Riboswitches are structured noncoding RNA domains used by many bacteria to monitor the concentrations of their target ligands and regulate gene expression accordingly. In the past 20 years, over 55 distinct classes of natural riboswitches have been discovered that selectively sense small molecules or elemental ions, and thousands more are predicted to exist. Evidence suggests that some riboswitches might be direct descendants from the RNA-based sensors and switches that were likely present in ancient organisms before the evolutionary emergence of proteins. Herein, we provide an overview of the current state of riboswitch research, focusing primarily on the discovery of riboswitches, and speculate on the major challenges facing researchers in the field. UTR) of an mRNA, where it forms at least one ligand-binding aptamer domain whose occupancy dictates the folding of an overlapping expression platform. Ligand-induced alternative folding of the expression platform is exploited to regulate transcription, translation, or other gene expression process NA World ,12 organNA World -15.The structural features and/or ligand specificity of each aptamer domain are used as a basis for organizing riboswitches into classes . To dateAlthough the current list of validated riboswitch classes is small compared to the total predicted, this collection presumably includes the most widespread and abundant examples ,15. Thisnatural Boolean logic gates [The validated riboswitch classes are also likely to showcase ligand binding and gene control characteristics that are widely exploited by those yet to be discovered. Some of the known riboswitches are remarkably simple in both structure and function ,19, wheric gates -25. ThesJudging by the riboswitch finds over the last two decades, we believe it is worth the time and resources required to further explore these types of noncoding RNA domains. As additional classes are discovered, much more is likely to be learned about the full capabilities of RNA, both as chemical sensors for diverse ligands and as genetic switches. These attributes could have been greatly exploited by early forms of life during the RNA World -16. FurtA decade after the first metabolite-sensing riboswitch validation studies were reported -4, two pBefore we discuss details of the known riboswitch classes and the efforts to discover more, it is important to consider how the current state of knowledge compares with what remains unknown. If only a few classes remain to be discovered, then perhaps researchers should move on to new discovery challenges. However, if there indeed are thousands of undiscovered classes, then it seems important to devise approaches that can rapidly shed light on this hidden world of molecular sensors and switches. For the reasons noted below, we believe there are compelling reasons to continue or even increase efforts to discover additional riboswitches and to establish their mechanistic and biological functions.RNA thermometers that evaluate temperatures [When estimating the total number of distinct riboswitch classes in modern cells, one must first consider how these RNAs are being classified . When maeratures . The seqeratures ,16.ykkC motif [However, as described later, some riboswitches have resisted researchers’ attempts to classify them, which means that the current rank-order list of validated riboswitch classes is imperfect. Furthermore, it seems certain that additional riboswitches will be found that call into question the above-defined boundaries between ‘classes’ and ‘types’. For example, a series of variants of the long-standing riboswitch candidate called the kC motif  were inikC motif ,29,38,39Even with these challenges, we believe the current imperfect methods for counting and classifying riboswitches can yield important observations and predictions -17,40-43Although only a tiny sampling of the full diversity of natural riboswitch classes is likely known, several important matters can be evaluated by examining this incomplete list. For example, the expression platforms associated with known riboswitch classes already reveal what mechanisms will commonly be employed by most other riboswitches to regulate genes , as discRegarding this question, the distribution of ligands sensed by the validated riboswitch classes  providese.g., TPP, AdoCbl, SAM, or THF), or electrons in the form of hydride units . These and other enzyme cofactors have been proposed to have emerged in an RNA World [Consistent with the ‘riboswitches are ancient’ hypothesis is the observation that numerous riboswitch classes have been discovered that sense RNA-like compounds . SeveralNA World ,12 Thereribozymes that also selectively bound this fundamental building block of RNA [Also notable are riboswitch classes that sense the building blocks of RNA. For example, a riboswitch class for the nucleotide biosynthetic precursor phosphoribosyl pyrophosphate (PRPP) has been discovered ,46,47. Pk of RNA . Regardlk of RNA ,48,49.+) and coenzyme A (CoA), as well as the universal energy currency of all cells, ATP. Over the last ten years, there has been some progress in shortening this list of conspicuously missing riboswitch classes. A detailed discussion of the ligands sensed by experimentally validated riboswitch classes is presented elsewhere [It is interesting to consider the implications of the list of known ligands sensed by riboswitches, but perhaps just as intriguing are the compounds absent from the list. A decade ago, it seemed surprising that there were some fundamental biomolecules that lacked a validated riboswitch class . These ilsewhere  and, bel+ biosynthesis. One class, called NAD+-I, appears to use two similar aptamers to recognize NAD+, wherein the second aptamer overlaps a ribosome binding site to suppress gene expression when ligand is bound [+, whereas the second aptamer is predicted [+ that does not actually involve direct contact with the nicotinamide moiety. We think it is more likely that the second aptamer will prove to bind the nicotinamide moiety tightly and selectively, rather than use a mechanism to measure the levels of NAD+ without making physical contact with the chemically unique part of the coenzyme. No such controversy is likely to occur for the second NAD+ riboswitch class, called NAD+-II [Most notably, two distinct riboswitch classes ,51 have is bound . Atomistis bound ,53 confiredicted  to bind Currently, there are no validated riboswitch classes for CoA or any of its acylated derivatives, such as acetyl-CoA or succinyl-CoA. One possibility is that riboswitches exist for CoA derivatives but, given that acylated versions varying greatly in fatty acyl chain length exist in cells, there is no single common riboswitch class that monitors the CoA pool. Unless the riboswitch is common, it will be difficult to discover by bioinformatic, genetic, or biochemical search strategies. Alternatively, if many of the common riboswitch classes are direct descendants of ancient RNA devices it is possible that RNA World organisms simply had no need for CoA aptamers. It has been proposed that fatty acid (and therefore phospholipid) metabolism emerged late in the evolutionary progression from the RNA World to today’s organisms . If trueSeveral other fundamental compounds are also missing from the growing list of riboswitch ligands , of whicglmS ribozyme class [Other unexpected gaps in riboswitch sensing also exist, and perhaps most noteworthy are several observations listed below. First, there are other near universal coenzymes, namely pyridoxal phosphate, heme, and lipoic acid, that currently lack validated riboswitch classes . Second,me class -58, whicme class . Similar2O and CO2. Given that water (~55 molar) is the primary solvent for biological systems, there is an ample supply of hydroxyl groups that can provide an oxyanion nucleophile for hydrolysis or hydroxylation reactions. The removal of an oxygen atom during dehydration reactions produces a water molecule that simply adds to the surrounding solvent.Ligands relevant to oxygen management are also notably scarce. Only molybdenum cofactor (MoCo) is counted as a riboswitch ligand whose primary function is relevant to this important task. The predominant role for MoCo-dependent enzymes is to promote oxygen transfer reactions involving redox processes . Perhapsalarmone called ZTP [Although most natural folate derivatives are considered members of a carbon management system, one derivative, 10-formyl-THF (10f-THF) functions as a carrier of an incompletely oxygenated carbon unit. A shortage of this enzyme cofactor leads to the accumulation of the purine biosynthetic intermediate AICAR, which can be further phosphorylated to form the bacterial lled ZTP . ZTP is lled ZTP . Thus, r2 has become a necessity as a recipient of electrons in aerobic organisms that derive energy from reduced electron carriers  via oxidative phosphorylation. However, conspicuously absent on the list of validated riboswitches classes are those for diatomic oxygen carriers such as heme molecules. Perhaps these are scarce because RNA World organisms might have thrived in an era without atmospheric molecular oxygen [The reasons provided above might, at least in part, account for the scarcity of riboswitches directly related to oxygen management. There is a notable absence of riboswitches that either directly bind diatomic oxygen or that can coordinate with this molecule. Or oxygen ,63, obvistructure-activity relationship (SAR) analyses. Instead of conducting months of expensive analog binding assays with each riboswitch aptamer, quality structural models based on x-ray crystallography or NMR data can provide near comprehensive insight into the nature of each newfound ligand binding pocket.Riboswitch research has benefited greatly from the advanced state of RNA structural biology and biophysics. It once seemed possible that the pace of novel riboswitch discoveries would easily be greater than the speed at which atomistic models of their aptamers could be solved by the application of biophysical approaches such as x-ray crystallography or NMR. However, structural models are currently available for nearly all the natural aptamers bound to their ligands ,64-67. Ie.g., Although the structural models of riboswitch aptamers are immensely valuable, they provide only a static image of a portion of a riboswitch when bound to its ligand. However, riboswitches also carry an expression platform and are likely to dynamically fold as they are being synthesized during transcription. The important kinetic parameters -77 of a RNA-seq technologies, where the effects of mutations or the status of individual riboswitches in vitro [in vivo [Single-molecule biophysics studies can be complemented by various in vitro ,87 or ev[in vivo -90 can bOf all the bacterial riboswitch classes that have been experimentally validated over the last two decades, only one class has convincingly been shown to function naturally in eukaryotic species. TPP riboswitches ,3, whichORF) [For example, some fungal TPP riboswitches have been observed to regulate gene expression by retaining or removing an intron located 5′ of the main open reading frame (main ORF) . These iORF) ,96. In aORF) . In planORF) . This TPin vitro selection for RNA aptamers beginning with pools transcribed from natural genomic DNA sequences from eukaryotes was used to identify numerous RNAs that bind adenosine [in vivo structure probing method also was used to identify putative eukaryotic aptamers for the coenzyme FMN [Because many eukaryotic species extensively employ alternative RNA splicing ,103, thedenosine ,105, GTPdenosine ,107, or denosine . An in vzyme FMN . Howeverzyme FMN  and sperzyme FMN  lack sufe.g., signaling molecules and elemental ions) rather than the fundamental metabolites that are so commonly sensed by bacterial riboswitches [Even bioinformatic searches have yet to reveal strong candidates for additional eukaryotic riboswitches. As each new bacterial riboswitch is validated, we typically seek homologs in eukaryotes, but usually without success . Also, uswitches .power law projection for riboswitch abundance [Both conventional thinking and the bundance ,15,17 (Ba) and