Polypyrimidine tract binding protein 1 (PTBP1) is one of the most well-described RNA binding proteins, known initially for its role as a splicing repressor before later studies revealed its numerous roles in RNA maturation, stability, and translation. While PTBP1’s various biological roles have been well-described, it remains unclear how its four RNA recognition motif (RRM) domains coordinate these functions. The early PTBP1 literature saw extensive effort placed in detailing structures of each of PTBP1’s RRMs, as well as their individual RNA sequence and structure preferences. However, limitations in high-throughput and high-resolution genomic approaches  precluded the functional translation of these findings into a mechanistic understanding of each RRM’s contribution to overall PTBP1 function. With the emergence of new technologies, it is now feasible to begin elucidating the individual contributions of each RRM to PTBP1 biological functions. Here, we review all the known literature describing the apo and RNA bound structures of each of PTBP1’s RRMs, as well as the emerging literature describing the dependence of specific RNA processing events on individual RRM domains. Our goal is to provide a framework of the structure–function context upon which to facilitate the interpretation of future studies interrogating the dynamics of PTBP1 function. While it is now known as a ubiquitous regulator of RNA biogenesis, polypyrimidine tract binding protein 1 (PTBP1)—also known as hnRNP I (heterogeneous nuclear ribonucleoprotein 1) ,3,4,5. EStructurally, PTBP1 is comprised of an N-terminal bipartite nuclear localization and export signal (NLS/NES) ,26,27,28As mentioned above, PTBP1 contains four RNA recognition motifs. Canonically, an RRM contains approximately 90 amino acids arranged in an αβ-sandwich with β1α1β2β3α2β4 topology ,35, in wPTBP1’s four RRMs are arranged in a “beads on a string” orientation, where each RRM is joined to its neighbor by linker regions of variable length. The linker regions following RRM1 (42 residues) and RRM2 (58–84 residues) are long, and these domains have been shown to move independently of their neighbor in solution ,33,52,53PTBP1 was initially described to bind short, single-stranded pyrimidine sequences , including those found at splice junctions ,11,63,64RRM1 of PTBP1 demonstrates the canonical RRM topology of β1α1β2β3α2β4 ,65; howeBased on these data, the preferred binding register for RRM1 is YCU with only the C nucleotide engaging in a canonically described interaction with conserved features of the RNA binding interface. The first position of this binding register can be occupied by either pyrimidine (C or U), because Q129 on β4 can act as either a hydrogen bond donor or acceptor, and can thus accommodate either pyrimidine . Of noteIntriguingly, several recent studies have demonstrated that the C-terminal region of the RRM1 domain is a critical allosteric regulator of RNA binding ,72. ThesUnlike RRM1, RRM2 demonstrates an extension of the canonical RRM topology with an additional fifth β-strand that sits adjacent and anti-parallel to β2, stabilized by a stacking interaction between Y275 (β5) and H201 (α1 helix) ,73. β4 aStudies characterizing RRM2′s interactions with various short pyrimidine sequences  identified that RRM2 binds the C3U4 doublet, as well as U6 ,53. As wBased on these data, the minimal binding register of RRM2 is CU(N)N, which is extended by an additional nucleotide as compared to the minimal binding register for RRM1 due to the U6 interaction with the C-terminal extension of this domain (the β5 and pseudo-sixth strand) not seen in RRM1. An additional study using longer sequences than the minimal binding register has also confirmed that there are significant chemical shift perturbations upon RNA binding in the β4-β5 loop, the β5 strand, and the pseudo-sixth strand, as reported above . AlthougThe linker region C-terminal to RRM2 (between RRM2 and RRM3) has important structural considerations. Notably, this linker region appears to be arranged in a relatively compact globular conformation, as compared to the RRM1-RRM2 linker, such that RRM2 and RRM3 are in closer proximity than RRM2 and its N-terminally adjacent domain RRM1 . AlternaNote that all residue numbers correspond to the longest isoform (PTB-4) that contains the entirety of exon 9 . RRM3 and RRM4 of PTBP1 (RRM3-4), like their N-terminal neighbors, demonstrate several non-canonical modifications. RRM3, like RRM2, contains a β5 strand that sits antiparallel to β2, and is connected to β4 by a long loop that is positioned above the RNA binding interface. RRM4, like RRM1, contains the canonical RRM topology, with four anti-parallel β-strands comprising the β-sheet ,32,33. HDue to the heterodimeric form adopted by RRM3-4, RRM: RNA interactions for the individual RRM3 and RRM4 domains were determined using the RRM3-4 didomain, in order to provide a more accurate structural interrogation of RNA interactions. As with RRM1 and RRM2, short pyrimidine sequences (C1U2C3U4C5U6) were utilized to identify the minimal binding register of each tandem domain and map specific interactions ,53. RRM3RRM4, like RRM1, binds the U2C3U4 triplet in a similar fashion but with several key differences . Like RRM1, C3 stacks on H483  RNP2 po. A laterThe organization of these RRMs (juxtaposed so that their β-sheets face in opposing directions) is unique among RNA binding proteins with tandem RRMs, and has important functional implications. All other RNA binding proteins with tandem domains organize these domains such that they bind immediately adjacent stretches of nucleotides on target RNA ,40,41. TIn all RRMs of PTBP1, although there are substitutions of aromatic amino acids at key RNA interacting positions within RNP1 and RNP2, the residues at these conserved positions still contact target RNA as reflected in seminal studies mapping RNA: RRM interactions ,53. RNP2The idea of separation of function among the RRM domains of PTBP1 was proposed in the early PTBP1 literature, and has been refined by numerous subsequent studies over the roughly four decades since PTBP1’s initial discovery. These early PTBP1 studies had suggested that RRM3 and RRM4 (RRM3-4) were the major drivers of RNA binding , whiSubsequent biochemical studies revealed that all four RRMs of PTBP1 bind RNA, demonstrating that the separation of functions within this protein is not as clearly delineated as “protein interacting” domains and “RNA binding” domains. Instead, these studies suggest that separation of functions within the PTBP1 protein is a result of unique sequence and structure preferences of each RRM for target RNA. While all four RRMs of PTBP1 utilize the residues at RNP2 position 2  and RNP1 position 5 to coordinate RNA, the network of hydrogen bonds and hydrophobic interactions that drive the specificity of RRM: RNA interactions are unique between the RRMs. Therefore, as described in the previous sections of this review, each RRM has a unique binding register, secondary structure preference and, especially in the case of RRM3-4, the ability to manipulate the structure of target RNA. This, combined with the solution behavior of the full PTBP1 protein , supports the idea that each RRM may provide unique contributions to independently modulate RNA specificity and, ultimately, PTBP1 function. In this section, we summarize the current evidence for domain-specific functions of each of PTBP1’s RRM domains (or didomain in the case of RRM3-4).Of the four RRMs of PTBP1, RRM1 has the most numerous examples in the literature of discrete domain-specific functions. As described above, a C-terminal α3 helix folds upon binding to an RNA hairpin in the ECMV IRES, and this helix was critical for PTBP1-mediated enhancement of IRES activity in vitro . While tIt is known that PTBP1 regulates multiple aspects of the biogenesis of several apoptotic genes ,88,89,90Finally, RRM1 has also been shown to have a critical function in regulating genes that control invasive potential in cancer. A study by Wang et al. demonstrated that PTBP1 binds the 5′ UTR (untranslated region) of the hypoxia inducible factor 1α (HIF-1α) transcript via its RRM1 and RRM3 domains, destabilizing HIF-1α and promoting an invasive phenotype in a non-small cell lung cancer (NSCLC) model . More reIt has been well described that RRMs can serve as protein–protein interaction interfaces in addition to their canonical RNA binding function ,62,96. IWhile