated herpesvirus (KSHV) is a nuclear double-stranded DNA virus that belongs to the gammaherpesvirus family. It infects human B lymphocytes and endothelial cells and causes Kaposi’s sarcoma (KS) as well as the lymphoproliferative disorders primary effusion lymphoma (PEL) and some cases of multicentric Castleman’s disease (MCD) –4. The Kpromoter upstream transcripts  are polyadenylated, non-coding RNAs with no or few introns that are made from bi-directional protein-coding promoters [In eukaryotes, RNA QC systems play an essential role in RNA metabolism –13. Messromoters –16. RNA romoters –19. Otheromoters . Some prromoters , 21, 22.romoters , 21, 22.The PABPN1 and PAPα/γ-mediated RNA decay (PPD) pathway is an RNA decay pathway that degrades polyadenylated nuclear transcripts , 21, 22.Nuclear transcripts can alternatively be eliminated by a pathway called the CBCN-mediated decay pathway that involves two complexes: the cap-binding complex (CBC) and the nuclear exosome targeting (NEXT) complex . The CBCKSHV transcripts resemble those of their host in that they are capped and polyadenylated. However, 70% of KSHV genes are intronless while the majority of human protein-coding genes contain multiple introns , 29. MosIn order to avoid transcript decay in the nucleus, KSHV has evolved mechanisms that allow efficient expression of its genes in the absence of splicing. The KSHV ORF57 (Mta) protein is a post-transcriptional regulator of gene expression that is essential for virus replication –42. HomoHere we used RNAi to target host nuclear decay pathways and monitored their contributions to the degradation of PANΔENE. Depletion of cellular factors involved in PPD and depletion of ARS2 stabilized the transcript, but knockdown of NEXT components had no effect on decay. In transcription pulse-chase assays, ORF57 protects PANΔENE from ARS2-mediated decay, but it is unclear whether it protects PANΔENE from PPD. In the context of viral infection in cultured cells, inactivation of both decay pathways restored the expression of a subset of ORF57-dependent viral genes produced from an ORF57-null bacmid. Thus, our data suggest that ORF57-mediated protection of viral RNAs from these nuclear decay pathways is important for viral replication. Mechanistically, we show that ORF57 inhibits recruitment of the exosome co-factor hMTR4 to viral RNAs. In addition, our data suggest that ORF57 promotes the recruitment of ALYREF to some viral RNAs to increase the stability of the viral transcript presumably by impeding the interaction between ARS2 and hMTR4. We conclude that ORF57 contributes to KSHV gene expression by hampering the activity of host-mediated nuclear RNA QC pathways.To identify host-mediated nuclear RNA decay pathways that target KSHV viral transcripts for degradation and to determine whether ORF57 acts on these pathways, we used a well-defined transcription pulse-chase assay , 70. In PPD degrades polyadenylated RNAs with few or no introns, at least partially due to their inefficient export . MoreoveWe next investigated the role of the CBCN-mediated nuclear RNA decay pathway in PANΔENE decay and ORF57 stabilization. To do so, we monitored PANΔENE stability upon CBCN inactivation by knocking down the NEXT components RBM7 or ZCCHC8 in the presence or in the absence of ORF57. Both RBM7 and ZCCHC8 were efficiently depleted at the mRNA level  and funcOur data suggest that both PPD and an ARS2-dependent decay pathway target PANΔENE for degradation but ORF57 preferentially inhibits the latter. Therefore, we tested whether these two pathways are redundant or they are part of the same pathway. We co-depleted PAPα/γ and ARS2 and monitored PANΔENE stability by northern blot. Inactivation of both decay pathways in the absence of ORF57 stabilized PANΔENE to a greater extent than PAPα/γ or ARS2 alone compare . FurtherThe nuclear exosome requires cofactors to carry out its functions including the RNA helicase hMTR4 that has been implicated to be part of both CBCN and PPD , 71, 72.To expand our studies of the role of ORF57 in nuclear decay to viral infection, we generated an ORF57 knockout using the KSHV infectious clone BAC16 . The intORF57 has been implicated in both transcriptional and posttranscriptional regulation, either of which could be affected in the context viral infection. Because much our work here focuses on two well characterized ORF57 target genes, PAN RNA and ORF59 mRNA, we tested whether the reduced expression of these transcripts in iSLK-ΔORF57 cells was due to a transcriptional or a post-transcriptional defect. To do so, we treated cells for 10 minutes with 4-Thiouridine (4SU), a uridine analog that is efficiently incorporated into RNA by all three cellular polymerases. After harvesting RNA, 4SU can be specifically biotinylated and newly made transcripts can be captured by streptavidin selection. The levels of nascent and newly made ORF59 and PAN RNA were unchanged between iSLK-WT and iSLK-ΔORF57 cells . We concOur pulse-chase reporter assays Figs  and 2 arWe next tested whether PPD promotes viral RNA decay during lytic reactivation. We knocked down PABPN1, PAPα/γ, or ZFC3H1 to inactivate PPD in iSLK-WT and iSLK-ΔORF57 cells . ConsistOur data suggest that viral transcripts are targeted for degradation by an ARS2-dependent decay pathway and PPD during lytic