the ligand it binds (variable b), and the best circumstance is to have high confidence in the answers for these two variables before starting an experimental campaign to prove them. Bioinformatics search algorithms are likely to remain the most effective strategy to identify RNAs that are strong candidates, as well as generate strong clues regarding the precise RNA constructs and the most likely ligand candidates to test. Unfortunately, most other riboswitch discovery strategies proceed without knowledge of either variable, or with knowledge of only one of the two variables. These efforts almost always lead to experimental failures as detailed below for several such methods.Given that the list of natural riboswitches most probably includes many exceedingly rare classes ,15,17 F, it is lEvidence for the existence of riboswitches was first encountered, unknowingly, via the use of genetic analyses conducted by researchers interested in the regulation of specific metabolic pathways. The first reports we can now recognize as hinting at the existence of riboswitches were related to lysine biosynthesis ,116, anddirected evolution methods used to create novel RNA aptamers [SELEX methods [A search strategy like the aptamers ,131 has  methods  use geno methods -108 do f methods ,15,17. Fin vivo or in vitro RNA structure probing methods [in vivo [in vitro [Another single-species approach to discovering natural riboswitch aptamers involves the use of  methods -90. Thes[in vivo  or in viin vitro  but, to transcriptomics data [Yet another single-species method that has been demonstrated for the discovery of novel noncoding RNA domains involves the analysis of ics data ,135. Sucics data ,44,136. A search strategy wherein the researcher arbitrarily chooses both the RNA construct and the candidate ligand perhaps has the lowest chances for success. Usually, the choice of which gene to study is due to the researcher’s interest in how a particular ligand candidate might regulate gene expression. They then identify a gene that logically might be regulated by the chosen compound. Genetic and biochemical assays ensue, and marginal data is sometimes embraced as evidence for riboswitch function. Again, the problem is that the projected number of riboswitch classes is not in favor of success. Imagine the researcher was fortunate to choose a bacterial species that has a single novel riboswitch class associated with one of its ~4,000 genes. Without additional care, the probability that the researcher has correctly chosen to work on the gene regulated by the riboswitch is 1 in 4,000, and the probability that they have also chosen the correct ligand might be only modestly better. Most validated riboswitch ligands are fundamental metabolites or elemental ions, but some ligands were recognized as biologically relevant only after the candidate riboswitch was identified. Thus, it is possible to choose the right riboswitch construct by chance but not have sufficient knowledge to test its matching ligand.Given these highly unfavorable probabilities, one should be very skeptical of riboswitch claims based on this search strategy ,137. Unfcomparative sequence analysis approach [Most methods described above could yield novel riboswitch class discoveries, but they are not the high-probability, scalable methods for the discovery of novel riboswitch classes like those needed to substantially advance the field. In contrast, bioinformatics methods ,138-144 approach ,146 to rapproach -149. MotNatural riboswitches can be exploited in several ways for practical applications, and a full treatment of the latest advances merits a separate review. Herein, we mention only a few major areas in which natural riboswitches can participate in therapeutic and biotechnology advances, and comment on key issues to consider as these technologies mature.e.g., Antibacterial compounds that trigger riboswitch function in a manner that is deleterious to the host cell have been developed -154. Fore.g., Although some compounds developed to trick riboswitches have been tested in animals e.g., -160, theSynthetic biologists have been working to create novel aptamers and RNA switches for use as designer gene control devices for more than two decades -165. Onein vitro or in vivo [Molecular engineers have created various RNA devices that function as switches  in vivo , but app in vivo ,166, but in vivo . Perhaps in vivo ,168, as  in vivo -101 (BoxMany riboswitch classes monitor or regulate such fundamental biochemical pathways that we and others have concluded that they are likely to be of ancient origin -15. ThusEscherichia coli strain carrying a plasmid vector expressing a fluoride riboswitch fused to a β-galactosidase reporter gene was used [E. coli strain carrying a ZTP riboswitch-reporter fusion construct was used to identify compounds from a chemical library that disrupt the folate cycle [Riboswitch-reporter fusion constructs also have been utilized to discover novel compounds that perturb biological processes. For example, an was used  to identte cycle . As the orphan riboswitch candidates [Natural riboswitches sense a remarkable diversity of ligands and thereby help cells monitor biologically relevant chemicals that are of fundamental importance to all forms of life ,16. Thisndidates  and to indidates -141. PerIn addition to finding riboswitch classes that sense more elemental ions , that reallosteric ribozymes, which seem almost certain to have once existed in the RNA World. In contrast, there is no shortage of riboswitches arranged in tandem that mimic the operation of Boolean logic gates [Descriptions of riboswitch mechanism diversity are provided elsewhere ,44,101, ic gates -25. At lWithout enhanced methods, the pace of riboswitch discovery and validation might slow substantially. Traditional laboratory methods are proving inadequate to find more than a few candidates and computational searches will be increasingly frustrated by numerous false positive hits, all due to the rarity of each undiscovered riboswitch class and the abundance of structured noncoding RNAs whose biochemical functions do not involve ligand sensing and gene control. If these problems can be overcome, the next 10 years are likely to reveal many additional surprising structures, functions, and uses of natural riboswitches.


The interaction of RNA with specific RNA-binding proteins (RBP) leads to the establishment of complex regulatory networks through which gene expression is controlled. Careful consideration should be given to the exact environment where a given RNA/RBP interplay occurs, as the functional responses might depend on the type of organism as well as the specific cellular or subcellular contexts. This requisite becomes particularly crucial for the study of long non-coding RNAs (lncRNA), as a consequence of their peculiar tissue-specificity and timely regulated expression. The functional characterization of lncRNAs has traditionally relied on the use of established cell lines that, although useful, are unable to fully recapitulate the complexity of a tissue or organ. Here, we detail an optimized protocol, with comments and tips, to identify the RNA interactome of given RBPs by performing cross-linking immunoprecipitation (CLIP) from mouse embryonal hearts. We tested the efficiency of this protocol on the murine pCharme, a muscle-specific lncRNA interacting with Matrin3 (MATR3) and forming RNA-enriched condensates of biological significance in the nucleus.Key features• The protocol refines previous methods of cardiac extracts preparation to use for CLIP assays.• The protocol allows the quantitative RNA-seq analysis of transcripts interacting with selected proteins.• Depending on the embryonal stage, a high number of hearts can be required as starting material.• The steps are adaptable to other tissues and biochemical assays. Graphical overviewIdentification of RNA/protein interactions from developing heartsThe dynamic interaction between RNA and proteins controls many aspects of gene expression and disease . RNA binding proteins (RBP) have been shown to affect every aspect of RNA metabolism by positively or negatively regulating transcription and splicing, cytoplasmic export, and stability of different classes of RNAs [3]. RBP can also regulate mRNA subcellular localization and promote localized translation [4]. On the other hand, non-coding regulatory sequences in the mRNA, such as 5′ or 3′ untranslated regions (UTR), can reciprocally influence protein fate by controlling protein translation [5]. The synergistic interplay between molecular partners often occurs through the formation of dynamic RNA and protein-containing condensates. In the nucleus, this spatial distribution offers many advantages for the processes of RNA transcription and processing since it concentrates the molecular machinery necessary for gene expression in the three-dimensional space. Notably, the importance of RNA/RBP interactions was shown to be favored by noncoding RNAs. Long non-coding RNAs (lncRNAs), in particular, can guide and influence their protein interactome in both nuclear and cytoplasmic compartments due to their structural versatility and highly context-specific expression [6–12]. Over time, many different approaches have been developed to study RNA–protein interactions [13–14]. These methodologies can be classified into two main types: protein-centric or RNA-centric. The first class is based on the possibility to precipitate a specific protein of interest with an antibody to identify its RNA interactome, while the second class uses modified oligos to precipitate an RNA of interest and then characterize the proteins bound to it. However, several examples in the literature highlight the high rate of false positives that characterize these techniques. Indeed, there is a high possibility to observe artefact interactions between RNA and proteins that can occur in vitro after cell lysis . For this reason, one of the essential advancements made in the identification of in vivo RNA/protein interactions is to catch direct binding between molecules while they are occurring in viable cells thanks to the application of different cross-linking methods. In particular, the UV light wavelength of 254 nm is highly efficient for the cross-linking of RNA and proteins in close contact [17] during a cross-linking immunoprecipitation (CLIP) experiment. Another improvement is in the use of fresh tissues as starting substrate to capture the RNA/protein interactions in a more physiological context. However, since the purification of viable cells from tissues can be challenging, few examples of their use are available [18–20].Here, we present an optimized protocol that, starting from fresh tissue , allows the preparation of in vivo cardiac extracts suitable for UV cross-linking and subsequent immunoprecipitation assays. The described steps allow the maintenance of in vivo RNA/protein interactions, which is an essential prerogative to identify functional molecular partners involved in the regulation of a biological process.Biological materialsC57BL/10 wild-type (WT) mouse ReagentsEthanol Dulbecco’s phosphate buffered saline (PBS) Protein inhibitor complex (PIC) Ribolock Phenylmethylsulfonyl fluoride (PMSF) Dynabeads Tri reagent HEPES KCl EDTA NaF NP40 Tween 20 Dithiothreitol (DTT) Tris–HCl NaCl SDS Proteinase K Laemmli sample buffer Anti-MATR3 antibody Anti-IgG antibody TM RNA MiniPrep Direct-zolSuperScript VILO cDNA Synthesis kit PowerUp SYBR-Green MasterMix Primers:pCharme FW 5′-tttctgtttgccctggacac-3′pCharme RV 5′ - gcactcttccttctctccga- 3′mCharme FW 5′-ggcacagacaccaaggccag-3′mCharme RV 5′ - gcactcttccttctctccga- 3′Gapdh FW 5′-tgacgtgccgcctggagaaa-3′Gapdh RV 5′-agtgtagcccaagatgcccttcag-3′SolutionsPBT (PBS + 0.02% Tween 20)Dissociation media (see Recipes)NP40 lysis buffer (see Recipes)HighSalt NP40 wash buffer (see Recipes)Proteinase K buffer (see Recipes)RecipesDissociation media (1 mL)Note: This volume is intended for 10–12 embryonal hearts. Prepare fresh on the day of the experiment.NP40 lysis buffer (50 mL)Note: Solution can be stored at +4 °C for up to six months. On the day of the experiment, add 0.5 mM DTT (1 M stock), 1× PIC, and 1:200/400 Ribolock to