the three isoforms of PTBP1 formed by alternative splicing at exon 9 are well described, there is also a fourth PTBP1 isoform—formed by the removal of exons 2-10—that contains only RRM3 and RRM4 ,75,102. As described above, the geometric orientation of these domains (in which their RNA binding interfaces face in opposite directions) is well poised to induce RNA looping and, indeed, prior studies have confirmed this ability of the RRM3-4 didomain ,83. It hSince its initial identification in 1988, PTBP1 has taken a center stage as a model protein for studying RBP biology and function. Its roles in virtually all stages of RNA maturation and utilization  have provided a rich substrate for study, as by these ubiquitous functions PTBP1 impacts a wider range of biological processes than any other RBP. It has been described as a critical regulator of gene expression homeostasis in various tissues, has been implicated in regulating development in multiple tissue types, and its dysregulation has been shown to drive phenotypes in multiple disease states including, but not limited to, various cancers.Despite PTBP1’s position as a main character in the field of RBP biology, there has been little description of the dynamic anatomy of this protein and how its various components  coordinate its diverse functions. In the early PTBP1 literature, the available genomic techniques did not allow for high-resolution and throughput analyses in cellular-based assays to elucidate PTBP1’s intra-protein functional dynamics in an endogenous environment. However, recent advances in genomic technology now provide a toolbox to answer these previously unanswered questions. The advent of CRISPR-based technology now allows the capability of deleting portions of or entire protein domains to interrogate their biological function endogenously. More efficient and cost-efficient immunoprecipitation and sequencing techniques allow higher resolution mapping of binding sites and even interrogation of RNA structure, when it is bound to proteins of interest. These approaches allow us the enhanced molecular resolution necessary to begin mapping modular roles of individual domains of RBPs on a target-specific basis.The current lack of knowledge of the dynamics of RBP—and specifically PTBP1—coordination of functions has rendered these proteins essentially “undruggable” and, therefore, most effort in therapeutic development has focused on interventions downstream of RBPs—e.g., targeting the dysregulated RNA or protein product. Deconstructing the modularity of RBPs will allow more targeted upstream interventions of dysregulated RBP function. For example, if a particular domain of an RBP drives a pathologic splicing event—e.g., RRM2 and α-tropomyosin—a small molecule inhibitor can be designed for the specific ligand (RNA): RRM that does not impact the function of the RBP at other targets. Our goal is that the information contained in this review will frame the structure–function context of each RRM of PTBP1 and facilitate the next generation of PTBP1 interrogation—moving from studying the macroscopic cellular function of the entire protein, to a higher-resolution view of how the protein coordinates these observed functions. Ultimately, we hope that this will provide a foundation for understanding how individual RBD domains (and their specific RNA interactions) can be targeted to treat developmental defects and disease.


Conditional control of RNA structure and function has emerged as an effective toolkit. Here, a strategy based on a one‐step introduction of diacylation linkers and azide groups on the 2′‐OH of RNA is advance. Selected from eight phosphine reagents, it is found that 2‐(diphenylphosphino)ethylamine has excellent performance in reducing azides via a Staudinger reduction to obtain the original RNA. It is demonstrated that the enzymatic activities of Cas13 and Cas9 can be regulated by chemically modified guide RNAs, and further achieved ligand‐induced gene editing in living cells by a controllable CRISPR/Cas9 system. In the current study, a diacylation crosslinking strategy is developed to regulate CRISPR systems and DPPEA is used to restore the RNA structure to conditionally control gene editing when needed. In these biological processes, interactions between RNA and gene expression play a crucial role at the post‐transcription level. In addition, RNA‐RNA interactions are the basis for the construction of many significant regulatory networks. Due to the great complex biological roles of RNA, there is no hesitation in developing chemical toolboxes to study the functions and interactions of RNA. Chemical strategies in controlling RNA functions are applied as post‐transcriptional modifications, which include dimethyl sulfate acylation and 2′‐OH acylation (SHAPE). The development of chemical probing in manipulating single guide RNA (sgRNA) has prevailed in recent years. The type II CRISPR system, CRISPR‐Cas9, requires a single nuclease and one guide RNA for site recognition of specific target sequences via Watson‐Crick base pairing. Cas13a, an RNA‐guided ribonuclease, exhibits a “collateral effect” upon target recognition. Moreover, guide RNA (gRNA) is an essential part of the CRISPR system, which has been used to establish versatile methods to regulate the activity of Cas proteins. To accurately manipulate the CRISPR/Cas9 system and achieve conditionally controlled gene expression as well, many efforts have been made to broaden CRISPR/Cas9 for regulation of a specific activity, such as study based on photo‐responsive strategies, or small molecules to control Cas9 protein activity and gRNA structure. Based on CRISPR technologies, a large number of research areas have been developed, which include severe acute respiratory syndrome coronavirus 2 (SARS‐CoV‐2) detection, genome editing, and gene therapy. Various chemical crosslinking approaches have been utilized to study nucleic acid interactions, such as formaldehyde, carbazole derivatives, 4‐thiouridin, 1,4‐phenyldiglyoxal, and psoralen crosslinking methods. It was recently reported that bis‐nicotinic azide (BIN) probes reacted with 2′‐OH groups of different RNA strands and, via a Staudinger reduction of the azide, triggered the masking group release. Given the outstanding performance of this crosslinking strategy, we anticipate that this crosslinking design will promote the application of small molecules to conditionally manipulate RNA structure and then conditionally control the CRISPR systems. Conditional control of gene editing could therefore be performed through gRNA modifications with a crosslinking strategy.RNAs play key roles in biological processes including gene expression, transcript splicing, messenger transcription, and translocation, as well as catalysis.1 ReceFigure3, here we showed that the BIN crosslinking probes had the following merits: 1) high modification efficiency, 2) low amounts of reduction agents, 3) highly feasible operations, and 4) excellent off‐to‐on switching. In addition, benefiting from the crosslinking approach of BIN2, we achieved the desired sgRNA modification and function of Cas9 in Hela cells for genome editing.In this project, we synthesized three types of BIN compounds with different spacer lengths between the linker and reaction sites. Compared to the crosslinking efficiency of BIN2 and BIN3 in sgRNAs of the CRISPR‐Cas system, we found that BIN2 exhibited excellent performance. This prevented sgRNA from pairing with the target DNA sequence until its protective groups were detached by phosphine reduction agents upon azide reduction Figure1. Unli22.1 we constructed a platform for chemically‐controlled CRISPR‐Cas genome editing via an RNA crosslinking strategy in which sgRNA was modified with azide‐substituted groups , BIN3 (12.5 mm), or BIN‐C (25 mm) in SHAPE buffer  for various durations at 37 °C. The results, shown in Figure m DPPEA was sufficient to drive the Cas9‐mediated cleavage of the target DNA to completion  and trans‐activating crRNA (tracrRNA) were assembled into sgRNA. To apply the crosslinking strategy in this combining system, with tracrRNA modified by BIN2, crosslinking bands were separated using denaturing PAGE Figure ,E. At th2.2 Subsequently, we applied the crosslinking method to 21 nt RNA strands to determine whether it had an impact on RNA binding between intermolecular RNA. Surprisingly, 21 nt RNA treated with BIN2 (crosslinked R‐21 nt) in the absence or presence of