infection, but ORF57 counters these activities. To examine redundancy between the pathways, we inactivated both decay pathways by simultaneously knocking down PAPα/γ and ARS2 in iSLK-WT and iSLK-ΔORF57 cells. Following lytic reactivation, PAPα/γ and ARS2 co-depletion restored ORF59 mRNA levels to a greater extent than PAPα/γ or ARS2 alone in iSLK-ΔORF57 cells . SimilarThe steady-state measurements of PAN RNA and ORF59  do not dTo elucidate the mechanism of ORF57 in RNA stabilization, we first focused our attention on ARS2. Previous reports showed that like the CBC-associated ARS2, ORF57 binds RNAs near their 5′ end  and inteWe next tested whether ORF57 affects hMTR4 recruitment to viral RNAs. Initially, we used native RIP to test the association of ORF57  and hMTRA major caveat of native RIP experiments is the reassortment of protein-RNA complexes that often occurs subsequent to cell lysis . To circin vitro approach to further test the model that ORF57 inhibits hMTR4 recruitment to RNA. Whole cell lysates from cells transfected with either an ORF57 expression plasmid or an empty vector control were incubated with an in vitro transcribed, radioactively labeled PAN RNA substrate. The probe included the ORF57 responsive element in the 5ʹ-end of PAN RNA and an m7G cap [Finally, we used an  m7G cap , 62. Foltrans with an ORF57 expression plasmid. We used 293i cells because they allow robust transfection and overexpression compared to iSLK derivatives. Interestingly, ALYREF overexpression partially restored ORF59 levels in 293iΔORF57 while PAN RNA accumulated to levels even higher than those observed in 293i-WT cells , ALYREF binding promotes their nuclear export, reducing their exposure to nuclear decay pathways and increasing the abundance of the transcript. However, our data here as well as previously published reports suggest this is not the case. First, in the absence of ORF57, ORF59 is subject to similar nuclear decay pathways as PAN RNA , and it Unlike other ORF57-dependent transcripts, ORF6 expression cannot be restored in ORF57-null cells upon inhibition of nuclear decay. Instead, our data suggest that ORF6 is regulated transcriptionally . BecauseThe recent definition of the NEXT, CBCA, and PAXT complexes represents a major step forward in understanding nuclear RNA decay in mammalian cells , 24, 27.During viral infection, the deletion of ORF57 results in viral RNAs being subject to two decay pathways, PPD and another involving ARS2 . PPD tarThe interrelationships between PPD and ARS2-mediated decay processes for viral or host RNAs remain somewhat obscure. We further note that the abbreviations PAXT and PPD were proposed independently , 19, butOur data suggest that ORF57 neutralizes the action of nuclear RNA decay pathways by preventing hMTR4 interaction with viral transcripts . InteresIt remains to be determined how cellular RNA decay pathways affect the expression of most KSHV genes. As KSHV genes are transcribed and processed in the host nucleus, all of the resulting transcripts must avoid RNA QC pathways. A complete understanding of these mechanisms will be challenging as they involve RNA decay, processing, and its coordination with transcription. In fact, the exosome and its cofactors have been implicated in regulating transcription of cellular and viral genes , 89. The2 in DMEM (Sigma) supplemented with 10% Tet-Free fetal bovine serum , 1x penicillin-streptomycin (Sigma), and 2 mM L-glutamine (Fisher). iSLK-WT and iSLK ΔORF57 cells were grown in the presence of 0.1 mg/mL G418 (Fisher), 1 μg/mL puromycin (Sigma) and 50 μg/mL hygromycin. 293i cells were maintained in media containing 50 μg/mL hygromycin. HEK293 cells  were cultured in the same conditions, but regular FBS (Sigma) was used instead.293A-TOA , 293i an- recombination-deficient E. coli strain that contains a heat-inducible Red recombinase system and an arabinose-inducible I-SceI homing endonuclease gene. Bacteria were grown at 30–32° except where noted. GS1783 bacterial cells and a plasmid containing a kanamycin-resistance gene with an adjacent I-SceI site (GS1439) were generous gifts of Dr. Greg Smith (Northwestern University) and Dr. Nikolaus Osterrieder (Freie Universität Berlin).The markerless Red Recombination system was used to make the BAC16-ΔORF57 bacmid –92. Ampl600~0.5 in 30 μg/mL chloramphenicol LB broth, then transferred to 42° water bath for 5 min to induce recombination enzymes and subsequently chilled in an ice bath for 20 min. The cells were harvested, washed twice in ice-cold 10% glycerol, then resuspended in 50 μL of the same. One hundred nanograms of the gel-purified I-SceI/kanamycin/ORF57 targeting PCR product was then electroporated in 1 mm cuvettes . The bacteria were then grown in 1 mL of LB for 2hr/30° and plated on LB plates with chloramphenicol and kanamycin. Colonies were allowed to grow for two days then individual clones were selected and bacmid DNA was isolated by miniprep using standard techniques.For the first step of the two-step Red recombination system, we performed PCR using primers NC1175 and NC1176 with GS1439 as a template to create a PCR product that has an I-SceI site next to the kanamycin resistance gene. In addition, the primers contain unique left and right homology regions to target the ORF57 locus of BAC16. They also contain