the needed volume .HighSalt NP40 wash buffer (50 mL)Note: Solution can be stored at +4 °C for up to six months.Proteinase K buffer (50 mL)Note: Solution can be stored at room temperature for up to six months. On the day of the experiment, add 0.5 mM DTT (1 M stock), 1× PIC, and 1:200/400 Ribolock to the needed volume.Laboratory supplies1.5 mL tube 15 mL tube Petri dish 100 mm, tissue culture treated Cell strainer 70 μm Cell lifter Surgical scissors Jewelers’ forceps, Dumont No. 5, L 4 1/4 Tweezer Micropestle Microscope UV-Crosslinker Magnetic rack Rotating wheel Lab roller Sonicator Thermomixer (Eppendorf)This protocol is suitable for the preparation of cellular extract from UV-crosslinked cells isolated from embryonal hearts. Cardiac tissue can be highly heterogeneous and composed of different cell types, such as cardiomyocytes, cardiac fibroblasts, and endothelial cells. Due to their size (lower than strainer cutoff), all these cells are kept during the filtering step, which is mainly required to remove clumps and tissue debris. To maintain in vivo interactions, we recommend the use of freshly collected hearts as cells need to be cross-linked while they are still viable. The cardiac extracts can be used as input for CLIP assay. An IgG negative control should always be run in parallel to the specific IP to check the specificity of the antibody (see sections D–G and [11]). It is important to note that at least 1 mg of total extract per condition (IP-specific and IgG negative control) is necessary. In Taliani et al. (2023) [11], a total of ~60 hearts (E15.5) were collected for CLIP and yielded ~5.3 mg of protein extract (0.09 mg for each E15.5 heart).Embryonal hearts isolation2 or cervical dislocation as approved by the Institutional Animal Use and Care Committee.Sacrifice the pregnant mouse by COWet the skin and the fur of the mouse with 70% ethanol to avoid samples contamination with mice hair.Position the mouse under the hood. Pinch the skin with tweezers, pull up and incise and pull apart the skin to expose the abdomen. Cut the peritoneum: you will see all the embryos contained in the placenta , do not store more than 5–6 hearts in a single 1.5 mL tube.Manual dissociationAttention:Prepare 1 mL of dissociation media. This volume is intended for 10–12 E15.5 embryonal hearts.Carefully remove as much PBS as possible to let the hearts settle down to the bottom of the tube , left.Add 500 μL of cold dissociation media to each tube (5–6 hearts).Mash the hearts with a pestle for 2–4 min on ice.Further dissociate the tissue by pipetting the solution with a 1,000 μL tip until no big clumps of tissue can be observed , right.Tip: For a faster dissociation, trap the hearts between the pestle and the edge of the 1.5 mL tube . It is pCardiac cells preparationPlace the 70 μm strainer  on a 15 mL tube on ice.Slowly add the cardiac homogenates to the strainer with a P1000 pipette. The solution will be filtered with gravity .Wash the strainer with PBS until 5 mL of volume is reached in total.Use a cell lifter to facilitate the filtration process by gently scraping the top of the strainer. Be careful not to break the filter membrane .Pour the 5 mL of filtered cardiac homogenates in one Petri dish (100 mm of diameter) at room temperature and check cells’ viability.Tip: Slightly move the plate, and consequently the media, to better visualize the translucent cells under a brightfield light microscope. Cells should stay in suspension . To visuExtract preparation2 on ice.Remove the plate lid and UV-crosslink the cells in a Spectrolinker UV Crosslinker at 254 nM with 4,000 μJ/cmHarvest the cells using a cell strainer and transfer the cell suspension to a 15 mL tube.g for 5 min at 4 °C.Centrifuge at 600× Attention: To proceed with extract preparation, scale up the number of hearts and repeat sections A–B.Remove the supernatant very gently to avoid disturbing the cell pellet and snap-freeze the cell pellet on liquid nitrogen. Pellets can be stored at -80 °C for up to 12 months. Resuspend the frozen pellets in 3 mL of NP40 lysis buffer.Pipette up and down with a P1000 pipette until the solution becomes homogeneous. Split the volume evenly in three 1.5 mL tubes .Incubate the tubes for 10 min at 4 °C with gentle shaking.Perform six cycles of sonication at low intensity  for 30 s at 4 °C with a Bioruptor Plus sonication device to ensure nuclear membrane lysis.g for 5 min at 4 °C and collect the supernatant, which represents the total cellular extract. Attention: After centrifugation, the supernatant should be clear. If the sample is still turbid, perform an additional centrifugation  and repeat steps D5–D9 using a lower quantity of NP40 lysis buffer (1 mL instead of 3 mL).Centrifuge at 16,000× Quantify the protein extract concentration with Bradford assay. Then, dilute the sample in supplemented NP40 lysis buffer to adjust the final concentration to 1 mg/mL.Beads loadingAttention:Prepare 1 mL of PBT supplemented with 1× PIC and 1:200/400 Ribolock. This volume is intended for the 2 h incubation of two reactions (IP and IgG). The washing volume is not included.For each immunoprecipitation, use two 1.5 mL tubes labeled IP (protein-specific) and IgG (negative control). Gently mix the Dynabeads Protein G magnetic particles and add 30 μL of beads to each tube.Wash the beads by adding 1 mL of PBT per tube.Place the tubes on the magnetic rack. Let the beads settle towards the magnetic rack and carefully remove all the supernatant.Repeat steps E2–E3. Carefully, aspirate the last PBT wash without touching the beads.Remove tubes from the magnetic rack and resuspend the beads in 400 μL of PBT by pipetting.Attention: Use the same Ab isotype for IP and IgG in order to use the same beads in the following section (F).Add the same amount of IP-specific or IgG antibodies to the corresponding tubes. Use antibody (Ab) amounts recommended by the manufacturer . Incubate at room temperature on a rotating wheel for 2 h.Immunoprecipitation (IP)Wash the IP-specific and IgG-loaded beads twice at room temperature with 1 mL of PBT. After each wash, place the tubes on a magnetic rack, let the beads settle, and carefully remove all the supernatant.Take 10% volume of extract as INPUT (INP) and keep on ice.On ice, mix 1 mL (1 mg) of the extract with the IP-loaded beads and 1 mL of the extract with the IgG-loaded beads.Incubate at 4 °C overnight on a rotating wheel.The day after, wash the beads three times with 1 mL of HighSalt NP40 wash buffer.Resuspend the beads in 100 μL of NP40 lysis buffer.Bring the INP sample to 100 μL of final volume with NP40 lysis buffer.Attention: It is possible to split the 100 μL in different combinations. In Taliani et al. (2023) [11], a 3:1 v/v ratio was used for RNA (75 μL) and protein (25 μL) extraction.For each sample , split the volume in two tubes for RNA and protein extraction. RNA preparationAdd 125 μL of proteinase K buffer and 50 μL of proteinase K enzyme to the 75 μL of resuspended beads (RNA tube) to reach a final volume of 250 μL.Incubate at 50 °C for 30 min with gentle shaking.Spin the tubes and prepare the sample for RNA extraction by adding 3–5 volumes of Tri reagent.Attention: Make sure that no residue of beads is left before proceeding. Bead contaminations can interfere with RNA extraction.Let the beads settle toward a magnetic rack and transfer the supernatant to a new tube. Repeat twice. TM RNA MiniPrep kit. Other methods can also be used . INP usually yields ~1 μg of RNA in a total of 30 μL of RNase/DNase free H2O.Extract and purify the total RNA to use in gene expression analyses (see section I). In Taliani et al. (2023) [11], the RNA was extracted using the Direct-zolProtein preparationAdd 4× Laemmli sample buffer (10 μL) and 50 mM DTT  to 25 μL of resuspended beads (protein tube) to reach a final volume of ~40 μL.Heat at 95 °C for 5 min.Let the beads settle toward a magnetic rack and transfer the supernatant to a new tube.Store at -80 °C until western blot analysis (see section I).CLIP experiment analysisCheck the efficacy of the target protein immunoprecipitation by western blot assay. Load half (20 μL) of the volume of each sample  into the SDS-PAGE gel. Proceed with protein separation by gel electrophoresis, transfer, and immunodetection. Western blot analysis for MATR3 is shown in After confirmation of protein recovery, proceed with RNA analysis by RT-qPCR or RNA-sequencing. In Taliani et al. (2023) [11], the RNA samples were retrotranscribed using SuperScript VILO cDNA Synthesis kit and quantified by RT-PCR using PowerUp SYBR-Green MasterMix ; melt curve: 15 s at 95 °C, 1 min at 60 °C, 30 s at 95 °C, 15 s at 60 °C). RNA was then sequenced on an Illumina Novaseq 6000 Sequencing system.Tip: Check the outcome of CLIP by performing RT-qPCR analysis on RNA transcripts used as controls. RNA samples from CLIP assays are suitable for both RT-qPCR and RNA-sequencing. For RT-qPCR, the enrichment of target RNAs can be graphed as the ratio of IP over INP. In the case of RNA sequencing, a detailed description of the analysis is presented in the Material and Methods section of Taliani et al. (2023) [11] . For the identification of reliable RBP RNA interactors, at least two biological replicates must be processed.This protocol (or parts of it) was used and validated in the following research article(s):Taliani et al. (2023) [11]. The long noncoding RNA Charme supervises cardiomyocyte maturation by controlling cell differentiation programs in the developing heart. Elife . Other tissues can also be used depending on the efficacy of viable cell purification.TroubleshootingProblem 1: The quantity of cells obtained from the hearts is lowPossible cause: Low starting materialSolution: Prepare and freeze multiple cell pelletsProblem 2: The solution does not flow through the 70 μm strainerPossible cause: The remaining cell clumps in the sample clog the strainerSolution: Recover the solution from the strainer, use a micropipette to further disrupt cell clumps, and add PBS to dilute the sample. Load the solution in the strainer, keeping the micropipette perpendicular to the strainer membrane, and apply some pressure on the filter without breaking the membrane.


Circular RNAs (circRNAs), a circular form of cognate RNA generated by back-splicing, are abundant in the brain and accumulate with age. Hence, they are assumed to play a role in neurodegeneration. Emerging evidence has demonstrated that age-related dysregulation of RNA editing and changes in circRNA expression are involved in ALS pathogenesis. Herein, we review the potential associations between age-dependent changes in circRNAs and RNA editing, and discuss the possibility of developing new therapies and biomarkers for ALS based on age-related changes in circRNAs and dysregulation of RNA editing.Amyotrophic lateral sclerosis (ALS) is an incurable motor neuron disease caused by upper and lower motor neuron death. Despite advances in our understanding of ALS pathogenesis, effective treatment for this fatal disease remains elusive. As aging is a major risk factor for ALS, age-related molecular changes may provide clues for the development of new therapeutic strategies. Dysregulation of age-dependent RNA metabolism plays a pivotal role in the pathogenesis of ALS. In addition, failure of RNA editing at the glutamine/arginine (Q/R) site of GluA2 mRNA causes excitotoxicity due to excessive Ca TARDBP), fused in sarcoma (FUS), and chromosome 9 open reading frame 72 (C9ORF72) quinoxaline-7-sulfonamide disodium salt (NBQX) and perampanel, and AMPA receptor-specific RNA aptamers have been shown to protect against the motor neuron death of ALS model animals ,176,177.ant mice , and theant mice ,176. Howant mice . Since nant mice ,180 and del mice  but has del mice . Mexiletdel mice ,184, butdel mice . Ezogabidel mice ,187. TheAging, circRNAs, and dysregulation of RNA editing are closely associated with one another. As aging is a major risk factor for neurodegenerative diseases, including ALS, scrutiny of the critical roles of age-dependent circRNAs and dysregulation of RNA editing in the pathogenesis of ALS is required to establish new disease-specific therapeutic approaches and biomarkers for ALS as well as for brain aging.