complementary R‐21nt‐c‐FAM showed that as the modification level increased, the RNA hybridization was gradually abolished . Interestingly, with an increased concentration of DPPEA, two obvious binding bands appeared on the native PAGE  is a rapid and sensitive RNA‐guided and RNA‐targeting system. To achieve artificial control of the CRISPR‐Cas13a system in vitro, we used BIN2‐modified gRNA to regulate the targeted RNA cleavage. It was anticipated that the gRNA treated with BIN2 would show a time‐dependent inhibition of RNA cleavage. The results showed that RNA cleavage was completely halted after 20 min, which implied that the crosslinking strategy markedly affected Cas13a activity  was incubated with DPPEA, target RNA, and Cas13a at 37 °C for 2 h, after which DPPEA dose‐dependent cleavage bands were exhibited  reacted with DPPEA at 37 °C for 1 h, and then reporter RNA and Cas13a were added into the system, inducing an increased fluorescence signal within 12 min. In addition, 128 µm DPPEA had almost no impact on Cas13a activity. Therefore, we confirmed that our crosslinking method effectively controlled Cas13a function.As a powerful RNA detection platform, CRISPR‐Cas13a ribonuclease can be applied to detect RNA viruses such as Zika, Dengue, and SARS‐CoV‐2.19 CRI Ultimately, we suggested that the crosslinking of gRNA, rather than the simple modification, affects the binding with Cas proteins.To clarify the mechanism by which the crosslinked gRNA regulated the CRISPR‐Cas13a system, we carried out a binding assay to investigate the interaction between the crosslinked gRNA and target RNA. In this experiment, gRNA containing a part of a sequence (23 nucleobases) completely paired with the targeted RNA with a 5′ FAM chromophore. We observed bands migrating more slowly than others on native PAGE, which illustrated that the unmodified gRNA bound well with the targeted RNA , Bst DNA pol, primer, dNTPs, and various concentrations of DPPEA were incubated at 37 °C for 3 h, and fully extended products were observed  almost completely recovered the original RNA  and SLX4 interacting protein (SLX4IP), were chosen as the targeted genes. According to the transfection protocol, the untreated and treated sgRNA were transfected into Hela‐OC cells with Lipofectamine 3000 for 4 h and then the medium was refreshed. After 24 h, the efficiency of the gene editing (insertion/deletion (indel) mutation) was evaluated by a T7EI nuclease assay. Before performing conditional control gene editing in live cells, we first assessed the cytotoxicity of DPPEA to Hela‐OC cells. We observed that DPPEA caused no toxicity in cells up to 512 µm  to explore the interaction between sgRNAs and protein. The HaloTag ligand was labeled on nuclease‐deficient Cas9 protein (dCas9) to facilitate the imaging analysis. This dCas9 protein retained gRNA binding activity. The EMSA results showed that the untreated sg‐GFP with dCas9 formed RNP complexes  were transfected into Hela‐OC cells for 4 h, and then different concentrations of DPPEA were added to the system and incubated for another 24 h. A reactivated indel efficiency was exhibited by DPPEA. Compared with untreated sg‐HBEGF, the crosslinked sg‐HBEGF, treated with 256 µm DPPEA, recovered up to 70% of the gene indel rates . This caging and decaging tendency was consistent with the editing efficiency of sg‐HBEGF. Moreover, these results showed that gene editing in cells, mediated by CRISPR‐Cas9, can be conditionally regulated by the chemical activation of sgRNA.To verify that the crosslinking approach regulated gene editing in a conditional manner, untreated sg‐HBEGF or crosslinked sg‐HBEGF , Bst DNA Polymerase (Bst DNA pol), M‐MuLV Reverse Transcriptase (M‐MuLV RT), and T7 Endonuclease I were commercially available from New England Biolabs (USA). Oligonucleotide sequences containing primers are provided in Table T4 DNA polymerase, Cas9 Nuclease,  And the synthetic routes are shown in supporting information.The target probes of BIN‐C, BIN2, and BIN3 are synthesized according to a procedure described in a previous study.16 Andm MgCl2, 30 mm DTT, 2 mm NTPs, 2 mm spermidine, 0.1 µg µL−1 T7 RNA polymerase and 100 mm HEPES‐KOH pH 7.9. Through phenol/chloroform/isopropanol extraction and ethanol precipitation to obtain sgRNAs. Notes, all sgRNAs were dispersed in ultra‐pure water and preserved at −20 °C for further use.The sgRNAs forward primers and reverse primers at the aid of T4 DNA polymerase to produce DNA template, which then were transcribed in vitro by T7‐RNA polymerase at 37 °C for 3 h in transcription aid buffer, containing 20 mm), BIN2 (12.5 mm), or BIN3 (12.5 mm); and 4‐dimethylaminopyridine  were reacted in SHAPE buffer at 37 °C for different periods of time, and then 90 µL ice ethanol was added to halt the reaction. Standard ethanol precipitation was used to obtain the crosslinked RNA at 100 ng µL−1, which was stored at −20 °C for the following experiments. Genomic DNA extracted from Hela cells was used to augment the target DNA using the following PCR primers: t‐SLX4IP‐F and t‐SLX4IP‐R for t‐SLX4IP; t‐HPRT1‐F and t‐HPRT1‐R for t‐HPRT1; and t‐HBEGF‐F and t‐HBEGF‐R for t‐HBEGF. In addition, GFP target DNA was extracted from the PEGFP‐C1 vector (Clontech) using t‐GFP‐1F and t‐GFP‐1R for t‐GFP1, and t‐GFP‐2F and t‐GFP‐2R for t‐GFP2). For the crosslinked sg‐SLX4IP‐inhibited DNA cleavage system, 40 ng of untreated sg‐SLX4IP and crosslinked sg‐SLX4IP were incubated with t‐SLX4IP (40 ng) and Cas9 (0.1 µm) in NEBuffer 3.1  at 37°C for 6 h, then heated to 75 °C for 5 min to quench the reaction. For the DPPEA activation of the DNA cleavage system, 40 ng of untreated sg‐SLX4IP or crosslinked sg‐SLX4IP together with t‐SLX4IP (40 ng) and Cas9 (0.1 µm) were treated with DPPEA at various concentrations for 6 h, and the other procedures were similar to the previous procedures. For the sg‐HBEGF, sg‐HPRT1, and sg‐GFP systems, 50 ng untreated sgRNAs or crosslinked sgRNAs were reacted with target DNA (50 ng) and Cas9 at 37 °C for 6 h, and the subsequent steps were the same as described previously in this section.sgRNAs were used as starting materials to prepare crosslinked sgRNAs. First, sgRNAs (3 µg); BIN‐C , and then injected on a 20% native PAGE . For the activation of RNA‐RNA combining experiment: Crosslinked R‐21nt‐c‐FAM incubated with various concentrations of DPPEA at 37 °C for 4 h, and following the R‐21 nt was added into. The following operation is likely to be above.21 nt RNA strand at the 5′ end with FAM labeling (R‐21nt‐c‐FAM) was used to test the modified RNA whether it could bind well with the complementary RNA (R‐21 nt). The crosslinked R‐21 nt was conducted similarly to the above general protocol. For the inhibition of RNA‐RNA hybridization assay: Untreated R‐21 nt (50 ng) and crosslinked R‐21nt‐c‐FAM with different modification levels were annealed in binding buffer  was utilized in the current study. First, the crosslinked gRNA was prepared according to the previously described protocol. For the inhibition system, crosslinked gRNA (20 ng) with different modification levels was reacted with target RNA (10 ng) and Cas 13a (45 nm) in Cas13a buffer  at 37 °C for 1 h. The reaction was quenched by adding formamide (4.0‐fold of the reaction volume). For off‐to‐on switching of CRISPR‐Cas13a, the crosslinked gRNAs , target RNA (10 ng), Cas13a (45 nm), and a series of concentrations of DPPEA were incubated at 37 °C for 1 h. Fluorescence dynamic assays were used to identify the high reactivity of Cas13a. In this assay, untreated gRNA or crosslinked gRNA (100 ng) and target RNA (50 ng) were rapidly mixed with reporter RNA (200 ng) and Cas13a in 1 × Cas13a buffer. The data were collected every second at room temperature for 10 min using a fluorescence spectrophotometer . For the DDPPEA conditional control reporter RNA cleavage assay, the crosslinked gRNA was incubated with various concentrations of DPPEA at 37 °C for 4 h, then the target RNA, reporter RNA, and Cas13a were added, and the data were collected every second.Cas13a obtained from m Tris‐HCl, 100 mm NaCl, 0.1 mm EDTA, and 0.01% Triton X‐100, pH 8.0 @ 25°C) were incubated with RNase I (0.01 U) or RNase T1 (0.005 U) for 10 min on ice atmosphere. The reaction was quenched by the addition of a stopping mixture (4.0‐fold reaction volume of formamide and containing 0.1% SDS). Notes, the RNase T1 buffer contained 100 mm Tris‐HCl and 50 mm EDTA, pH 7.4 @ 25 °C. These enzymatic reactions were characterized by denaturing PAGE analysis .In vitro crosslinked RNAs stability testing were carried out as described below. 