the desired ORF57 deletion sequence flanking the I-SceI containing kanamycin resistance gene for recombination in step 2 (see below). We obtained GS1783 cells carrying BAC16 from Dr. Jae Jung  . These bThe clones were screened by three independent validation strategies. Proper targeting was verified by colony PCR using primers NC1179/NC1178 and NC1177/NC497  which spAfter selection of clones with proper targeting and full-length TRs, we performed the second step of the Red recombination approach that removes the kanamycin resistance gene and creates the ORF57 deletion. Overnight cultures of the positive clones were diluted (10 μL into 1 mL of LB+chloramphenicol), grown for 2 hr at 30°, then diluted 1:1 with LB/chloro containing 2% arabinose to induce I-SceI expression. This generates a dsDNA break at the I-SceI site flanking the kanamycin-resistance gene. One hour after arabinose addition, the cells were shifted to 42° for 5 min then placed at 30° for an additional 3 hours. This step induces the recombination machinery to drive homologous repair between the duplicated ORF57-deletion sequences that flank both sides of the kanamycin-resistance gene. Colonies were grown on chloramphenicol plates lacking kanamycin and screened by: 1) verification of loss of kanamycin-resistance, 2) PCR using NC1179 and NC497, 3) PFGE. Bacmid DNA was prepared from the positive clones using the Qiagen Large-Construct kit and the resulting bacmids were once again validated with PFGE and the junction was sequenced to verify the precise 33-bp ORF57 deletion. BAC16 and deletion bacmid were then transfected into iSLK cells using FuGene and selected with 300 μg/mL of hygromycin . MultiplAll the proteins studied here are essential, so care was taken to optimize siRNA conditions to produce the most efficient functional knockdowns with the lowest cell toxicity. iSLK cells were transfected with 20 or 40 nM siRNA  using RNAiMAX transfection reagent (Invitrogen) per manufacturer’s instruction. Specifically, we used final concentrations of 40 nM siRNAs for ZFC3H1, RBM7, ZCCHC8 and hMTR4 and 20 nM siRNAs for PABPN1 and ARS2. For PAPα/γ, we used 20 nM each of siRNAs that target PAPα or PAPγ for a total of 40 nM siRNA. Twenty-four hours post siRNA transfection, cells were split into new plates and allow to grow for another 24 hours, after which doxycycline and NaB was added to induce lytic reactivation. Total RNA was harvested 72 hours post siRNA transfection and 24 hours post lytic reactivation. For PAPα/γ knockdown, we observe a more robust effect after 4 days of silencing . HoweverFor pulse-chase assays in 293A-TOA cells, knockdowns were performed following the conditions described for iSLK cells. Twenty-four hours post transfection, cells were split into new plates and allow to grow for another 24 hours, after which they were transfected with the appropriate reporter construct. The one exception was hMTR4. In this case, cells were allowed to grow for 48 hours before being transfected with the reporter construct. For DNA (PANΔENE reporter) transfections, cells grown in a single well of a 12-well plate were transfected with 0.8 μg of DNA using TransIT-293 (Mirus) following manufacturer’s protocol. siRNAs are listed in Transcription pulse-chase assays were performed as previously described , 61. Bri2, 4 mM spermidine, 10 mM DTT, 200 ng T7-driven template, 0.5 mM ATP, CTP, and GTP, 2 U/μL RNasin, 50 μM UTTPαS (TriLink Biotechnologies), 25 μCi of α-32P-UTP (800 Ci/mmol), and T7 RNA polymerase. Probes were made from PCR-generated DNA templates or by enzymatic digestion of a plasmid   and 250 μM PMSF. Proteins were resolved by SDS-PAGE and western blotted using standard procedures. Antibodies used are given in RNA was harvested using TRI reagent according to the manufacturer’s protocol. Following extraction, RNA was treated with RQ1 DNase (Promega). Random hexamers were used to prime cDNA synthesis with MuLV reverse transcriptase (New England Biolabs). Real-time reactions used iTaq Universal SYBR Green Supermix (Biorad). Primers are listed in Lytic reactivation of iSLK derived cells was achieved by adding doxycycline (1 μg/ml) and NaB (1mM). Ninety-six hours post induction, media from iSLK-WT, iSLK-ΔORF57 and iSLK-ΔORF57 + pLenti-ORF57 cells were collected, centrifuged for 5 min at 1000 xg and passed through a 0.45 um filter. Polybrene was added  and 300 μL of pre-warmed virus-containing media were applied to HEK293 cells grown in a 12-well plate. Cells were centrifuged for 45 min at 30°C and then incubated in 5% CO2 at 37°C for 2 hours. After this, media was replaced and 24 hours later, cells were analyzed by FACS.2, 0.5% Triton X-100) supplemented with protease inhibitors and RNasIn Plus (Promega). Next, CaCl2 was added to 5 mM with 45U of RQ1 DNase and incubated for 15 min at 25°C. For RNA digestion, Micrococcal Nuclease  was diluted 1:200 and 10 μL of this freshly diluted stock was added to the extract. RNA digestion proceeded at 25°C for precisely 10 min, after which 50 μL of 300 mM EGTA was added to stop the reaction. After clarification of the lysate by centrifugation at 16000 xg for 10 min, the protein-RNA complexes were immunoprecipitated for 1 hr with anti-hMTR4, ARS2, ORF57, ALYREF or IgG1 antibody . Samples were heated at 65°C for 5 min and immediately placed on ice for 