Ziziphus jujuba var. spinosa  and its phylogenomic relationship with Chinese jujube were investigated. Plastomes of 21 sour jujube individuals were sequenced and assembled. The length of the sour jujube plastomes ranged between 159399 and 161279 bp. The plastomes exhibited collinearity of structure, gene order and content. The most divergent regions were located in the intergenic spacers, such as trnR-UCU-atpA and psbZ-trnG-UCC. Sliding window analysis demonstrated that the sequence variation among the sour jujube plastomes was relatively low. Sixty-two to 76 SSRs with 4 motif types were identified in the sour jujube plastomes with a predominant motif type of A/T. Three protein-coding genes exhibited higher nonsynonymous/synonymous substitution ratios, indicating that these genes may undergo positive selection. A total of 80 SNPs were detected and 1266 potential RNA editing sites of 23 protein-coding genes were predicted. In the phylogenomic tree constructed, sour jujube has a sister relationship to Chinese jujube, which indicates that Chinese jujube may have originated or been domesticated from sour jujube. The present study explicitly investigated the individual-level plastome variation of sour jujube and provides potential valuable molecular markers for future genetic-related study of this lineage.Comparative plastomics can be used to explicitly dissect various types of plastome variation. In the present study, the plastome variation pattern of  The chloroplast is a plant organelle originating from a primitive cyanobacterium and performs a fundamental role in plant photosynthesis and other functions . The conZiziphus jujuba var. spinosa, also known as sour jujube, is a deciduous shrub plant species belonging to the Rhamnaceae and is widely distributed in Northern China (Z. jujuba var. spinosa is classified as a varietas of Z. jujuba (Chinese jujube). However, based on its morphology and other evidence, some researchers have suggested that sour jujube should be classified as an original lineage and Chinese jujube might have originated or been domesticated from sour jujube . Sequence libraries were quantified using a Bioanalyzer 2100 . Each sample was barcoded with a unique index and libraries were pooled, and whole genome sequence data were generated using an Illumina HiSeq 2500 platform.Z. jujuba var. spinosa (KX266830) as a reference, the plastomes of 21 individuals sampled in this study were assembled using NOVOPlasty 3.094 (https://www.geneious.com). The circular plastomic maps of the 21 individuals were drawn using OGDRAW with default values  as outgroups based on the 73 common protein coding sequences extracted by Geneious 10.2.2 and aligned with MAFFT. All the other species’ plastomes were directly downloaded from NCBI  was located at another copy of ycf1 and was adjacent to ndhF. A little inconsistent deviation of JSB from ndhF was observed. The junction of LSC/IRb (JLB) spanned rps19 in all Ziziphus plastomes, and was proposed to be a putative ancestral JLB state in angiosperm species  were detected throughout the 21 aligned sour jujube plastomes . More work is needed to understand how these changes may lead to functional changes between different populations (While the plastome is generally viewed as highly conserved and little changed, other findings have suggested that the plastome contains ample SNPs for resolving various issues (Ziziphus acidojujuba C. Y. Cheng et M. J. Liu, of sour jujube may not be appropriate from the perspective of plastome phylogeny. Z. jujuba may have originated or been domesticated from Z. jujuba var. spinosa (The classification and delimitation of intra-Rhamnaceae lineages are controversial, resulting from the large number of species, highly variable morphology as well as discrepancy between different data types from various studies , but a mZ. jujuba var. spinosa were assembled and comparative plastome and phylogenomic analyses were conducted. The conservative nature of the plant plastome was detected in the sour jujube plastomes, but some intriguing variations were also uncovered, such as some highly divergent loci and functional genes under positive selection. The CpSSRs and SNPs identified in sour jujube plastomes provide informative molecular markers for future studies. Phylogenomic analysis indicated the sister relationship of sour jujube to Chinese jujube. The plastome contains sufficient information about variation and has been shown to be effective in plant biology research (In the present study, plastomes of plad006_suppl_Supplementary_Table_S1-S5_S7-S9Click here for additional data file.plad006_suppl_Supplementary_Table_S10Click here for additional data file.plad006_suppl_Supplementary_Table_S6Click here for additional data file.


Fabaceae species, BBWV2 RNA1 NCBI accession KC790225, and RNA 2 MW556592 both from China. This is the first BBWV2 genome study reported in Australia and forms part of the strategy to integrate versatile diagnostic genomics tools at the border to safeguard the Australian grains industry.High-throughput sequencing (HTS) is an important tool for plant virus detection and discovery. HTS technologies such as Nanopore sequencing has been rapidly developing in recent years, and offers new possibilities for fast routine diagnostic applications. This study compared MiSeq (Illumina) RNA-Seq with the MinION sequencer (Oxford Nanopore Technologies) (ONT) direct-RNA-Seq methods to detect and sequence the broad bean wilt virus 2 (BBWV2) genome. The Illumina BBWV2 RNA1 and RNA2 genome segments were intact and matched with ONT with 99.1% and 98.8% nucleotide identity (nt) match, respectively. The RNA1 genome derived from ONT had deletions within the nucleoside-triphosphate binding and protease factor open-reading frames, and the upstream of five untranslated regions. However, its RNA2 was intact, and no sequencing errors were observed. The ONT and Illumina RNA1 and RNA 2 BBWV2 genome segments clustered together phylogenetically along with other none-Globally, viral diseases impair the growth and vigor of cultivated crops such as grains, leading to a significant reduction in quality, marketability, and competitiveness. As an island nation, Australia has a distinct advantage in using its border to prevent the introduction of damaging viruses, which threaten the continental agricultural sector. However, breeding programs in Australia rely on imported seeds as new sources of genetic diversity. As such, it is critical to remain vigilant in identifying new and emerging viral pathogens, by ensuring the availability of accurate genomic diagnostic tools at the grain biosecurity border. High-throughput sequencing offers game-changing opportunities in biosecurity routine testing. Genomic results are more accurate and informative compared to traditional molecular methods or biological indexing. The present work contributes to strengthening accurate phytosanitary screening, to safeguard the Australian grains industry, and expedite germplasm release to the end users. Integrated diagnostic technologies such as high-throughput sequencing (HTS) offer immense opportunities in plant viruses disease management strategies. HTS approaches, such as RNA-Seq Illumina short-read sequencing, have been widely used to detect and discover RNA viruses –4. HowThe direct RNA long-read sequencing capability has proven to be robust in unravelling cell transcriptional features . It follFabavirus of the subfamily Comovirinae, family Secoviridae and is an important pathogen causing damage in many economically important horticultural and ornamental crops worldwide (Broad bean wilt virus 2 (BBWV-2) belongs to the genus orldwide . The BBWorldwide . The RNAorldwide . Due to orldwide , this viVicia faba (faba bean) leaf sample “166_NSW” collected in 2001 from New South Wales, Australia, and preserved with CaCl2 was extracted using the RNeasy Plant Mini Kit (Qiagen), according to manufacturer’s instructions. The total RNA integrity was verified and used to prepare Illumina RNA-Seq library followed by sequencing  with average sequence depth of 92,061. The BBWV-2 RNA2 had a total of 3,711,210 reads mapping to it and yielded a 3,594-nt contig that aligned to 89.33% nt of BBWV-2 RNA 2 (MW556592) with a sequence depth of 73,085. The ONT direct RNA sequencing generated 1,380,000 raw reads, and 1,191,358 reads passed quality control. The ONT had a total of 75,982 reads mapping to BBWV-2 RNA1 and yielded a 5,549-nt contig that aligned to 92.6% nt of BBWV-2 RNA1 (KC790225) with a sequence depth of 2,609. The ONT BBWV-2 RNA2 had 61,847 reads mapping to it and yielded a 3,605-nt contig length that aligned to 88.0% nt of BBWV-2 RNA 2 (MW556592), with a sequence depth of 2,318.Fabaceae BBWV isolate from China  with 94.1% nt identity and BBWV2 RNA2 matched 89.3% to NCBI (MW556592). The ONT-derived 5,549 nt BBMV-2 RNA1 contig contained deletions within the NTP binding, protease factor ORFs, and the upstream of five untranslated regions (UTRs). However, the BBWV2 RNA2 3,605 nt contig generated from ONT was intact, and no sequencing errors were observed when compared to the reference sequence MW556592 and the Illumina BBWV-2 RNA2-derived genome segment. The Illumina and ONT hybrid assembled contigs for the 166_NSW BBWV-2 RNA1 and RNA2 genome segments phylogenetically grouped together and most closely aligned with the RNA 1 (KC790225) and RNA 2 (MW556592) of a none-om China .The ONT direct RNA long-read sequencing offers an attractive option for detecting RNA viruses within a routine diagnostic framework due to native generation of long reads . Also, i