50 ng crosslinked RNAs (R‐21nt‐c‐FAM) in the RNase I buffer . Then the samples were analyzed on JASCO‐810. The signals were recorded every centigrade degree from 4 °C to 95 °C at 260 nm. Untreated R‐21 nt binding with R‐21 nt was seen as a positive control.In the circular dichroism analysis assay, R‐21 nt and R‐21 nt were used, which can completely combine with each other through Watson‐Crick pairing. R‐21 nt and different levels of modification R‐21 nt were heated to 95 °C for 5 min and then cooled to room temperature naturally in CD buffer ‐2,5‐diphenyltetrazolium bromide (MTT). Cells were incubated with MTT (5 mg mL−1) for 4 h, and then the medium was discarded. Following 150 µL DMSO was added, and shaking at 70 rpm for 30 min at 37 °C. Finally, optical density was measured at 492 nm with a microplate reader .Hela‐OC cells (stable expression Cas9 protein) were cultured in a complete medium containing 89% (v/v) basal medium , 10% (v/v) fetal bovine serum  and 1% (v/v) pen strep (Genview) at 37 °C in a 5% CO5 per well) were seeded into 24 well plates and incubated for 24 h before transferring the sgRNAs into the cells. sgRNA  and DMEM (24 µL) were mixed with a solution containing Lipofectamine 3000  and 23.5 µL DMEM, which were incubated for 15 min at room temperature. The transfection was started when the 50 µL mixture was added to the cells. After 4 h, cells were washed with PBS buffer and fresh complete medium was used to culture cells for another 24 h. The genome was extracted using a FastPure Cell DNA Isolation Mini Kit (Vazyme) and mutations were detected using a T7EI nuclease assay. Prior to mutation detection, the target sites were amplified using PrimeSTAR HS DNA Polymerase with primers . The PCR product was purified by a PCR purification mini kit (TOROIVD), and 50 ng of the product was used as a substrate for T7 Endonuclease I (1.5 × 10−3 U for t‐SLX4IP or 3.5 × 10−3 U for t‐HBEGF) digestion. The reaction was performed at 37 °C for 1 h and quenched by adding loading dye containing 20 mm EDTA.Cells (1 × 10For blocking the gene editing assay, different crosslinking levels of sg‐HBEGF and sg‐SLX4IP were transfected into cells, and the rest of the procedure was similar to the previously described method.For DPPEA activation of the gene editing assay, sgRNAs were transfected into cells for 4 h, and then cells were treated with a reduction solution containing different concentrations of DPPEA. The cells were cultured for another 24 h, and the remaining procedure was similar to the previously described method.The authors declare no conflict of interest.H.L., T.Z., X.Y. contributed equally to this work. T.T. and X.Z. conceived the original idea, designed the studies and led the project. H.J.L., X.F.Y. and R.C.F. conducted all of biological studies. T.Y.Z. synthesized BIN‐C, BIN2, and BIN3, and analyzed the NMR data. W.X. synthesized HBC620 and HBC525. All the authors provided feedback on the study and on the manuscript. The authors declare no competing financial interests.Supporting informationClick here for additional data file.


Mutations in the RNA-binding protein FUS are linked to amyotrophic lateral sclerosis and frontotemporal dementia (ALS/FTD). FUS mutants mislocalize and aggregate in dying neurons. We previously established that FUS proteinopathy is linked to changes in the histone modification landscape in a yeast ALS/FTD model. Here, we examine whether FUS’ RNA binding is necessary for this connection. We find that overexpression of a FUS mutant unable to bind RNA is still associated with reduced levels of H3S10ph, H3K14ac and H3K56ac. Hence, FUS’ ability to bind RNA is not required in the mechanism connecting FUS proteinopathy to altered histone post-translational modifications. IntroductionSOD1, c9orf72, FUS, and TDP-43genes have all been implicated in ALS/FTD pathologyAmyotrophic lateral sclerosis (ALS) and frontotemporal dementia (FTD) form a ruinous neurodegenerative disease spectrumFUS(Fused in Sarcoma) encodes for the RNA-binding protein FUSEpigenetics refers to heritable changes to a phenotype occurring without directly affecting an organisms’ genetic background, typically through changes in chromatin structure and compositionSaccharomyces cerevisiaeFUS ALS/FTD modelPrevious work has revealed links between ALS/FTD and several histone modifying enzymes. For instance, the histone deacetylase (HDAC) HDAC1 mislocalizes to the cytoplasm in a FUS-ALS modelResults and DiscussionRNA binding by FUS is not necessary for histone PTM dysregulation)). We confirmed that FUS expression was comparable for both constructs via western blotting .We have previously shown that the levels of H3S10ph, H3K14ac and H3K56ac are significantly decreased in a FUS overexpression ALS/FTD yeast model). We did not observe a statistically significant difference in the levels of these two PTMs between FUS and RRM yeast. Similarly, we observed a reduction in H3S10ph levels in both FUS and RRM yeast compared to the control, with no significant difference between FUS and RRM yeast . Linear range assays for each of these antibodies is provided as Extended Data. These results suggest that dysregulation of histone PTMs occurs independently of FUS’ ability to bind RNA.We assessed the levels of H3S10ph, H3K14ac and H3K56ac in yeast overexpressing FUS, RRM and a control plasmid. Surprisingly, histone acetylation levels on H3K14 and H3K56 were decreased in both FUS and RRM yeast compared to control . Notably, we observed this effect despite protein overexpression. These results further support that FUS’ connection to PTM dysregulation and subsequent impact on total cellular RNA levels is independent of FUS-RNA binding.Collectively, the dysregulated PTMs are involved in active gene expressionAltogether, we find that histone PTM dysregulation is independent of RNA binding by FUS in a yeast overexpression model. It is important to note that some of the histone modifications we study are cell cycle dependent, and thus, without flow cytometry experiments we cannot completely discard the possibility that overexpression of FUS or RRM is altering cell cycle distribution. Moreover, our studies are limited to yeast models, and thus, mechanistic verification in other model systems is still necessary . Proteomic analysis of putative FUS binding partners in SH-SY5Y cells revealed a significant number of RNA processing, splicing and binding proteinsConclusionsSpecific histone PTM alterations are connected to FUS proteinopathy in yeast models, though the molecular mechanisms behind this association are unclear. RNA-binding through the RRM domain is not necessary for FUS aggregate formation in yeast, but it is necessary to elicit a growth suppression phenotype. Surprisingly, we have shown that FUS does not need to bind RNA to elicit changes in histone PTM levels or global RNA levels. Hence, it appears that histone PTM changes are linked to aggregation of FUS. Overall, our results suggest aggregation – rather than defects in than RNA binding – as an important feature connecting FUS to the epigenome in ALS/FTD.Materials and MethodsYeast Strains, Media and PlasmidsAll yeast were W303a .Protein Overexpression.Yeast strains were grown to saturation overnight in raffinose-supplemented dropout media at 30 °C and 200 rpm. Overnight cultures were then diluted to an OD600of 0.30 in galactose-supplemented synthetic dropout media and induced for 5 hours at 30°C. Yeast cultures were then standardized to the lowest OD600. Cells were then pelleted at 850 rcf at 4°C and washed three times with sterile distilled water and harvested. The supernatant was removed and the pellets were flash frozen in liquid nitrogen and stored at -80°C.Serial dilution growth assays.Yeast were grown to saturation overnight in raffinose-supplemented dropout media at 30°C. Overnight cultures were diluted 2-fold, then serially diluted 5-fold. Yeast were spotted onto solid synthetic dropout medium containing glucose or galactose with a pin-frogger. Yeast were grown at 30°C for 3 to 4 days before imaging.Western blotting.Western blottingwas performed as previously describedMicroscopy.Control, FUS, and RRM yeast pellets were thawed and resuspended in 70% (w/v) ethanol for 30 minutes at room temperature. Yeast were pelleted again and washed twice with PBS. Pellets were then resuspended in a small volume of PBS and imaged