2 min. After this, 400 μL of RIPA correction buffer  were added to each sample and run through a QIAGEN QiaShredder column 2 times. Next, CaCl2 was added to 5 mM with 45U of RQ1 DNase and incubated for 15 min at 25°C. For RNA digestion, Micrococcal Nuclease (2x106 Gel Units/ml) was diluted 1:200 and 10 μL of this freshly diluted stock was added to the extract. RNA digestion proceeded at 25°C for precisely 10 min, after which 34 μL of 300mM EGTA was added to stop the reaction. After clarification of the lysate by centrifugation at 16000 x g for 10 min, the Protein-RNA complexes were immunoprecipitated for 1 hr with anti-hMTR4, ORF57 or IgG1 antibody. Twenty microliters of washed Protein A Dynabeads were added to each sample for 1 hr. Beads were then washed with RIPA buffer five times and then eluted as described for native RIP. Following elution, samples underwent PCA and chloroform extraction and were ethanol precipitated. The resulting RNA was then analyzed by qRT-PCR.CLIP was performed as previously described , 76 withSeventy-two hours after PAPα/γ and ARS2 knockdown and 24 hours post lytic induction, iSLK-WT and iSLK ΔORF57 cells were treated with 500 μM 4-Thiouridine (4SU) for 10 minutes. RNA was then harvested using TRI reagent and treated with RQ1 DNase. 80 μg of RNA were biotinylated at room temperature for 3 hours in a solution of 10 mM Tris-HCl pH 7.5, 1mM EDTA, 0.1% SDS and 0.2 mg/mL Biotin-HPDP (Thermo Fisher). Biotinylated RNA was extracted with chloroform three times, and then ethanol precipitated. Streptavidin selection was performed using 20 μL of Dynal MyOne Streptavidin T1 bead slurry (Thermo Fisher). Prior to incubation with sample, beads were washed 3 times with MPG 1:10-I  and then blocked for 1 hour in the same solution supplemented with 0.1 μg/μL poly(A), 0.1 μg/μL cRNA, 0.1 μg/μL ssDNA and 0.1% SDS. Biotinylated RNA and blocked beads were nutated for 1 hour at room temperature and then washed 10 times as previously described . Samples32P-UTP (800 Ci/mmol), 1U/μL of RNasIn Plus, 1 mM each of ATP, CTP and GTP, 50 μM UTP and transcription buffer . Reaction was incubated at 37°C for 2 hours and then run through a MicroSpin G-50 column (GE). RNA was ethanol precipitated, resuspended in 15 μL of water and capped using the Vaccinia Capping system (NEB) following manufacturer’s recommendations. Capped RNA was extracted 1X with PCA, 1X with chloroform, ethanol precipitated and, then, separated on a urea-PAGE. RNA band of the correct size was excised and eluted with G-50 buffer  overnight. Eluted RNA was extracted with PCA, ethanol precipitated and resuspended in water. RNA yield was determined by scintillation counting.Body labeled RNA probe was made using a plasmid expressing KSHV PAN RNA as template in a 20 μL reaction containing 50 units of T7 RNA polymerase, 2.5 μL of 2, 50 mM DTT, 10 mM ATP and 3 mg/mL torula yeast RNA), 0.5 μL RNasin Plus and 105 CPM of radioactively labeled RNA probe. Once the binding reactions were completed, volumes were increased to 250 μL by adding 225 μL of 1X IP buffer . Specific antibodies  or 1.6 μg of FLAG-ALYREF + 0.4 μg ORF50 using TransIT-293 reagent following manufacturer’s protocol. Similarly, 293i ORF57 cells were transfected with either 1.6 μg of pcDNA3 + 0.4 μg of ORF50. Thirty-six hours post transfection, total RNA was harvested using TRI reagent. The resulting RNA was then analyzed by qRT-PCR.S1 Fig(A) Quantitative western blots showing PABPN1, hMTR4 (arrowhead), ARS2 and transient transfected ORF57 protein levels following PAPα/γ, PABPN1, ZFC3H1, hMTR4, ARS2, RBM7 or ZCCHC8 knock down. Actin serves as loading control. (*) unspecific band. (B) Knock-down efficiency of PAPα, PAPγ, ZFC3H1, RBM7 and ZCCHC8 was determined by qRT PCR due to the lack of robust antibodies. Bar graphs showing results from qRT-PCR of PAPα, PAPγ ZFC3H1, RBM7 and ZCCHC8 obtained from 293A-TOA cells where PAPα, PAPγ, ZFC3H1, RBM7 and ZCCHC8 were depleted using siRNAs. Values were normalized to β-actin and plotted relative to siCtrl. (C-D) Because RNA knockdown does not necessarily correlate with protein loss, we assayed for loss of functional activity. To do so, we measured the RNA levels of two known PPD targets (SNHG4 and SNHG19) and two known NEXT complex targets (proEXT1 and proMGST3) by qRT-PCR. (C) Bar graphs showing results of SNHG4 (white) and SNHG19 (black) obtained from 293A-TOA cells where ZFC3H1 was depleted or PAPα and PAPγ were co-depleted. (D) Bar graphs showing results of PROMPTs EXT1 (black) and MGST3 (white) obtained from 293A-TOA cells where RBM7 or ZCCHC8 were depleted. All values are average and the error bars are standard deviations (n = 3).(TIF)Click here for additional data file.S2 Fig(A) Representative northern blot of transcription pulse-chase assay in cells expressing an empty vector or ORF57 and transfected with a two-siRNA pool targeting RBM7. 