Moreover, we uncovered a dichotomy of NAD-RNAs in translation where they are detected with mitochondrial ribosomes but minimally on cytoplasmic ribosomes indicating their propensity to be translated in mitochondria.Accurate identification of NAD-capped RNAs is essential for delineating their generation and biological function. Previous transcriptome-wide methods used to classify NAD-capped RNAs in eukaryotes contain inherent limitations that have hindered the accurate identification of NAD caps from eukaryotic RNAs. In this study, we introduce two orthogonal methods to identify NAD-capped RNAs more precisely. The first, NADcapPro, uses copper-free click chemistry and the second is an intramolecular ligation-based RNA circularization, circNC. Together, these methods resolve the limitations of previous methods and allowed us to discover unforeseen features of NAD-capped RNAs in budding yeast. Contrary to previous reports, we find that 1) cellular NAD-RNAs can be full-length and polyadenylated transcripts, 2) transcription start sites for NAD-capped and canonical m Two methods NADcapPro and circNC are presented for precise identification of NAD-caps and non-canonical caps in eukaryotic RNAs. Historically only eukaryotic transcripts were believed to contain 5ʹ end modifications since the original analyses of bacterial RNA composition did not detect modification on the 5′ end3. However, seminal work in the late 1970s4 and early 20005 established that nucleotide metabolites, by virtue of their adenosine nucleotide moieties (NAD and FAD), could be utilized by E. coli and T7 RNA polymerases as initiating nucleotides for RNA synthesis in vitro. The initial evidence that NAD caps are indeed incorporated into RNA in cells was provided by mass spectrometry (LC-MS) in bacteria6. Nonetheless, due to the inability of MS analysis to provide the sequence context, the physiological role of these noncanonical caps remained elusive.The presence of modified nucleosides is a characteristic feature of the majority of cellular RNAs. Modified nucleosides generally ascend from the chemical modification of a genetically encoded nucleoside. One exception is the addition of an N7-methylguanosine to the 5ʹ end of select eukaryotic RNAs transcribed by RNA polymerase II referred to as “m7. This method not only corroborated the presence of NAD-capped RNAs (hereafter referred to as NAD-RNA) in bacteria but also divulged the identity of the transcripts. NADcapSeq transpired to be a decisive advance in assessing the function of these caps in RNA metabolism. Subsequent NADcapSeq analyses of yeast8, plant9, human10, and archaeal RNA12 have led to the identification of a significant number of transcripts with a NAD cap and have established the presence of the NAD cap as a 5′ non-canonical cap in all three domains of life.To assess the significance of the presence of the NAD cap in cellular RNA, a chemo-enzymatic and Next Generation Sequencing (NGS) based method, known as NAD captureSeq (NADcapSeq), was introduced in 2015Aplysia californica to replace nicotinamide with alkynyl alcohol, in a transglycosylation reaction13. The ‘clickable’ transglycosylation product is next biotinylated by a copper-catalyzed azide-alkyne cycloaddition (CuAAC)7. After enriching the biotin-linked RNAs using streptavidin beads, the RNAs are subjected to NGS-based RNA Seq analysis to identify the NAD-RNAs. Although NADcapSeq is a robust method for cataloging NAD-RNAs, recent studies have highlighted two of its key limitations; (a) the dependence on the use of copper ions as a metal catalyst for the Huisgen cycloaddition reaction14, and (b) ADPRC promiscuity16. Metal ions, including copper, can induce RNA fragmentation, which could hinder the detection of low-abundant RNAs, and manifest a bias toward shorter fragmented reads from the 5′ termini of relatively abundant RNA17. A further limitation of profiling NAD-capped RNAs by NADcapSeq in eukaryotes involves the residual promiscuity of the ADPRC enzyme on the m7G-capped RNAs16 (hereafter referred to as m7G-RNA). Depletion of m7G-RNA by an anti-m7G cap antibody has been used as one strategy to minimize the level of m7G-RNA in the NAD-RNA population16. Although this is an improvement, additional approaches to streamline NAD cap detection are indispensable for the rapid and accurate profiling of the NAD-RNAs in eukaryotes. Furthermore, the current lack of high-resolution mapping of the NAD position in eukaryotes precludes mechanistic determination of NAD cap addition.NADcapSeq leverages the unique feature of Adenosine Diphosphate Ribosyl Cyclase (ADPRC) from 14 to identify NAD-capped transcripts in Arabidopsis16, which depended on m7G cap antibody depletion to minimize contaminating m7G-RNAs. NADcapPro Seq circumvents the antibody depletion by coupling the robust m7G decapping activity of Dcs118 prior to the SPAAC reaction. Dcs1 abolishes the unfavorable residual enzymatic activity of ADPRC towards canonical m7G cap. Moreover, direct visualization of the NAD-capped RNAs was possible with streptavidin-conjugated IRDye. A second method, circNC, provides an independent mechanism to detect non-canonical capped RNAs at the nucleotide level of precision. Analysis of NAD-capped RNAs initially detected by NADcapPro subjected to circNC enabled us to extrapolate that NAD caps are not only incorporated as noncanonical initiating nucleotides in place of ATP but, can also be post-transcriptionally added in eukaryotic cells. Furthermore, NAD caps were found to be associated with mitochondrial ribosomes, raising the possibility that they can be translated in mitochondria.Here we report two methods, NAD cap profiling (NADcapPro) Seq, and intramolecular circularization of Noncanonical-Capped (circNC) RNA. NADcapPro Seq builds on the recently reported use of copper-free Strain-Promoted Azide-Alkyne Cycloaddition (SPAAC)Huisgen cycloaddition reaction14. An alternative approach based on copper-free chemistry, the strain-promoted azide-alkyne cycloaddition (SPAAC) reaction19 was recently reported16. Instead of using 4-pentyn-1-ol (NAD capSeq), 3-azido-1-propanol is used for the ADPRC-catalyzed transglycosylation reaction  to obviate the need of using copper ions  RNAs were subjected to the SPAAC reaction and the biotin-conjugated RNA (corresponding to NAD-RNA); was visualized by near-infrared (IR) IRDye® 800CW streptavidin-based detection. As observed in Fig. 10. Moreover, the sensitivity of IR dye for the detection of NAD-RNAs is in the fmol range. SPAAC reactions with different concentrations of in vitro transcribed NAD-RNAs demonstrated that biotin-conjugated RNAs can be detected by the IR dye at levels as low as 15.6 fmol  base21). Consistent with previous reports16, detection by IR dye of biotinylated RNA analyzed by SPAAC revealed residual activity on m7GpppA capped RNAs  and a 7-methyl-guanosine monophosphate18 prior to SPAAC modification. We termed this approach NAD cap profiling (NADcapPro).To profile bona fide NAD-RNAs more accurately in eukaryotes, we developed a modified SPAAC approach to enzymatically remove m7GpppA-capped RNA and 40 nts NAD-RNA at molar ratios of ~50:1 was used in the absence or presence of Dcs1 treatment. Visualization of the RNA following the SPAAC reaction was carried out with DBCO-Cy3 to visualize the clicked RNAs, avoiding the prerequisite to blot the RNA onto a membrane. As shown in Fig. 7GpppA-capped RNA in the absence of Dcs1 treatment with robust detection of the m7GpppA-capped RNA. In contrast, treating the RNA mixture with Dsc1 specifically abolished the reactivity of the ADPRC towards the m7GpppA capped RNA without affecting the detection of NAD-RNAs, demonstrating the feasibility of NADcapPro for identifying NAD-RNAs in eukaryotes. This was next validated by using endogenous polyA+ RNAs from WT cells treated with and without Dcs1 prior to the SPAAC reaction 22. Differential expression analyses using DESeq223 identified transcripts enriched in SPAAC or NADcapPro relative to standard RNA-seq. We classified a gene as potentially capped if it was enriched by at least twofold over the standard RNA-seq at a q-value ≤0.01  were identified as NAD-RNAs by NADcapPro  to that of WT cells revealed 346 transcripts that were directly responsive to Xrn1 deNADding  term analysis was carried out to ascertain the potential biological processes the refined list of NAD-RNAs may be involved in Figs. e and 3f.25. Although there is in vitro evidence in support of this hypothesis, cell-based evidence is lacking in eukaryotes. To directly address whether NAD caps can be added by a transcription initiation independent mechanism we developed an approach to map the precise NAD metabolite nucleotide addition site within transcripts validated by NADcapPro to possess NAD caps. We built on two previously reported methodologies, CapZymeSeq26 and RNA end circularization27 to develop an orthogonal approach which we termed intramolecular circularization of Noncanonical-Capped RNA, circNC   Fig.  which isDE) Fig.  or triphDE) Fig. . To veriDE) Fig. . The seq10. However, definitive interpretations of these data were complicated by the fact that an exogenous transcript was used rather than an endogenous NAD-RNA. Support for the translation of NAD-RNAs was subsequently reported in Arabidopsis thaliana where NAD-capped transcripts were detected within cytoplasmic polysomes9. However, as these studies relied on the NADcapSeq approach, it was not clear whether the detected RNAs were indeed NAD-capped as previously reported16.Since nuclear encoded NAD-RNAs have intact 5′ and 3′ UTR and are polyadenylated akin to canonical capped transcripts, we next tested if the NAD transcripts are translated. Initial reports using a transfected NAD-capped RNA in mammalian cells did not detect any appreciable level of translationTo assess whether NAD-RNA can associate with actively translating pools of RNA, we isolated RNAs from polysome gradients and subjected them to NADcapPro Fig. . PolyA+ S.cerevisiae indicated a predominance of NAD caps on mitochondrial encoded transcripts8, and two, Cox2 and 21 S rRNA, were subsequently validated by an independent method24. Yeast mitochondria encode 8 major protein-coding genes – Cox1, Cox2, Cox3, COB, SCE1, ATP6, ATP9, and VAR1 along with two mitochondrial ribosomal RNAs -15 S and 21 S rRNA, and 24 tRNAs. Since mitochondrial transcripts lack a polyA tail, they were not included in our above NADcapPro analysis. We next set out to establish a comprehensive NAD-RNA profile of protein-coding and rRNA yeast mitochondrial transcripts by SPAAC analysis of total RNA isolated from purified mitochondria and assessed NAD transcripts by Northern blotting using the RNA eluates from the affinity purified NAD-RNAs. The lack of m7G caps on mitochondrial transcripts precluded the need to employ NADcapPro. Only 5 NAD mitochondrially encoded transcripts were detected to be NAD-capped  in more detail. This method renders direct visualization of both NAD-capped and uncapped RNA populations by specifically impeding the mobility of NAD-RNAs in the gel due to the transient formation of diesters between immobilized boronic acid and the ribose moiety30. DNAzyme-mediated RNA cleavage was used to generate 5′-end containing sub-fragments of defined length22, and a Northern blot analysis with the 32P labeled transcript-specific probes was used to corroborate the presence of NAD-capped transcripts. Remarkably, unlike nuclear-encoded transcripts, both ATP9 and Cox2 NAD mRNAs are associated with the translationally active mitochondrial ribosomes  is detected in the mature ribosome, but not the 21 S rRNA  that favored 5′ RNA fragments. However, more pertinently, its extrapolation into analyzing eukaryotic mRNAs with m7G caps was problematic due to the promiscuity of the ADPRC reaction. To address this issue, Hu et al16 reported the use of copper-free SPAAC-mediated NAD-RNA isolation and proposed the use of m7G-capped RNA depletion to enrich for NAD-RNA. Herein we expanded on the use of copper-free SPAAC-mediated NAD-RNA isolation and coupled the SPAAC reaction with a more robust enzymatic approach to eliminate m7G-capped RNAs to accurately profile NAD-RNAs in eukaryotes and further leveraged the highly sensitive IRDye® 800CW streptavidin-based detection to visualize NAD-RNAs at concentrations as low as 15 fmol. Of note, a recent report also used an enzymatic approach