immediately. Slides were observed using a wide-field Axio Observer 7 Inverted Microscope (Zeiss) with a x63/1.4-numerical aperture (NA) Plan-Apochromat (oil immersion) objective. The resulting images were processed using both Zen Microscopy Software (Zen) and ImageJ (Fiji). Average percentages of budding vs. non-budding cells were calculated from five representative images for three biological replicates of each strain.RNA Purification.Frozen pellets were thawed and then treated with zymolyase 20-T . Total RNA was then purified from samples using a Qiagen RNeasy Miniprep Kit . RNA concentrations were measured using a NanoDrop Lite . RNA concentrations from FUS and RRM yeast were then compared to control yeast to obtain fold change values (sample/control), which were used for statistical analysis.Statistical Analysis.Statistical analysis of data was performed in GraphPad Prism 9 using the built-in stats package . Significant differences between nuclear intensity and histone modifying enzymes levels were determined using Welch’sttest withp = 0.05 as the cutoff. Significant differences between FUS, RRM, and control groups were determined using one-way ANOVA followed by Tukey’s test for pairwise comparison of the group means withp = 0.05 as the cutoff for significance. Error bars on the graphs represent standard deviation (SD) calculated from values obtained in the data analysis steps described above. All graphs were constructed with GraphPad Prism 9 10.22002/s6mgq-cqe22Description: Linear Range Assays for Selected Antibodies and Light Microscopy. Resource Type: Dataset. DOI:


Red clover necrotic mosaic virus (RCNMV) is a segmented positive-strand RNA virus consisting of RNA1 and RNA2. Previous studies demonstrated that efficient translation of RCNMV RNA2 requires de novo synthesis of RNA2 during infections, suggesting that RNA2 replication is required for its translation. We explored a potential mechanism underlying the regulation of replication-associated translation of RNA2 by examining RNA elements in its 5′ untranslated region (5′UTR). Structural analysis of the 5′UTR suggested that it can form two mutually exclusive configurations: a more thermodynamically stable conformation, termed the 5′-basal stem structure (5′BS), in which 5′-terminal sequences are base paired, and an alternative conformation, where the 5′-end segment is single stranded. Functional mutational analysis of the 5′UTR structure indicated that (i) 43S ribosomal subunits enter at the very 5′-end of RNA2; (ii) the alternative conformation, containing unpaired 5′-terminal nucleotides, mediates efficient translation; (iii) the 5′BS conformation, with a paired 5′-end segment, supresses translation; and (iv) the 5′BS conformation confers stability to RNA2 from 5′-to-3′ exoribonuclease Xrn1. Based on our results, we suggest that during infections, newly synthesized RNA2s transiently adopt the alternative conformation to allow for efficient translation, then refold into the 5′BS conformation, which supresses translation and promotes efficient RNA2 replication. The potential advantages of this proposed 5′UTR-based regulatory mechanism for coordinating RNA2 translation and replication are discussed. The genomes of positive-strand RNA viruses serve as direct templates for both ribosomes that translate viral proteins and viral RNA-dependent RNA polymerases (RdRps) that synthesize minus-strand RNA intermediates, the first step in genome replication. These two processes cannot occur simultaneously on the same RNA template due to their opposing directions, i.e., ribosomes travel in the 5′-to-3′ direction, while viral RdRps travel in the 3′-to-5′ direction. Consequently, regulation of these incompatible processes is required to allow for efficient viral proliferation during infections ,2,3,4.Dianthovirus, family Tombusviridae) possesses a segmented positive-strand RNA genome composed of RNA1 (3.9 kb) and RNA2 (1.5 kb)  A 5,6].  kb A . A. ADianth(1.5 kb) A 11,12],12DianthBoth RNA1 and RNA2 genomes lack a 5′-cap structure and a 3′-poly(A) tail and thus rely on alternative translation strategies for expression of their viral proteins . TranslaThe 81 nt long 5′UTR of RNA2 was previously investigated and shown to be involved in both RNA2 replication and translation of MP . In thatThe full-length cDNA clones of wild-type RCNMV RNA1 and RNA2 genome segments (kindly provided by Tim Sit and Steven Lommel) were used to generate all RCNMV constructs used in this study. R2HA is an RNA2 variant encoding a C-terminally HA-tagged MP. Mutants were derived by standard site-directed PCR-based mutagenesis, using the Q5 High-Fidelity DNA Polymerase kit (NEB). Sequencing was performed to ensure that only the intended modifications were present in mutants. Specific changes introduced into the RCNMV genomes are shown in the accompanying figures.4OAc on ice. RNA concentrations were measured by spectrophotometry, and transcript integrity was verified by agarose gel electrophoresis. 5′-monophosphorylated RNAs were generated as described previously [Uncapped viral RNAs were in-vitro-transcribed from SmaI-linearized RCNMV RNA1 and RNA2 constructs using the T7-FlashScribe transcription kit (Cellscript) as described previously . Brieflyeviously . RNA conSelective 2′-hydroxyl acylation analyzed by primer extension (SHAPE)  was perf5 protoplasts were transfected with infectious viral transcripts (3 μg of RNA1 and/or 1 μg of RNA2) using PEG-CaCl2 [32P-radiolabeled DNA oligonucleotide probes that are complementary to the 3′-end of RNA2 , as described previously [Cucumber protoplasts were isolated from 6-day-old cotyledons and isolated as described previously . ApproxiEG-CaCl2 . Transfeeviously . Viral RViral protein accumulation was detected via Western blotting. Total proteins were separated by 12% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and were transferred to PVDF membrane (Amersham Hybond P 0.45 PVDF). Ponceau S staining was performed to ensure equal loading of total proteins. C-terminally HA-tagged MP was detected with horseradish peroxidase (HRP)-conjugated anti-HA high affinity (3F10) rat monoclonal antibodies (Roche) at 1:4000 dilution. MP-HA bands were detected using ECL Select Western blotting detection reagent  and captured through MicroChemi imager (DNR Bio-Imaging Systems). MP levels were calculated relative to that of the reference genome (R1HA) and protein levels were normalized relative to corresponding RNA levels determined through Northern blotting. MP levels were quantified using Quantity One software. Experiments were repeated three times to generate averages and corresponding standard error of the means (SEMs).35S-methionine, as described previously [Uncapped RNA2 transcripts (0.5 pmol) were assessed for translation using a wheat germ extract in vitro translation system (Promega). MP accumulation was monitored by incorporation of eviously . TranslaIn vitro degradation assays were performed as described previously  using twThe 5′UTR was shown previously to be important for regulating translation of MP from RCNMV RNA2 . In thatPrevious analysis of the 5′UTR of RNA2 indicated that formation of 5′BS inhibited translation of MP, possibly by impeding ribosome entry at the 5′-end . To deteThe uAUGs and corresponding non-uAUG controls were introduced into the 5′UTR of R2HA  by single-nucleotide substitutions, so as to limit effects on higher order structure. Transfection of cucumber protoplasts with wt RNA1 (denoted as R1) and R2HA led to readily detectable accumulation of R2HA RNA in infections via Northern blotting B. ImportTo assess if 5′-end accessibility was altered in the uAUG mutants, the RNA2 transcripts were subjected to the yeast 5′-to-3′ exoribonuclease Xrn1, which requires a minimum of three single-stranded 5′-terminal nucleotides to engage its RNA substrate ,38. The Based on the finding that ribosomes enter and scan from the 5′-terminus of RNA2 , the posOur compensatory mutational analysis clearly established that formation of 5′BS in RNA2 has opposite effects on translation (inhibitory) and replication (stimulatory) . The 5′-To explore this possibility, we examined predicted suboptimal structures for the 5′UTR of RCNMV RNA2. A lower stability alternative conformation was identified in which sequences involved in forming the 3′-half of 5′BS were instead paired to nucleotides located 12 residues from the 5′-terminus, leaving a 5′-terminal segment unpaired B. In botTo evaluate the