7SK serves as loading control. (B) Decay curves of biological replicates of the transcription pulse-chase assays after siRBM7; each point is a mean value with standard deviation (n = 3). See (TIF)Click here for additional data file.S3 Fig(A-B) Quantitative western blots showing PABPN1, hMTR4 (arrowhead), ARS2 and ORF57 protein levels following PAPα/γ, PABPN1, ZFC3H1, hMTR4, ARS2, RBM7 or ZCCHC8 knockdown in iSLK-WT (A) and iSLK-ΔORF57 cells (B). Actin serves as a loading control; (*) unspecific band. (C) Bar graphs showing results from qRT-PCR of PAPα, PAPγ, ZFC3H1, RBM7 and ZCCHC8 obtained from iSLK-WT (green) and iSLK-ΔORF57 (black) cells where PAPα, PAPγ, ZFC3H1, RBM7 and ZCCHC8 were depleted using siRNAs. (D-G) See (TIF)Click here for additional data file.S4 Fig(A-C) Western blot of protein from native RIP of ARS2 (A), ORF57 (B) and hMTR4 (C). (D-E) Western blot of protein from CLIP of ORF57 (D) and hMTR4 (arrowhead) (E). (*) unspecific band. All samples were collected 24 hours post lytic reactivation of iSLK-WT and iSLK-ΔORF57 cells.(TIF)Click here for additional data file.S5 Fig(A) Western blot of proteins from ALYREF overexpression in 293iΔORF57 cells and knock-down efficiency of ARS2 and hMTR4 (arrowhead). All samples were collected 36 hours following ORF50 transfection to induce lytic reactivation. (*) unspecific band. (B) Bar graphs showing results from qRT-PCR of PAPα (black), PAPγ (white) and SNHG19 (gray) where PAPα and PAPγ were depleted using siRNAs. Values were normalized to β-actin and plotted relative to siCtrl. All values are average and the error bars are standard deviations (n = 3). See (TIF)Click here for additional data file.S1 TableAll siRNAs were purchased commercially .(XLSX)Click here for additional data file.S2 TableAll templates for in vitro transcription were generated by PCR with a T7 promoter on the reverse primer. The one exception was the ORF47 probe which was made by a cut plasmid as indicated.(XLSX)Click here for additional data file.S3 TableAll antibodies were commercially available as indicated.(XLSX)Click here for additional data file.S4 TableTarget, sequence, and primer number (ID) for all PCR primers used herein.(XLSX)Click here for additional data file.


ZIKV is a recently emerged mosquito-borne flavivirus that can cause devastating congenital Zika syndrome in pregnant women and Guillain-Barré syndrome in adults. The molecular mechanism of ZIKV virion assembly is largely unknown. Here, we report that ZIKV NS2A plays a central role in recruiting viral RNA, structural protein prM/E, and viral NS2B/NS3 protease to the virion assembly site and orchestrating virion morphogenesis. One mutation that impairs these interactions does not significantly affect viral RNA replication but selectively abolishes virion assembly, demonstrating the specific role of these interactions in virus morphogenesis. We also show that the 3ʹ UTR of ZIKV RNA may serve as a “recruitment signal” through binding to NS2A to enter the virion assembly site. Following a coordinated cleavage of C-prM-E at the virion assembly site, NS2A may present the viral RNA to C protein for nucleocapsid formation followed by envelopment with prM/E proteins. The results have provided new insights into flavivirus virion assembly. The flavivirus virion consists of an envelope outer layer, formed by envelope (E) and membrane (M) proteins on a lipid bilayer, and an internal core, formed by capsid (C) protein and genomic RNA. The molecular mechanism of flavivirus assembly is not well understood. Here, we show that Zika virus (ZIKV) NS2A protein recruits genomic RNA, the structural protein prM/E complex, and the NS2B/NS3 protease complex to the virion assembly site and orchestrates virus morphogenesis. Coimmunoprecipitation analysis showed that ZIKV NS2A binds to prM, E, NS2B, and NS3  in a viral RNA-independent manner, whereas prM/E complex does not interact with NS2B/NS3 complex. Remarkably, a single-amino-acid mutation (E103A) of NS2A impairs its binding to prM/E and NS2B/NS3 and abolishes virus production, demonstrating the indispensable role of NS2A/prM/E and NS2A/NS2B/NS3 interactions in virion assembly. In addition, RNA-protein pulldown analysis identified a stem-loop RNA from the 3ʹ untranslated region (UTR) of the viral genome as an “RNA recruitment signal” for ZIKV assembly. The 3ʹ UTR RNA binds to a cytoplasmic loop of NS2A protein. Mutations of two positively charged residues (R96A and R102A) from the cytoplasmic loop reduce NS2A binding to viral RNA, leading to a complete loss of virion assembly. Collectively, our results support a virion assembly model in which NS2A recruits viral NS2B/NS3 protease and structural C-prM-E polyprotein to the virion assembly site; once the C-prM-E polyprotein has been processed, NS2A presents viral RNA to the structural proteins for virion assembly. Many flaviviruses are significant human pathogens, including Zika (ZIKV), dengue (DENV), yellow fever (YFV), West Nile (WNV), Japanese encephalitis (JEV), and tick-borne encephalitis (TBEV) viruses. Flaviviruses are formed by 180 copies of envelope (E) and membrane (M) proteins that are anchored onto a host-derived lipid bilayer .The molecular mechanism of flavivirus assembly remains largely unknown. Flavivirus and other RNA virus assemblies are coupled with RNA replication , 4. Duri7–The current study uses ZIKV as a model to study flavivirus assembly. We demonstrate a central role of ZIKV NS2A in recruiting viral RNA, prM/E complex, and NS2B/NS3 complex to virion assembly sites. A 3ʹ UTR stem-loop RNA of the viral genome may serve as a “recruitment signal” to bind to a cytoplasmic loop of NS2A. Mutations that impair the interactions of NS2A with viral RNA, prM/E, or NS2B/NS3 are lethal for virion assembly. These