to minimize m7G capped RNA within the NAD population by eluting biotin-conjugated NAD-RNAs with a NAD decapping enzyme31.The presence of NAD as a 5′ noncanonical cap in cellular RNAs has been established in various organisms7GpppA-caps in eukaryotes by exploiting the robust m7G decapping activity of yeast Dcs118. Dcs1 treatment of RNA prior to the ADPRC reaction eliminates ADPRC reactivity towards the m7GpppA-capped RNAs  whose 5′ ends are generated by exoribonucleases and cannot undergo ab initio capping10. In the present study, we expand on these results and demonstrate in addition to snoRNAs, cytoplasmic mRNA can also be NAD-capped post-transcriptionally suggesting the presence of dedicated capping machinery at least in S.cerevisiae. The demonstration of both human and plant snoRNAs with NAD caps33 also suggests that post-transcriptional NAD capping is likely a general mechanism in eukaryotes. Our data is consistent with two modes of NAD cap addition, one being ab initio addition and the second through a yet uncharacterized NAD capping mechanism. Collectively, our findings reveal the complex nature of cellular NAD capping and the limitation of our current understanding of this intriguing modification.The best-characterized mechanism of NAD cap addition currently is the ab initio addition of non-canonical caps including NAD caps. This is primarily based on in vitro studies demonstrating that the RNA polymerases from both prokaryotes and eukaryotes can add a NAD cap in place of ATP at the 5′-end during transcription initiation10 we demonstrated that nuclear-encoded NAD-RNAs minimally interact with a translationally competitive pool of cytoplasmic ribosomes and do not appear to be engaged in translation in yeast cells under normal growth conditions. Remarkably, a more dramatic outcome was detected in mitochondria. Mitochondrial protein-coding NAD-RNAs were detected to be engaged within the translationally competent pool of mitochondrial ribosomes, consistent with undergoing active translation. One possibility for the differing states of translatability between the cytoplasmic and mitochondrial transcripts could be the requirement of the m7G cap for the cytoplasmic translation1 but not mitochondrial translation34. However, the formal possibility that these NAD transcripts are not engaged in translation but function as regulatory RNAs cannot be ruled out. Future studies will address any potential role of NAD cap in mitochondrial translation.The ability to more accurately identify NAD-RNAs enabled us to assess their presence on actively translating ribosomes and address whether NAD-RNAs can be translated. Consistent with our previous assessment that in vitro generated exogenous NAD-RNAs are not translated22 were grown at 30 °C in . All yeast strains used in the present study are listed in Supplementary Table Yeast cells (BY4741 strain background) WT and xrn1-H41A7G cap structures were synthesized by in vitro transcription from synthetic double-stranded DNA template ɸ2.5-NAD-40, ɸ2.5-NAD-75 containing the T7 ɸ2.5 promoter, and a single adenosine within the transcript positioned at the transcription start site  PPP (5′)-A RNA Cap Structure Analog (New England Biolabs) was included in the transcription reaction, whereas for NAD-RNA, NAD was used instead of ATP. In vitro transcription was carried out at 37 °C overnight, using HiScribeTM T7 High yield RNA Synthesis kit (New England Biolabs (NEB)). Following in vitro transcription, RNA was purified using Monarch®RNA Cleanup Columns (NEB) as per the manufacturer’s instructions.RNAs containing NAD and m2, pH 7.9). Reactions were incubated at 37 °C for 1 hour, and the deNADed RNA was purified using Monarch®RNA Cleanup Columns (NEB) as per the manufacturer’s instructions.For deNADding, the NAD-RNAs were incubated with 100 nM SpRai1 recombinant protein in NEB buffer  which we referred to by the SGD nomenclature, Dcs1, was used. Typically, the reaction was performed in 50 µL volume using 4 units per µg of capped RNA following the manufacturer’s protocol. The decapped RNA was purified using Monarch®RNA Cleanup Columns (NEB) as per the manufacturer’s instructions.For decapping of both in vitro transcribed m35 and treated with DNase (Promega) according to the manufacturer’s protocol. Total RNA from human cells was isolated using TRIzol reagent (Thermo Fisher Scientific) according to the manufacturer’s protocol and treated with DNase as explained above to eliminate DNA. To enrich for poly-adenylated RNA, total RNA purified as above was used as the input for the Poly(A) Purist MAG Kit (Thermo Fisher Scientific) following the manufacturer’s protocol. We retrieved ~1.5 µg of polyA RNA from 100 µg of total yeast RNA.Total RNA from yeast strains was isolated with the acidic hot phenol method2 (pH 7.0), and 40 U of RNasin® Ribonuclease Inhibitor (Promega) at 37 °C for 1 h. After the ADPRC treatment, RNAs were incubated with 250 µM biotin-PEG3-azide, freshly mixed 1 mM CuSO4, 0.5 mM THPTA, 2 mM sodium ascorbate in a 100 µL reaction with 50 mM HEPES, 5 mM MgCl2 (pH 7.0), and 40 U of RNasin® Ribonuclease Inhibitor (Promega) at 30 °C for 30 min. The RNA was then precipitated with ethanol in the presence of 2 mM EDTA and 2 M ammonium acetate36. For reactions using the SPAAC approach, ADPRC was first used to replace the nicotinamide and add an azide residue onto the NAD-RNA. Typically, these reactions were performed in a 100 µL volume with either total RNA or polyA-purified RNA dissolved in DEPC water, 10% (v/v) of 3-azido-1-propanol (Sigma) together with 10 µL of (125 µg/mL) of ADPRC (Sigma) in ADPRC reaction buffer (50 mM HEPES (pH 7.0) and 5 mM MgCl2). The reaction was carried out at 37 °C for 1 hour. The transglycosylated RNA with the azide residues was next purified using Monarch®RNA Cleanup Columns (NEB) and eluted in 20 µL of DEPC water. For the SPAAC reaction, dibenzocyclooctyne-PEG4-biotin  was used as the alkyne moiety for the cycloaddition. Following the ADPRC reaction, both total RNA and polyA enriched RNA in pure water were mixed with 1x volumes of MK-Gel Loading Buffer  and 500 μM DBCO-biotin (Sigma). Typically, these reactions were 20 μL MK-Gel Loading Buffer, 18 μL RNA (from ADPRC reaction), and 2 μL 10 mM stock of the DBCO reagent. The reactions were performed at 55 °C for 15 minutes to denature the RNA and stopped by adding 310 μL water followed by purification of the conjugated RNA using Monarch®RNA Cleanup Columns (NEB). The products were analyzed by gel electrophoresis or subjected to affinity purification as described below.To biotinylate NAD-capped RNAs using CuAAC, 50 µg of total RNA was first incubated with 10 µL of 4-pentyn-1-ol (Sigma-Aldrich), 10 µL of (125 µg/mL) of Adenosine diphosphate-ribosylcyclase (ADPRC) in a 100 µL reaction containing 50 mM HEPES, 5 mM MgClFor blotting analysis of either enriched RNA following biotin-mediated affinity purification or directly after the SPAAC reaction, RNA was incubated at 90 °C for 2 min in formaldehyde loading buffer (FLB) , 0.5 µg/mL ethidium bromide and 10% glycerol). Samples were then loaded onto a 1% agarose gel and resolved by electrophoreses at 120 V for 1 h. Before transferring the gel onto a 0.45 μm nitrocellulose membrane (Cytiva), total RNA was first visualized using a UV gel transilluminator. RNA was transferred overnight by passive, upward transfer facilitated by the capillary flow of the 10X SSC buffer. After transfer, RNA was cross-linked to the NC using UV-C light (0.2 J/cm2). The membrane was next blocked with Odyssey Blocking Buffer, PBS (Li-Cor Biosciences) for 30 minutes at room temperature (RT). After blocking, IRDye® 800CW streptavidin (Li-Cor Biosciences) was diluted to 1:7,000 in Odyssey Blocking Buffer and stained the NC membrane for 30 minutes at RT. The membrane was next washed three times with PBST . Before scanning, the membranes were briefly rinsed in 1x PBS and scanned by an Odyssey Fc (Li-Cor Biosciences) with the software prearranged to auto-detect the signal intensity for both the 600 and 800 channels. The 600 channel was used for ethidium bromide and the 800 channel was used for the detection of IRDye® 800CW streptavidin.7GpppA capped RNA, we mimicked the in vivo differential by mixing in vitro transcribed 40 nts NAD-RNA and 75 nts m7GpppA capped RNA at molar ratios of ~50:1. The mixture was subjected to the SPAAC- reaction as discussed above with the only difference that instead of DBCO-biotin, DBCO-Cy3 was used for cycloaddition. The RNA was purified over the Monarch®RNA Cleanup Columns (NEB). The eluted RNA was next mixed with an equal amount of 2X MK- Gel Loading Buffer and denatured at 65 °C before resolving on a 7 M 15% PAGE gel. The ethidium bromide-stained RNA was first visualized in the gel using a UV gel transilluminator and the Cy3 conjugated RNA was detected using the Cy3 channel on a Typhoon scanner (GE)To assess the residual activity of ADPRC towards the mxrn1-H41 mutant and were treated with Dcs1 as described above to remove m7G-caps. Samples without Dcs1 treatment (SPAAC) and ADPRC (minus ADPRC) were also processed similarly to assess the contribution from m7G caps and any background noise due to highly abundant RNAs. Following the SPAAC reactions, enrichment was achieved by selective affinity to streptavidin beads10. Briefly, RNAs were incubated at 25 °C for 30 min with 15 μL of magnetic Dynabeads™ MyOne™ Streptavidin T1, which were pre-blocked with 100 ng/µL of bacterial small RNAs in 100 μL of immobilization buffer containing 10 mM Tris-HCl (pH 7.5) 1 mM EDTA and 2 M NaCl. After rigorous washing with wash buffer  five times at 25 °C for 5 min each, biotinylated RNAs were eluted by incubating the beads with 20 μL of MK-Gel Loading Buffer at 90 °C for 2 min. The biotinylated RNAs were next purified using Monarch®RNA Cleanup Columns (NEB) as described above and eluted in 10 µL of nuclease-free water. The samples were next flash-frozen in liquid nitrogen and were sequenced at GENEWIZ (Azenta Life Sciences).Fifteen micrograms of polyA RNA were isolated from three independent biological replicates of both yeast WT and 37. Following this, in silico rRNA depletion was accomplished by using hisat238 to align the reads to rRNA sequences and save the reads that did not align. Transcript abundances were quantified by aligning these rRNA-free files to a reference transcriptome  using Kallisto39. The resulting counts were rounded and TPMs recalculated based on these rounded counts. Read count distributions for rounded count can be seen in Supplementary Fig. 40 and the Tidyverse set of packages41. Differential expression was performed using DESeq223 with the “apeglm” method of normalization42. The GO term analysis was performed using Gene Ontology Slim Term Mapper (https://www.yeastgenome.org/goSlimMapper). To determine if the number of overlaps observed in Fig. Raw reads were processed for quality control and adapter removal by fastp®RNA Cleanup Columns (NEB) as described above and eluted in 30 µL of nuclease-free