potential relevance of the alternative conformation A, RCNMV The primary sequences involved in the transition between the 5′BS conformation and the alternative conformation involve the first 20 nts of the 5′UTR and a segment just upstream of the AUG for MP A. The LSInterference with formation of both structures in min-7 yielded low levels of its RNA and high relative levels of MP B,C. FindThe low level of relative translation from min-R2HA versus R2HA suggested that deleting the LSL likely shifted conformational equilibrium towards the 5′BS C. ConverThis unexpected result indicated that further stabilization of the alternative structure in infections was doubly detrimental and suggested that a certain balance between the two conformations is required for optimal production of both RNA and protein. Interestingly, when the alternative-structure-stabilized min-R2HAs were tested for translation in vitro, the modifications led to increased MP production D, consisHere, we have examined further the roles of the 5′BS and 5′UTR in modulating RNA2 replication and translation in RCNMV. Three pieces of evidence support the formation and function of 5′BS: (i) solution structure probing results are consistent with the formation of 5′BS in RCNMV B, (ii) aBoth genome segments of RCNMV are uncapped, making them more vulnerable to cellular 5′-to-3′ exoribonucleases. In tombusviruses, both secondary and tertiary structural features in the 5′UTR of its uncapped genome form an RNA conformation that protects 5′-terminal nucleotides from Xrn attack by impeding its access . Our resThe balance between RNA2 replication and MP translation must be properly regulated during infections. Our results, along with those of Hyodo et al. , indicatIt is proposed that the 5′UTR of RNA2 can assume two mutually exclusive structural configurations: the 5′BS conformation, which prevents translation and permits replication, and the alternative conformation, which promotes translation and inhibits replication A. TransiA previous study showed that replication of RNA2 is required for translation of MP , and we Based on our results we propose a putative model to describe the involvement of the alternative conformation in regulating RNA2 replication and translation during infections. Upon RCNMV infection, RNA2 released from the virion would harbor the thermodynamically favorable 5′BS structure, which would both protect its 5′UTR from exoribonuclease attack and promote RNA2 replication. The presence of the 5′BS in invading RNA2 genomes would also prevent the ex-virion message from being immediately directed into translation. Unlike for RNA1, which encodes viral replication proteins p27/88) essential for genome amplification [8 essentiFollowing replication, to allow for translation of MP, newly synthesized RNA2s would form the alternative RNA structure transiently via co-transcriptional folding. This would both promote initiation of a first round of translation by increasing 43S subunit access and inhibit minus-strand synthesis (requiring 5′BS) that would interfere with translation. However, it is unlikely that a single round of translation would be able to provide the levels of MP needed for a successful infection. Thus, after the initial round of translation, additional rounds could follow if the 43S subunit traffic on the 5′UTR acted to delay the formation of 5′BS. Eventually, the nascent RNA2 messages would adopt the more stable 5′BS, and this would allow them to be replicated or packaged into virions. This model provides a possible mechanistic explanation for the observed requirement of RNA2 replication for MP translation during infections and offers new insights into the regulatory strategies utilized during RCNMV infections.


In plants, RNase III Dicer-like proteins (DCLs) act as sensors of dsRNAs and process them into short 21- to 24-nucleotide (nt) (s)RNAs. Plant DCL4 is involved in the biogenesis of either functional endogenous or exogenous  short interfering (si)RNAs, thus playing crucial antiviral roles.Saccharomyces cerevisiae, an RNAi-depleted organism, in which we could highlight the role of dicing as neither Argonautes nor RNA-dependent RNA polymerase is present. We have therefore tested the DCL4 functionality in processing exogenous dsRNA-like substrates, such as a replicase-assisted viral replicon defective-interfering RNA and RNA hairpin substrates, or endogenous antisense transcripts.In this study we expressed plant DCL4 in DCL4 was shown to be functional in processing dsRNA-like molecules in vitro and in vivo into 21- and 22-nt sRNAs. Conversely, DCL4 did not efficiently process a replicase-assisted viral replicon in vivo, providing evidence that viral RNAs are not accessible to DCL4 in membranes associated in active replication. Worthy of note, in yeast cells expressing DCL4, 21- and 22-nt sRNAs are associated with endogenous loci.S. cerevisiae. The results described herein show that S. cerevisiae can provide insights into the functionality of plant dicers and extend the S. cerevisiae tool to new biotechnological applications.We provide new keys to interpret what was studied so far on antiviral DCL4 in the host system. The results all together confirm the role of sense/antisense RNA-based regulation of gene expression, expanding the sense/antisense atlas of The online version contains supplementary material available at 10.1186/s11658-023-00469-2. Ribonucleic acid (RNA) has multiple roles in cellular functions, from coding genes to non-coding regulatory activities. The latter function is associated with the ability of RNA to form double-stranded (ds) or ds-like secondary structures. DsRNAs accumulate in the host cells in viral infections as a result of viral gene expression and/or viral replication. However, increasing evidence suggests that dsRNAs are not limited to viral origin, but can be produced from endogenous sources, such as retroelements, sense/antisense co-transcriptions –3 and, iPlant Dicer-like proteins (DCLs) are RNase III, double-stranded RNA (dsRNA)-specific endonucleases with specialized functions in producing short (s)RNAs of 21- to 24-nucleotides (nt), including micro (mi)RNAs and short interfering (si)RNAs of endogenous or viral origin. In turn, miRNAs and siRNAs guide the effector Argonautes (AGOs)-containing complexes to silence RNA target transcripts. RNA-silencing pathways contribute to viral defense, transposon silencing, heterochromatin formation, and post-transcriptional repression of cellular genes .Arabidopsis thaliana, DCL1, DCL2, DCL3 and DCL4 have specific roles in the biogenesis of distinct classes of endogenous short (s)RNAs: 21- and 22-nt miRNAs are predominantly generated by DCL1 , cDNAs were synthesized from 1 μg of total RNA extracted using the QuantiTect Reverse Transcription Kit (Qiagen) with the gene-specific primers: GFP, 5′-TGGAAGCGTTCAACTAGCAGA; ACT1, 5′-TCATGGTCGGTATGGGTCAA, following the manufacturer’s instructions. qRT-PCR reactions were carried out by means of a StepOnePlus real-time system (Applied Biosystems) and assembled in reaction with 1.5 μl cDNA, 10 μl SYBR® Select Master Mix (Applied Biosystem), 0.2 μl each of 100 mM of forward and reverse primer, and RNAse free water to 20 μl . Thermocycling was carried out with one cycle at 95°C for 10 min, followed by 40 cycles of 95°C for 45 s and 58°C for 1 min and 72°C for 45 s. The dissociation curve of the final products was checked to ascertain the presence of a single amplification product. qRT-PCR was performed on triplicate samples of each cDNA using the following primers: HpGFP, 5′-TGGAAGCGTTCAACTAGCAGA and 5′-CTAATTGGTAAAGATAGAGGAATTCGT; ACT1, 5′-TCATGGTCGGTATGGGTCAA and 5′-CCATATCGTCCCAGTTGGTGA, the S. cerevisiae Actin gene was selected as reference gene (GenBank: MK879550.1). Gene expression was calculated using the Delta–Delta cycle threshold method [For quantification of gene expression of yeast grown overnight at 25°C, with 0 μM and 250 μM [Cud method , 34.Saccharomyces cerevisiae strain YPH499 carrying pSAL-HADCL4 plasmid was sub-cultured in 10 mL of -LEU minimal media containing 2% dextrose at 30°C. The subculture was scaled up to two liters of the same medium containing 0.25 mM copper sulfate and grew O/N at 25°C until reaching the optical density at 600 nm of 0.8. Yeast cells were pelleted and frozen in dry ice upon storing at − 80°C. Frozen cells were resuspended in 5 mL TBS-Tween  of Complete EDTA-free protease inhibitors (Roche), with Tween®-20 (Merk) to a