results suggest that ZIKV NS2A recruits viral RNA, unprocessed C-prM-E polyprotein, and NS2B/NS3 protease to the virion assembly site where coordinated cleavage of C-prM-E initiates genomic RNA and C protein encapsidation followed by prM/E envelopment and virion budding.One major challenge in studying flavivirus NS2A is the lack of good antibodies against this protein. We failed to raise antibodies for flavivirus NS2A after many attempts using different approaches (data not shown). To overcome this challenge, we constructed a hemagglutinin (HA)-tagged NS2A ZIKV (HA-NS2A ZIKV) using an infectious cDNA clone of Cambodian strain FSS13025 . The HA cis-Golgi marker), indicating that NS2A is associated with the ER membrane. The HA-NS2A was also colocalized with viral C, prM, E, NS1, NS3, and NS4B proteins, as well as double-stranded RNA (dsRNA). Because NS5 mainly resided in the cell nuclei, it did not show a strong fluorescent signal in the cytosol to costain with NS2A protein  to examine the cells that had been infected with the E103A HA-NS2A ZIKV. A single-round infectious E103A HA-NS2A ZIKV was prepared by entation  in whichentation . The resentation . Both WTentation , whereasentation . The resTaking advantage of the HA-tagged NS2A protein, we performed coimmunoprecipitation (co-IP) to identify viral proteins that bind to NS2A in ZIKV-infected cells. In WT HA-NS2A ZIKV-infected cells, HA-NS2A selectively pulled down prM, E, NS2B, and NS3 proteins but not C, NS1, NS4B, or NS5 proteins , lane 7.Since prM/E and NS2B/NS3 form individual complexes, we examined if HA-NS2A interacts with one or both proteins from the prM/E and NS2B/NS3 complexes. HEK293T cells were cotransfected with two plasmids: one plasmid encoding HA-NS2A and the other encoding Flag-tagged prM, E, NS2B, or NS3 protein. Co-IP results showed that HA-NS2A pulled down individual prM-Flag , E-Flag trans complementation described in To explore the molecular defect of NS2A E103A on virion assembly, we performed co-IP on cells that were infected with E103A HA-2A ZIKV   . To dete2 h p.t. . Further2 h p.t. . These r2 h p.t.  and D, eTo further explore the role of ZIKV NS2A in virion assembly, we examined the ability of NS2A to bind viral RNA using an RNA immunoprecipitation (RNA-IP) assay. WT HA-NS2A ZIKV-infected cell lysates were precipitated by antibodies against HA, viral NS1 protein, or cellular protein claudin-2 . The amoin vitro RNA-protein pulldown assay to characterize the NS2A-viral RNA interaction (S-transferase (GST-NS2A) was expressed from Escherichia coli, partially purified, and reconstituted in detergent n-Dodecyl-β-d-maltopyranoside (DDM)-containing buffer. ZIKV 5ʹ UTR-C66-3ʹ UTR RNA  was in vitro transcribed and ligated to a biotinylated cytidine using T4 RNA ligase, resulting in 5ʹ UTR-C66-3ʹ UTR-biotin RNA. The 5ʹ UTR-C66-3ʹ UTR-biotin RNA efficiently pulled down GST-NS2A but not GST  (dK (dissociation constant) of 10.3 nM for ZIKV NS2A binding to viral RNA  than the WT , suggestin vitro RNA-protein binding assay to examine the role of R96 and R102 in NS2A binding to viral RNA. To focus on the RNA binding to the NS2A cytosolic loop region, we prepared recombinant protein GST-NS2A(58–133), representing amino acids 58 to 133 of NS2A with an N-terminal GST (66-3ʹ UTR-biotin RNA (66-3ʹ UTR-biotin (lane 4). As a negative control, E103A mutant GST-NS2A(58–133) did not compromise the RNA binding (lane 5). The results indicate that R96 and R102 are important for NS2A binding to viral RNA.We performed an inal GST . WT GST-otin RNA , lane 3,Next, we performed an RNA-IP assay to compare the viral RNA binding to NS2A between the WT and ΔSL-2 HA-2A RNA-transfected cells. At 24 h p.t., the cell lysates were precipitated by HA antibodies. The amount of indicated RNA coprecipitated with HA-NS2A was quantified by RT-qPCR. Compared with the cellular GAPDH and U1 RNAs, the WT viral RNA was ∼2-fold enriched in binding to HA-NS2A protein, whereas ΔSL-2 RNA was not significantly enriched . CollectFlavivirus infection induces double-membrane vesicles (Ve) where viral replication complexes reside –27. At t–7–The biological relevance of NS2A/prM/E and NS2A/NS2B/NS3 interactions was supported by the NS2A E103A mutation that abolished virion assembly as well as NS2A binding to prM/E or NS2B/NS3 complexes  and 3A. Besides losing binding to other viral proteins, the NS2A E103A mutation also led to the production of a truncated C protein, with the first 6 amino acids removed by viral NS2B/NS3 protease . We showOur biochemical and functional analysis indicates that the 3ʹ UTR of viral RNA may serve as an “RNA recruitment signal” for ZIKV assembly. Although the 3ʹ UTR of Kunjin virus was reported to bind to NS2A , the molDuring flavivirus assembly, the cleavage between C and prM is tightly regulated in a sequential manner by NS2B/NS3 protease and cellular signalase –34. The –37–Finally, it should be noted that, in addition to the viral proteins described here, cellular factors (including proteins and lipids) have been well documented to participate in virion assembly . Studiesl-glutamine, 100 U/ml penicillin, 100 g/ml streptomycin, and 10% fetal bovine serum . BHK-NS2A-HA cells .African monkey kidney epithelial (Vero E6) cells, baby hamster kidney (BHK-21) cells, and human embryo kidney 293T (HEK293T) cells were purchased from the American Type Culture Collection (ATCC) and cultured in high-glucose Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 2 mM HA cells  constituThe following antibodies were used in this study: mouse monoclonal antibody 4G2 cross-reactive with flavivirus E protein (ATCC); rabbit anti-E polyclonal antibody (GeneTex); rabbit anti-HA monoclonal antibody ; mouse anti-HA monoclonal antibody ; rabbit anti-Flag polyclonal antibody ; mouse anticapsid antiserum (generated in-house); rabbit anti-prM polyclonal antibody (GeneTex); rabbit anti-NS1 polyclonal antibody (GeneTex); rabbit anti-NS3 polyclonal antibody (GeneTex); mouse monoclonal antibody cross-reactive with ZIKV and DENV-2 NS4B (clone 44-4-7) ; mouse aRenilla luciferase (Rluc) reporter ZIKV replicon (pZIKV Rep WT) as reported previously (XhoI and KpnI restriction sites at positions 3018 and 5010 in the ZIKV genome were introduced into the ZIKV infectious cDNA clone (pFLZIKV) and the eviously . The DNA18) and the prM with a C-terminal Flag tag into the pXJ vector. C18 was retained to ensure the correct ER membrane targeting of prM. A DNA fragment encoding the last 17 amino acids of M protein (M17) and the E protein with a C-terminal Flag tag was cloned into pXJ vector, resulting in construct pXJ-E-Flag. M17 served as a signal peptide for translocation of E into the ER lumen. For the construct pXJ-HA-NS2A, a DNA fragment encoding a signal peptide from Gaussia luciferase (SPG), the last 16 amino acids of NS1 protein (C16), and NS2A with an N-terminal HA tag were fused and cloned into the pXJ vector. SPG and C16 sequences were engineered to ensure the correct processing and membrane topology of NS2A.A mammalian expression vector, pXJ , was useS-transferase (GST) tag in E. coli, the corresponding DNA fragments were cloned into the vector pGEX-4T-1 .For constructs expressing full-length NS2A or its various mutants or truncations with an N-terminal glutathione in vitro transcribed from full-length cDNA clone plasmids (linearized by ClaI restriction enzyme) using the T7 mMessage mMachine kit (Ambion) and electroporated into cells (Vero or BHK-21) using the Gene Pulser XCell electroporation system (Bio-Rad) to generate virus as mentioned previously .The replicon transient-transfection assay was performed as described previously . BrieflyCells were lysed in IP buffer  with rotation at 4°C for 1 h. Cell lysates were then clarified by centrifugation at 15,000 rpm at 4°C for 15 min. The supernatants were mixed with specific antibodies or IgG isotype control (1:200 diluted), after which additional sodium chloride was added to a final concentration of 300 mM. For RNase A treatment groups, RNase A was added to a final concentration of 60 μg/ml. The mixtures were rotated overnight at 4°C to form immune complexes. Immune complexes were captured by protein A/G magnetic beads (Millipore) with 1-h rotation at 4°C. Thereafter, bead-bound immune complexes were collected by a DynaMag-2 magnet (ThermoFisher Scientific) and washed five times with chilled phosphate-buffered saline (PBS) containing 0.1% Tween 20. Finally, the bead-bound immune complexes were eluted by boiling in 2× lithium dodecyl sulfate (LDS) sample buffer supplemented with 50 mM dithiothreitol (DTT) at 70°C for 15 min. The eluates were analyzed by SDS-PAGE and Western blotting as described previously . Goat antrans-complementation as shown in 2) for 2 h at room temperature and then washed twice with 100 mM cacodylate buffer. The monolayers were scraped off and centrifuged at 12,000 rpm for 5 min to pellet cells. The pellets were postfixed with 1% OsO4 in 100 mM cacodylate buffer, washed with electron microscopy (EM)-grade water, and en bloc stained with 2% aqueous uranyl acetate in water for 20 min at 60°C. The pellets were dehydrated in ethanol series (from 50% to 100%) and processed through propylene oxide before being embedded in Poly/Bed 812 (Polysciences). Ultrathin sections (70 nm) were cut on a Leica EM UC7 ultramicrotome (Leica Microsystems), stained with 0.4% lead citrate in water for 3 min, and examined with a Philips CM100 transmission electron microscope at 60 kV. Digital images were acquired with a bottom-mounted charge-coupled device (CCD) camera, Orius SC2001 .Vero cells were infected with single-round infectious E103A HA-NS2A ZIKV  with rotation for 1 h. Cell lysates were clarified by centrifugation at 13,000 rpm for 15 min, mixed with specific antibodies or IgG isotype control (1:200 dilution), and rotated overnight. Immune complexes were incubated with protein A/G magnetic beads with 1-h rotation, after which bead-bound immune complexes were washed five times with washing buffer . All procedures for RNA-IP were completed at 4°C or on ice. Finally, buffer RLT from the RNeasy minikit (Qiagen) was added to extract the total RNA that had been captured in the bead-bound immune complexes by following the manufacturer’s instructions.RNA immunoprecipitation (RNA-IP) was performed as described previously  with somin vitro-transcribed full-length ZIKV RNAs.Quantitative real-time RT-PCR (RT-qPCR) was performed using an iTaq Universal SYBR Green one-step kit (Bio-Rad) on the LightCycler 480 system (Roche) by following the manufacturer’s protocol. Primers ZIKV-6878F (5ʹ-CATGGTAGCAGTGGGTCTTC-3ʹ) and ZIKV-6980R (5ʹ-CTCCTCTCTCCTTCCCATTAGA-3ʹ) were used to determine ZIKV RNA level; primers GAPDH-F (5ʹ-AGGTCGGTGTGAACGGATTTG-3ʹ) and GAPDH-R (5ʹ-TGTAGACCATGTAGTTGAGGTCA-3ʹ) were used to determine the GAPDH mRNA level; primers U1-F (5ʹ-GGGAGATACCATGATCACGAAGGT-3ʹ) and U1-R (5ʹ-ATGCAGTCGAGTTTCCCACA-3ʹ) were used to determine the U1 level. The absolute quantification of ZIKV RNA was determined by standard curve method using E. coli by following a protocol as described previously (E. coli strain Rosetta 2(DE3)pLysS cells (Novagen). The cells were cultured in Terrific Broth medium (Invitrogen) with 100 μg/ml ampicillin and 34 μg/ml chloramphenicol at 37°C. Once the cell culture reached an optical density at 600 nm (OD600) of 1.0, isopropyl-β-d-thiogalactopyranoside (IPTG) was added to a final concentration of 0.5 mM to induce protein expression. After growing at 16°C overnight, the cells were pelleted down by centrifugation at 8,000 × g for 10 min at 4°C and resuspended in a lysis buffer , followed by ultrasonication at 4°C. The lysates were clarified by centrifugation at 20,000 × g for 10 min. The supernatants were further centrifuged at 39,000 rpm for 1.5 h in an SW40Ti rotor (Beckman Coulter) at 4°C to pellet down the membrane fractions. Afterward, the pellet was resuspended in a solubilization buffer  with agitation for 2 h at 4°C. After centrifugation at 39,000 rpm for 30 min to remove undissolved debris, the supernatants were then loaded onto a GSTrap FF column  preequilibrated with buffer A . After extensive washing in buffer B , the column-bound proteins were eluted in buffer A supplemented with 20 mM glutathione. Fractions containing GST-tagged NS2A proteins were then pooled. Ultrafiltration was performed in a 50-kDa centrifugal concentrator (Sartorius) to remove the glutathione from the eluates. Finally, the eluates were concentrated to about 5 mg/ml, aliquoted, and flash-frozen in liquid nitrogen before storage at −80°C.GST-NS2A, GST-NS2A(58–133), GST-NS2A(58–133) R96A/R102A, and GST-NS2A(58–133) E103A recombinant proteins were expressed and purified from eviously  with mod66-3ʹ UTR RNAs containing the 5ʹ UTR, the first 66-nucleotide sequence of the capsid gene, and the 3ʹ UTR of the ZIKV genome were in vitro transcribed from a DNA template containing T7 promoter using the MEGAscript T7 transcription kit (ThermoFisher Scientific). After transcription, the reaction mixtures were passed twice through the Illustra Microspin G-25 columns  to remove ribonucleoside triphosphates (rNTPs). The RNAs were recovered by phenol-chloroform extraction and ethanol precipitation. Biotinylated 5ʹ UTR-C66-3ʹ UTR RNA (5ʹ UTR-C66-3ʹ UTR-biotin) was prepared by ligating a biotinylated cytidine (bis)phosphate to the 3ʹ end of 5ʹ UTR-C66-3ʹ UTR RNA using the Pierce RNA 3ʹ-end biotinylation kit (ThermoFisher Scientific) according to the manufacturer’s instructions. The biotinylated RNAs were isolated by phenol-chloroform extraction and ethanol precipitation. The biotinylated 5ʹ UTR-C66-3ʹ UTR RNAs were finally resuspended in RNase-free water and stored at −80°C.The 5ʹ UTR-CRNA-protein pulldown assay was performed using the Pierce magnetic RNA-protein pulldown kit (ThermoFisher Scientific) according to the manufacturer’s instructions. Three micrograms of recombinant protein and 3 μg of biotinylated RNA were used in each pulldown reaction. After pulldown, the eluates were loaded onto SDS-PAGE gels. GST-tagged proteins in the eluates were detected by Western blotting using the rabbit anti-GST antibodies.66-3ʹ UTR RNAs was incubated with various concentration of recombinant GST-NS2A proteins in 50 μl binding buffer  for 15 min at 24°C. Afterward, 10 μl 6× gel loading dye without SDS (New England Biolabs Inc.) was added to the reaction mixtures. The mixtures were then loaded onto a 0.4% agarose gel containing 0.5× Tris-borate-EDTA (TBE) buffer to resolve the protein-RNA complexes and free RNAs by electrophoresis (120 V for 30 min) in 1× TBE buffer. Subsequently, RNAs in the agarose gel were transferred to a BrightStar-Plus positively charged nylon membrane (ThermoFisher Scientific) according to the instructions of the NorthernMax-Gly kit (Ambion). The transferred RNAs were then cross-linked to the membrane by irradiation with 200 mJ/cm2 of 254-nm UV light in a CX-2000 UV cross-linker . The biotinylated RNAs were detected using the chemiluminescent detection module (ThermoFisher Scientific). The chemiluminescence signals were detected using the ChemiDoc imaging systems (Bio-Rad).Electrophoretic mobility shift assay (EMSA) was performed to evaluate the binding affinity of GST-NS2A to viral RNAs. In brief, 100 pM biotinylated ZIKV 5ʹ UTR-CdK). The ratio of bound RNAs to the total RNAs (free plus bound RNA) versus the concentration of GST-NS2A proteins was plotted in Prism 8 (GraphPad). The value of dK was calculated using nonlinear regression with the least-squares fitting method, where the Bmax was set as 1. All data show the means and standard deviations (SDs) from three independent experiments.The intensities of the bands representing the bound RNAs (in the protein-RNA complexes) and free RNAs in the EMSAs were quantified using the software ImageJ (NIH) to estimate the equilibrium dissociation constant .All numerical data are presented as the mean ± SD. An unpaired Student