water. The dephosphorylated RNAs are next divided into 3 equal fractions of 10 µL each and treated separately with either no enzyme (Ctrl) or MDE (NEB) or Rai1. The MDE reaction was performed using NEB’s protocol at 37 °C for 1 hour, whereas the Rai1 reaction was performed in 50 µL volume with NEB buffer 3 and 0.5 µL RNAsin at 37 °C for 1 h. The reaction products were next purified using Monarch®RNA Cleanup Columns (NEB) and eluted in 15 µL DEPC water. All Ctrl, MDE, and Rai1 treated RNAs were next subjected to intramolecular ligation using T4 RNA ligase 1 (NEB). Typically, the ligation reaction was performed in a 50 µL volume with 15 µL of Ctrl, MDE, and Rai1 treated RNA following the manufacturer’s protocol in the presence of ~25% PEG8000 at 25 °C for 16 hours. The reaction products were next purified using Monarch®RNA Cleanup Columns (NEB) and eluted in 15 µL DEPC water and reverse transcribed with M-MLV reverse transcriptase and gene-specific primers with gene name -P1 in Fig. In total, 5 µg of yeast polyA RNA was first dephosphorylated using Quick CIP (NEB) to eliminate potential 5′ end phosphorylated RNA contamination in the subsequent reactions. Typically, these reactions were performed in a 100 µL volume with polyA-purified RNA dissolved in DEPC water, with 6 µL of (CIP) in 10X rCutSmart™ Buffer at 37 °C for 40 minutes. The dephosphorylated RNAs were next purified using Monarch37. The analysis of this sequencing data was dependent on our PCR products spanning the junction between the 5′ and 3′ ends of a transcript. We chose primer locations near the start and end of the coding sequences for each gene  individually and used the top 5 most abundant unique reads for multiple sequence alignments (using the R package msa46) which consisted of the top 5 from an MDE sample and from a Rai1 sample. Manual, visual analysis of the junction was carried out on the reads where the junction could be detected.Raw reads were processed for quality control and adapter removal by fastpene Fig. . However600nm~2) were first treated with 100 µg/mL cycloheximide for 10 minutes at room temperature. The cells were next harvested, washed once in ice-cold 1x PBS, and lysed using mortar and pestle in liquid nitrogen. The cell powder obtained after grinding was resuspended in polysome buffer A 47 and incubated at 4 °C for 30 min. Samples were then centrifuged for 10 min at 10,000 × g to remove the cell debris. Approximately 40 A260 units were layered onto 10–50% w/v sucrose gradients prepared in buffer A. The gradients were made with Gradient Master 107 (Biocomp). These gradients were next centrifuged for 3 h at 39,000 rpm and 4 °C in a Beckman-Coulter ultra-centrifuge using the SW40 rotor. After centrifugation, fractions corresponding to each peak  3 M sodium acetate (pH5.2) and 2X volume of 100% ethanol to get rid of sucrose. The ribonucleoprotein pellet was next resuspended in 400 µL of DEPC water and total RNA from each fraction was isolated using Trizol Reagent (Invitrogen) following manufacturers’ instructions. For circNC analysis, polyA RNA from each fraction was isolated as explained above.For polysome fractionation, exponentially grown yeast cells in 500 mL YPD ) and incubated at 4 °C for 30 min. The lysate was next clarified by centrifugation for 10 min at 5000 × g to remove the cell debris. The clarified lysate was then transferred to a fresh 15 mL centrifuge tube and the protein concertation was assessed using a Bradford assay. 5 mg of protein lysate was added to the pre-equilibrated (in 1X lysis buffer) anti-Flag M2 magnetic beads (Sigma). The mixture was rotated end-over-end at 4 °C for 3 h. Next, the beads were washed 3X with wash buffer  for 5 min at 4 °C and the FLAG-tagged protein was eluted by incubation with 200 μg/mL Flag peptide (Sigma). The eluate was next assessed for the presence of MRPS17 by Western Blotting using anti-FLAG antibody (Sigma) . RNA from the eluate was isolated using Phenol-Chloroform as described above.Translationally active mitoribosomes were isolated as described before using C-terminally 3X FLAG-tagged MRPS17 protein as a bait22. 30 μg of total cellular RNA or the RNA retrieved from the mitoribosomes eluates (as described above) were incubated with 1 μM of the corresponding DNAzyme in a 50 μL reaction containing 10 mM Tris-HCl pH = 8.0, 50 mM NaCl, 2 mM DTT, and 10 mM MgCl2. Samples were denatured at 85 °C for 2 min and gradually cooled to 37 °C. MgCl2 was added to a final concentration of 10 mM and incubated for 60 min at 37 °C. Reactions were stopped with 100 μL of stop solution , and RNA was precipitated with 500 μL ethanol by incubating for 30 min at –80 °C followed by centrifugation for 30 min at 16,000 × g at 4 °C. The supernatant was removed, and the pellet was resuspended in H2O.Boronate affinity electrophoresis was performed as described recently32P-labeled DNA probe complementary to the 5′-end fragments of target RNAs (Supplementary Table 32P] ATP (Perkin Elmer). Reaction products were visualized with Amersham Typhoon RGB Biomolecular Imager .NAD-capping was analyzed using DNAzyme-generated fragments of mitochondrial RNAs. The cleaved RNA was resolved on 8% urea polyacrylamide gels supplemented with 0.3% 3-acrylamidophenylboronic acid (Boron Molecular). RNA was next transferred to a positively charged Nylon transfer membrane  and incubated with a https://github.com/shahlab/NADcapPro. Differential expression analysis was conducted as described above using DESeq2 with the “apeglm” as the method of normalization to control for potential biases and improve the accuracy of the results.The present study utilized biological triplicates for all samples in the NAD CapPro Seq and regular RNA seq experiments to ensure statistical robustness and reproducibility. Statistical analysis was conducted using Student’s t-test (single-tail), and chi-squared test to assess the independence of categories in each comparison. The analyses were performed using the Tidyverse package and other general statistics packages of the R programming language version 4.2.1. The code used is available at Further information on research design is available in the Supplementary MaterialDescription of Additional Supplementary FilesSupplementary Data 1Reporting Summary


Nucleic acids extractedfrom biomass have emerged assustainableand environmentally friendly building blocks for the fabrication ofmultifunctional materials. Until recently, the fabrication of biomassnucleic acid-based structures has been facilitated through simplecrosslinking of biomass nucleic acids, which limits the possibilityof material properties engineering. This study presents an approachto convert biomass RNA into an acrylic crosslinker through acyl imidazolechemistry. The number of acrylic moieties on RNA was engineered byvarying the acylation conditions. The resulting RNA crosslinker canundergo radical copolymerization with various acrylic monomers, therebyoffering a versatile route for creating materials with tunable properties. Further, reversible-deactivationradical polymerization methods, such as atom transfer radical polymerization(ATRP) and reversible addition–fragmentation chain transfer(RAFT), were also explored as additional approaches to engineer thehydrogel properties. The study also demonstrated the metallizationof the biomass RNA-based material, thereby offering potential applicationsin enhancing electrical conductivity. Overall, this research expandsthe opportunities in biomass-based biomaterial fabrication, whichallows tailored properties for diverse applications. This is mainly because of therelatively low cost of biomass DNA and RNA compared with other typesof nucleic acids. Additionally, these sustainable natural biopolymerscould be extracted from any living organisms, thereby potentiallyreducing the use of petrochemicals.15 or ethylene glycol diglycidyl ether (EGDE).20 Additionally, electrostatic interactions,22 coordination,24 π–πstacking,27 and groove binding18 havebeen exploited. These noncovalent methods offer the advantage of reshapingand reprocessing the resulting material because of the weak and reversiblenature of the interactions. Furthermore, these biomass DNA-based materialswere metalized via treatment with metal precursors , which enabled their use as catalysts.20 Nevertheless, the simple crosslinking of biomass DNA constrainsthe possibility of engineering material properties since incorporatingfunctional molecules  is challenging.28Until recently, direct crosslinking ofbiomass DNA has been themain approach for fabricating biomass DNA-based material 1, top. I29 Consequently,acyl imidazole chemistry has been extensively studied for the structuralmapping of RNA  in vitro30 andin vivo.31 We have recently demonstratedthat RNA can be modified with acyl imidazole reagents to serve asan atom transfer radical polymerization (ATRP) initiator under themild and biocompatible polymerization conditions.32 Additionally, it is important to note that during the radicalpolymerization process vinyl monomers could undergo copolymerizationwith two or more different types of monomers. As a result, a distinctivematerial emerges with contributing properties from each monomer, whichleads to a tailored material for specific needs.34 Synthetic oligonucleotides may be equipped with polymerizable handles and incorporated into polymericnetworks through radical polymerizations37 or ring-openingmetathesis polymerizations (ROMP).40 However, such methodologyfor nucleic acid modification can only be achieved by leveraging phosphoramiditechemistry during solid-phase synthesis, which is not applicable tobiomass nucleic acids. Consequently, our strategy greatly broadensthe scope of possible functionalities through radical (co)polymerizationmethods for biomass RNA-based 3D-structured materials. This expansionallows access to outstanding potential applicability in various fields,including 3D printing, healthcare, electronics, and catalysis.To address this challenge, we sought to use biomassRNA, whichto the best of our knowledge has not been reported previously. Weaimed to functionalize biomass RNA with polymerizable vinyl groupsrather than simply crosslinking nucleic acids 1, bottomAAm-AI) was synthesized as a model acylating reagent in thereaction of 6-acrylamidohexanoic acid with 1,1′-carbonyldiimidazole(CDI) in DMSO . The freshly synthesized AAm-AI was added to commercially available biomass RNA extractedfrom the torula yeast. The resulting acrylamido RNA crosslinker wasisolated by precipitation with isopropanol. We confirmed through 2Dheteronuclear single-quantum coherence (HSQC) NMR analysis (1H–13C) of the biomass RNA before  and after  AAm-AI treatment that the acrylamide linkers weresuccessfully incorporated in the biomass RNA. The results of the protondiffusion-ordered spectroscopy (1H-DOSY) experiment alsoshowed that the protons in the RNA and the acrylamido moieties haveidentical diffusion coefficients, thereby indicating that they werein the same molecule . We