final concentration of 0.05%) in 15 mL Falcon tubes and then sonicated  in ice using output amplitude of 70%, 30 s per cycle for a total of 5 times per 2.5 min. The yeast lysate was splitted in four 2 mL eppendorf tubes and centrifuged at 16,000g for ten minutes at 4°C. The supernatant was transferred in a new 15 mL Falcon tube. The lysate was diluted 1:10 with TBS to a final concentration of Dithiothreitol (DTT) of 1 mM in a 50 mL Falcon tube and kept in ice. 0.4 mL of agarose-antiHA gel (Thermo Fisher), were washed twice with the same volume of TBS and added to 50 mL falcon tubes containing the yeast lysate. The yeast lysate was incubated with the agarose-antiHA gel O/N on the wheel at 4°C. The Falcon tubes have been centrifuged at 12,000 g for one minute and the pellet was washed three times with 1 mL ice-cold TBS-Tween (w/o DTT). The pellet was resuspended by gentle pipetting with wide-bore tips in 0.5 mL of TBS containing 1 mg/mL of HA peptide (Thermo Scientific) and incubated at 25°C with a slow shaking for 15 min. After centrifuging at RT for ten seconds at 12,000 g, the procedure was repeated for additional three times, thus obtaining three aliquots of supernatant  of 0.1 mL was transferred to a new eppendorf tube and kept on ice until used for the in vitro assay.Nicotiana tabacum BY2 cells were cultured at 23°C in Murashige–Skoog liquid medium. Evacuolated BY2 protoplasts were obtained by percoll gradient centrifugation and cytoplasmic extract (BYL) prepared as described earlier [32P-labeled in vitro transcribed RNA (1.5 pmol) [32P-labeled products were visualized by phosphor-imaging. earlier . Single-.5 pmol)  was incu.5 pmol)  or 5-µl S. cerevisiae S288C ; multireads were evaluated with RSEM [FASTQ quality was checked using FastQC . RNA-Seqith RSEM  and Multith RSEM  tools; dith RSEM , which uith RSEM . sRNA-Seith RSEM  on the Sith RSEM .A. thaliana col0 wild type, dcl2-4, dcl4-1 single mutants and dcl4dcl2 double mutants [https://www.ncbi.nlm.nih.gov/geo/ under accession number GSE136164. Row data underwent the elaboration used for SSC RNA-derived sRNAs and were aligned to TAS2 cDNA (locus AT2G39681).4 replicas of sRNA datasets from  mutants  are fromNicotiana benthamiana is widely considered a model plant in plant virology, the recognized host for study tombusviruses [ (CUP1) inducible promoter ; BYL was previously shown to contain endogenous DCL activities with a predominant action of DCL3 [To test whether the recovered HA-DCL4 was functional, we incubated DEYF at 25°C with typical substrates of plant DCLs, either a highly structured single-stranded (ss) DI-RNA  or a per of DCL3 . InteresS. castellii DCR1 into 23-nt sRNAs [Subsequently, we tested whether DCL4 was capable of processing RNA substrates in vivo under cell growth conditions. For this purpose, we co-expressed DCL4 with an RNA molecule as a substrate, which has been earlier determined as a ‘strong silencing construct’ (SSC) of the mRNA reporter of GFP as a means of inducing processing by nt sRNAs .RAD9 intron [The SSC consists of two complementary regions corresponding to a portion of the GFP mRNA separated by the 9 intron . Upon ga9 intron , after GS. cerevisiae RNaseIII-like protein) is not capable of producing 2-nt 3′ overhangs sRNA duplexes having the chemical features of Dicer products, i.e.5′- monophosphates and 3′-hydroxyls. To gain more detailed insight into the composition of the DCL4-mediated processing products of the SSC and DCL4 co-expressing yeast cells, we prepared sRNA libraries with standard protocols and sequenced them using next generation sequence (NGS) analysis by preparing sRNA-seq libraries . In turn, sequencing libraries were prepared representing the subset of sRNAs with 5′-monophosphates and 3′-hydroxyls, which are the chemical features of DCL4 products. As expected and according to previous studies [S. cerevisiae system, the (+) and (−) strands of the DI-RNA were stabilized cooperatively by the viral replication components [When components of the tombusvirus replicase complex are expressed, DI-RNA transcripts are amplified with the formation of (−) and ( +) strands and the synthesis of head-to-tail dimers. Considereported , 52 in t studies , the pre studies . This fimponents  and protmponents . All theAntisense transcription is recognized as an important inter-kingdom RNA-based regulatory mechanism of gene expression. The transcription of antisense RNAs is part of self-regulatory circuits that allow genes to regulate their own expression. The non-coding transcripts denoted as RNA of Unknown Functions 5 (RUF5-1 and 2) are well-known regulatory and antisense transcripts of genes encoding copper chelation proteins CUP1-1 and 2, respectively . Copper S. cerevisiae genome,  and sRNA production (sRNAseq) data to search for unknown or sense/antisense transcripts non yet annotated  and (−) strands were also detected in association with the IsoCitrate Lyase (ICL1) locus. ICL1 is involved in the glyoxylate cycle, and its expression enables the cell to grow on non-fermentable carbon sources (i.e. ethanol). Its expression is known to be de-repressed in the absence of glucose . ExpressThe above evidence prompted us to verify whether the presence of 21- and 22-nt sRNAs is associated with other yet unannotated antisense transcripts, as previously described by Yassour et al. . We haveS. cerevisiae is active in processing ss- and ds- defective-interfering RNAs of tomato bushy stunt virus in vitro , we could not find neither an analogue to plant DRB4 nor to animal dsRNA-binding proteins in S. cerevisiae. Accordingly, we observed the strong capacity of DCL4 to process the predicted dsRNA region of SSC RNA into 21-nt sRNAs followed by 22-nt sRNAs  we found that the size profile of sRNAs from the TAS2 transcript in Arabidopsis paralleled what we found in yeast  is a potent trigger of RNA silencing because the secondary structure of the molecule is similar to dsRNAs and, considering its ability to accumulate in infected tissues, is a preferred substrate of antiviral DCLs, including DCL4 [In mammals, studies have failed to detect significant levels of virus-derived siRNAs in several virus/host systems despite the presence of Dicers and Argonautes, which makes mammal cell RNAi compatible. Explanations come from recent studies showing that interferon response in mammalian somatic cells masks dsRNAi, which prevents Dicer sensing , 68. In  complex . In N. bing DCL4 .S. cerevisiae system the (+) and (−) strands of the DI-RNA are stabilized cooperatively by the viral replication components [Yeast systems have been exploited for studying the replication of tombusvirus DI-RNAs. Whether dsRNA replication intermediates are triggers of vsiRNA production is still an open question in both animal and plant virus systems. In the mponents  and the mponents  and protmponents . In the mponents , 74. IndS. cerevisiae, S. castellii, and presumably other budding yeasts. In the present study we extended previous applications of heterologous Dicers in S. cerevisiae and the potential to fully reproduce the ruler-cutting function of DCL4 with no lethal effects or abnormalities for the recipient. In accordance with the previous observation on the processivity of S. castellii DCR1, we observed a massive amount of sRNA by DCL4 from the yeast retrotransposon Ty-1 . For inAdditional file 1: Figure S1. RNAfold results in the case of aberrant SSC transcripts.Additional file 2: Table S1. Differentially expressed genes (DESeq2). DESeq2 results for differential expression analysis of RNA-Seq data. Legend: baseMean (mean read count) is highlighted in yellow if > 50; log2FC > 0.585 (corresponding to FC > 1.5) is colored in red, log2FC < -0.585 is blue; significant p-value < 0.05 is green.Additional file 3: Table S2. Sense-antisense annotated genes derived sRNAs. Results are shown for each replicate . Genes highlighted in yellow are represented in the paper: RUF5-1/CUP1-1 and RUF5-2/CUP1-2 . Genes highlighted in yellow are represented in the paper: CYC1, UTR1 and ICL1 . The schematic shows the proviral Ty1 reference (Acc. N. M18706.1) with long terminal repeats (LTRs) and gag/pol expression organization of mRNA.Additional file 6. Replicates Raw Western blots and nucleic acid gels. Replicates Raw Western blots and nucleic acid gels relative to the images at A) Fig. S. cerevisiae, B) Fig. , in vitro functionality of yeast crude extract or DEYF in processing Tombusviruses dsRNAs in vitro transcribed, D) Fig.


The testis is a key male reproductive organ that produces gametes through the process of spermatogenesis. Testis morphologies, sperm phenotypes, and the process of spermatogenesis evolve rapidly in mammals, presumably due to the evolutionary pressure on males to give rise to their own offspring. Here, we review studies illuminating the molecular evolution of the testis, in particular large-scale transcriptomic studies, which were based on bulk tissue samples and, more recently, individual cells. Together with various genomic and epigenomic data, these studies have unveiled the cellular source, molecular mechanisms, and evolutionary forces that underlie the rapid phenotypic evolution of the testis. They also revealed shared  and species-specific spermatogenic gene expression programs. The insights and available data that have accumulated also provide a valuable resource for the investigation and treatment of male fertility disorders – a dramatically increasing problem in modern industrial societies. •The testis is a rapidly evolving organ, on both the transcriptomic and translatomic layer.•The force of sperm competition has shaped the evolution of this organ.•Insights from new data provide a resource for future investigations of testicular molecular evolution and male fertility. The testis is the male reproductive gonad in all bilaterian animals, including humans and other mammals. The function of the testis is to produce sperm, through the process of spermatogenesis, but also androgens, primarily testosterone. The testis has evolved rapidly in mammals, which is likely mainly due to positive selection associated with the force of “sperm competition”, which reflects the evolutionary pressure on males to be reproductively successful; that is, to fertilize female eggs and thus give rise to their own offspring . This prStudies in the past two decades have revealed that the rapid evolution of the testis is reflected at the molecular level. In particular the advent of RNA sequencing (RNA-seq) protocols has facilitated global large-scale cross-species investigations. Here, we discuss the exciting fundamental insights that these novel approaches have provided into the molecular basis of testicular evolution and the peculiar features of spermatogenesis.We set the stage by introducing the spermatogenic/testicular cell types and highlighting the peculiar chromatin dynamics of the autosomes and sex chromosomes during spermatogenesis. We then summarize how bulk tissue RNA-seq and ribosome profiling (or Ribo-seq) studies have shed light on the consequences of these chromatin dynamics but also on the extensive translational regulation that is occurring during spermatogenesis. We proceed by discussing insights into the molecular evolution of the testis obtained from cross-species comparisons of these data types, to then move on to describe how recent evolutionary single-cell RNA-seq studies have led to groundbreaking findings regarding the cellular source and evolutionary forces underlying the rapid evolution of the testis. Next, we delve into the intricate and intriguing patterns and mechanisms of sex chromosome evolution pertaining to spermatogenesis. We conclude our account by discussing overall implications of the recent large-scale genomics investigations of the testis and by identifying promising future research avenues.Spermatogenesis takes place in seminiferous tubules, which crowd the testis . SpecifiDuring spermatogenesis spermatogonia differentiate into spermatocytes, where meiosis takes place . The meiSpermatogenesis displays several unique and peculiar molecular chromatin features. Chromatin in spermatogenic cells is indeed massively remodeled during spermatogenesis. This process involves repackaging of the DNA initially around testis-specific histones and histone variants, then around transition proteins, and in the end around small protamines, which allow dense packaging of the DNA into the compact sperm head  Fig. 2)Fig. 2). Molecular biologists have been aware for decades that the testis expresses many or even most genes – more genes than other organs – and have therefore, for example, often resorted to the testis for cloning experiments. Initial analyses of RNA-seq data derived from bulk tissue samples confirmed this notion at a genome-wide scale , indicatRecent genome-wide investigations of mRNA translation based on Ribo-seq  – a highGlobal comparisons of gene expression patterns across species based on bulk tissue samples provided initial insights into the rates of expression evolution in different mammalian organs. Early work, which was using microarrays to compare gene expression levels of corresponding one-to-one (1:1) orthologous genes between humans and chimpanzees for several organs, already showed that the testis had the greatest divergence in gene expression levels among the investigated organs . Later sNotably, our recent work has brought the investigation of gene expression evolution from the transcriptome to the next expression layer – the translatome – based on Ribo-seq data for several organs (including the testis) from representative mammals . This stRecent high-throughput single-cell (sc) or single-nucleus (sn) RNA-seq technologies have enabled detailed investigations of the cellular and molecular evolution of organs. We thus recently performed an extensive evolutionary single-cell transcriptome study for testes from eleven species that cover the three major mammalian lineages  and birds (the evolutionary outgroup), and include seven primates  Fig. 4)Fig. 4). To trace the cellular source of the rapid evolution of the testis, we built expression trees for the different cell types based on these data and found that the total branch lengths of the trees, which reflect the amount of evolutionary expression change (akin to the aforementioned bulk RNA-seq work), vary substantially between cell types. Interestingly, while the rate of expression evolution is similar in Sertoli cells and diploid spermatogenic cells (and lower than that in other somatic cell types), it was found to be substantially higher in the post-meiotic haploid cell types , consistent with previous work and a recent inference based on data for three placental mammals . PairwisWe sought to trace the evolutionary forces underlying the rapid evolution of late spermatogenesis and reasoned that two non-mutually exclusive patterns of natural selection may account for this finding . First, Specifically, signatures of functional constraints along spermatogenesis were assessed based on patterns of the tolerance to functional mutations, mouse knockouts, and rates of amino acid altering substitutions in coding sequences of genes expressed in different spermatogenic stages. All of these analyses showed a progressive reduction of constraint towards late stages of spermatogenesis across species, typically with the strongest reduction in early spermiogenesis; that is, in round spermatids .On the other hand, an examination of the temporal expression pattern of genes whose encoded protein sequences have been shaped by positive selection revealed a significant increase in percentages of positively selected genes used during spermatogenesis, with a peak in round spermatids . MoreoveBut what are the fundaments underlying the dynamic changes of selective forces and patterns of innovation during spermatogenesis? Given that the breadth of expression across tissues and developmental processes (termed “expression pleiotropy”) was proposed to represent a main determinant of the types of mutations that are permissible under selection , we asseIn our study , we alsoNotably, integrated analyses of our data and known fertility phenotypes  unveiledFinally, we used our single-nucleus spermatogenic data to scrutinize expression patterns of sex chromosomal genes and their evolution across mammals . Two setOur analyses provided several intriguing insights into the expression of sex chromosome-linked genes and their evolution . First, Here we have reviewed cross-mammalian transcriptome studies that – initially at the bulk-tissue level and more recently at the level of individual cells – have unveiled details of the unique and complex biology and molecular evolution of the testis and the process of spermatogenesis that takes place in this organ. The available data also provide a rich resource not only for future investigations of the molecular evolution across mammals but also for studies of the biology of the testis and associated fertility disorders – a dramatically increasing problem especially in industrialized regions of the world . In the Nils Trost designed and contributed figures and co-wrote the manuscript.Noe Mbengue designed and contributed figures and co-wrote the manuscript.Henrik Kaessmann wrote the manuscript.German Research Council  and the 10.13039/501100000781European Research Council  to H.K.This review and the new research in our group covered at the end of this review by Murat et al. were supported by grants from the