furthercharacterized the biomass RNA crosslinker using size exclusion chromatographyequipped with multiangle light scattering (SEC-MALS) as illustratedin Figure S4 .Acrylamide-functionalized acyl imidazole  to water.29 Therefore, we synthesizedacrylamido RNA crosslinkers under different acylation conditions (%DMSO v/v in water and the amount of AAm-AI), and thedegree of acylation was determined using 1H NMR, accordingto the previously reported procedure .32 As shown in AAm-AI resulted in an increased number of acrylamide incorporation in theRNA. It should be noted that up to superstoichiometric yield (12.8acrylamide groups per 10 ribonucleotides) could be achieved when RNAwas treated with 3 equiv (3×) of AAm-AI comparedwith ribonucleotide in 100% DMSO overnight. The enhanced acylationunder high % DMSO is attributed to the slower hydrolysis of AAm-AI under such conditions . The RNA crosslinkers were further characterized by UV–vis,as shown in AAm-AI reacted with nitrogenous bases in addition to 2′-hydroxylgroups.41 Of note, the most significantshift (∼5 nm) was observed for the 3×, 100% DMSO condition and tetramethylethylenediamine (TEMED)in water. After 5 min of polymerization at room temperature, the RNAhydrogels were stained with GelRed, a nucleic acid-intercalating dye,for visualization (25R100), fabricated using the RNA crosslinker preparedin 25% DMSO (v/v), could not stand independently outside of waterunder the polymerization conditions tested. This can be attributedto the lower number of crosslinking points compared with RNA crosslinkerssynthesized in 100% (100R100)or 50% (50R100) DMSO (v/v) .With the acrylamido RNA crosslinkersin hand, we proceeded to test the capability of acrylamido residuesin RNA to undergo polymerization via free radical polymerization (FRP).Three different RNA crosslinkers were synthesized in 100%, 50%, or25% DMSO in water (v/v) using 3× of SO (v/v) 2B. Enhan42 Inspiredby this, we envisioned that the different degrees of acylation wouldresult in distinct degradation behaviors of the RNA hydrogels. Totest our hypothesis, we examined the biodegradability of the threeRNA hydrogel variants  under biologically relevant conditions as shown in 25R100 and a relatively slowdegradation of 50R100, whichwas evidenced by the diffusion of GelRed to the supernatant. While 25R100 and 50R100 were completely degraded within 24 h, 100R100 remained nearly intact for 18 days ofincubation. The complete degradation of 100R100 was finally observed after incubation for 30 days.These results suggest that the stability and durability of the RNAhydrogel can be engineered by selecting an appropriate RNA crosslinker,which could be useful for the controlled release of cargo loaded inthe hydrogel.44Chemical modifications of nucleic acids can increasethe resistanceof nucleic acids to enzymatic degradation by introducing structuralalterations and steric hindrance.N-isopropylacrylamide(NIPAM) to make RNA-NIPAM hybrid gels  usedin the acylation process, “Comonomer” represents theabbreviation of the comonomer , and y and z indicate the weight percentages of RNA andthe comonomer in the gel, respectively. After the successful copolymerizationof RNA crosslinkers with NIPAM, we stained the hydrogels with GelRed. The degradation of the RNA-NIPAM copolymeric hydrogel in 15% FBSwas also examined . The copolymeric hybrid RNA gels exhibited retarded degradation,possibly attributed to a reduced RNA content within the gel and sterichindrance.To demonstrate the versatilityof our method, we investigated thecopolymerization of the RNA crosslinker with various acrylic monomers4A. RNA cs 2–4 in 4B. To fah GelRed4B by soaM45 2 in 4B and 50M45 3 in 4B was ob25R67AAmix33) was demolded. The resulting copolymeric RNA hydrogels exhibitedsolid structural integrity, thereby enabling them to free-stand andmaintain their shape in the air as a result of successful copolymerizationwith AAmix.We also carried out a reaction in a custom-designedmold by copolymerizingRNA crosslinker using the AAmix as the diluent 4C. The mS12). RNA crosslinkers synthesized in 25% or 100% DMSO (v/v)were copolymerized with 8% AAmix at the final concentrations of 10,30, and 90 mg/mL, respectively. R11AAmix89) showed no significant improvement in the mechanicalproperties .To quantify the RNA crosslinker-assisted enhancementof stiffness,we fabricated cylindrical RNA-acrylamide hybrid gels and conductedmechanical property tests in the compression mode (100R32MA68 (7 in 5 (MA with Irgacure2959) or 6 (MA and unmodified RNA with Irgacure 2959)in Figure S13) and thermogravimetry to confirm the identity ofthe copolymeric material. As shown in 100R100) and copolymeric RNA hybrid materials made with dimethyl acrylamide(100R29DMAAm71) or MA (100R32MA68). The distinct contactangles of the hydrogels imply that the copolymerization of appropriateacrylic monomers with the RNA crosslinker could tune the hydrophobicityof the RNA hydrogels.In additionto NIPAM and acrylamides, the capability to copolymerizehydrophobic monomers with the RNA crosslinker offers the potentialto engineer a wide range of material properties, including enhancedstability in organic solvents and hydrophobic characteristics. Importantly,we observed the high solubility of the RNA crosslinker in DMSO (>200mg/mL), which prompted us to investigate the copolymerization of theRNA crosslinker with hydrophobic monomers in organic solvent . As shown in A68 7 in 4E, wass45 Atom transfer radical polymerization(ATRP) and reversible addition–fragmentation chain transfer(RAFT) polymerization are among the most widely used RDRP techniquesenabling precise control over the properties of synthetic polymers,including molecular weights, molecular weight distributions, composition,and architectures .52 As a result, polymeric networks and multiscale materials made byATRP and RAFT have found various applications in electronics, adhesives,coatings, lubricants, and healthcare.50Reversible deactivationradical polymerization (RDRP) methods are changing the world by enablingthe precise synthesis of polymers and materials in a property-controlledmanner.53 we designed another acylatingreagent with methacrylate functionalities using hydroxyethyl methacrylate(HEMA) and CDI . The resultingHEMA-functionalized imidazole carbamate (HEMA-CM) wastreated to biomass RNA in 50% DMSO in water (v/v) overnight. Characterizationof the methacrylic RNA crosslinker product using UV–vis  and 1H NMR spectroscopy showed slightly reduced functionalizationefficiency of HEMA-CM (3.6 methacrylate groups per 10ribonucleotides) compared with AAm-AI 59 in PBS under green lightirradiation using eosin Y (EYH2) and oligo(ethylene oxide)methyl ether methacrylate  as the photocatalystand the model comonomer, respectively  could also occur on the polymeric RNAnetwork  by FRP and washed the gelby soaking it in water overnight under gentle shaking. The gel wasthen treated with 400 mM AgNO3  measurement results in 3 treatment played a crucialrole in inducing the conductivity of the RNA hydrogel. Moreover, weobserved stronger conductivity when more RNA was used in hydrogelfabrication. These results suggest that the treatment of metal precursorsolutions may provide an additional possibility to engineer the hydrogelproperty in a postsynthetic manner.By harnessing the metal doping onto the RNA hydrogel,we envisionvarious applications ranging from diagnostics,rid gels 6D,E. We ydrogels 6E. We plAAm-AI), which allows for the control over the mechanical properties anddegradability of the network. Increasing the RNA crosslinker concentrationcan also serve as an alternative method for achieving a densely crosslinkednetwork, which results in enhanced stiffness and a lower swellingratio. The resulting acrylamido RNA crosslinker could undergo radicalpolymerization with a diverse range of other acrylic monomers to regulatethe surface polarity of the copolymeric RNA hydrogel while also allowingthe control over the RNA content (0–100 wt %) within the gel.Our study also explored the polymerization of the methacrylic RNAcrosslinker via RDRP techniques , whichenables the fabrication of copolymeric RNA gels with customizableswelling ratios through the choice of the polymerization method. Themetallization of RNA with silver ions demonstrated its potential toenhance the electrical conductivity of biomass RNA-based materials.This work expands the possibilities for biomass RNA-based materialfabrication with tailored properties while overcoming the challengesassociated with conventional synthetic strategies. This techniqueis expected to accelerate advancements in the emerging field of biomassnucleic acid-based materials, thereby enabling diverse applications,such as drug release, electronics, and catalysis.Insummary, we report the successful conversionof biomass RNAinto an acrylic crosslinker using acyl imidazole chemistry. The numberof acrylic moieties on the RNA was engineered by varying the acylationconditions (% DMSO v/v in water and the amount of AAm-AI stock (1 M) was incubatedat room temperature for 30 min under gentle shaking.In 650 μL of DMSO was dissolved 161.2mg (1 mmol) of CDI.To the dissolved CDI in DMSO, 185.2 mg (1 mmol) of 6-acrylamidohexanoicacid was added, and the final volume was brought to 1 mL by the additionof DMSO. The resulting AAm-AI stock was mixed a40 mg portion of yeast RNA. For the acylation under the 25% or 50%v/v DMSO in water, an additional 1200 or 400 μL of nuclease-freewater was added, respectively. After the overnight incubation at roomtemperature under gentle shaking, the resulting RNA crosslinker wasprecipitated by the addition of 3 M sodium acetate (1/10 volume) andisopropanol (1.5 volume). The precipitated RNA crosslinker was isolatedby centrifugation  at 4 °C. The isolatedRNA crosslinker pellet was redissolved in water and further purifiedby additional precipitation and centrifugation. Finally, the purifiedRNA crosslinker pellet was dissolved in water, and the concentrationof RNA (mg/mL) was determined by measuring A260 [extinctioncoefficient = 40 (μg/mL)−1 cm–1]. To estimate of the degree of acylation, ∼3 mg of RNA crosslinkerwas dissolved in 600 μL of D2O followed by 1H NMR analysis.With400 μL of 1 M To homopolymerize the RNA crosslinker in a reactionvolume of 50 μL, acrylamido RNA crosslinker  was taken, and the volume was adjusted to44 μL by adding water. Subsequently, 1 μL of TEMED and5 μL of 10% APS in water were added, and the mixture was thoroughlymixed. The resulting mixture was incubated at room temperature for3 min to polymerize. For the copolymerization of the RNA crosslinker,3 M NIPAM stock (339.5 mg in 1 mL of DMSO) or 40% acrylamide mix [acrylamide/bis(acrylamide)= 29:1] was prepared. Next, acrylamido RNA crosslinker  was mixed with NIPAM stock  or acrylamide mix ,and the volume was brought to 44 μL by adding water. Subsequently,1 μL of TEMED and 5 μL of 10% APS in water were addedand thoroughly mixed. The resulting mixture was incubated at roomtemperature for 3 min to polymerize.