RNA viruses are a major threat to animals and plants. RNA interference (RNAi) and the interferon response provide innate antiviral defense against RNA viruses. Here, we performed a large-scale screen using Caenorhabditis elegans and its natural pathogen the Orsay virus (OrV), and we identified cde-1 as important for antiviral defense. CDE-1 is a homolog of the mammalian TUT4 and TUT7 terminal uridylyltransferases (collectively called TUT4(7)); its catalytic activity is required for its antiviral function. CDE-1 uridylates the 3ʹ end of the OrV RNA genome and promotes its degradation in a manner independent of the RNAi pathway. Likewise, TUT4(7) enzymes uridylate influenza A virus (IAV) mRNAs in mammalian cells. Deletion of TUT4(7) leads to increased IAV mRNA and protein levels. Collectively, these data implicate 3ʹ-terminal uridylation of viral RNAs as a conserved antiviral defense mechanism.
Subscribe to Journal
Get full journal access for 1 year
only $4.92 per issue
All prices are NET prices.
VAT will be added later in the checkout.
Tax calculation will be finalised during checkout.
Rent or Buy article
Get time limited or full article access on ReadCube.
All prices are NET prices.
Ding, S.-W. & Voinnet, O. Antiviral immunity directed by small RNAs. Cell 130, 413–426 (2007).
Goubau, D., Deddouche, S. & Reis e Sousa, C. Cytosolic sensing of viruses. Immunity 38, 855–869 (2013).
Schoggins, J. W. et al. A diverse range of gene products are effectors of the type I interferon antiviral response. Nature 472, 481–485 (2011).
Félix, M.-A. et al. Natural and experimental infection of Caenorhabditis nematodes by novel viruses related to nodaviruses. PLoS Biol. 9, e1000586 (2011).
Ashe, A. et al. A deletion polymorphism in the Caenorhabditis elegans RIG-I homolog disables viral RNA dicing and antiviral immunity. eLife 2, e00994 (2013).
Ashe, A., Sarkies, P., Le Pen, J., Tanguy, M. & Miska, E. A. Antiviral RNAi against Orsay virus is neither systemic nor transgenerational in Caenorhabditis elegans. J. Virol. 89, 12035–12046 (2015).
Guo, Y. R. et al. Crystal structure of a nematode-infecting virus. Proc. Natl. Acad. Sci. USA 111, 12781–12786 (2014).
Jiang, H. et al. Orsay virus utilizes ribosomal frameshifting to express a novel protein that is incorporated into virions. Virology 450-451, 213–221 (2014).
Franz, C. J. et al. Orsay, Santeuil and Le Blanc viruses primarily infect intestinal cells in Caenorhabditis nematodes. Virology 448, 255–264 (2014).
Sarkies, P., Ashe, A., Le Pen, J., McKie, M. A. & Miska, E. A. Competition between virus-derived and endogenous small RNAs regulates gene expression in Caenorhabditis elegans. Genome Res. 23, 1258–1270 (2013).
Guo, X., Zhang, R., Wang, J., Ding, S.-W. & Lu, R. Homologous RIG-I-like helicase proteins direct RNAi-mediated antiviral immunity in C. elegans by distinct mechanisms. Proc. Natl. Acad. Sci. USA 110, 16085–16090 (2013).
Fan, Y. et al. Structure of a pentameric virion-associated fiber with a potential role in Orsay virus entry to host cells. PLoS Pathog. 13, e1006231 (2017).
Duchaine, T. F. et al. Functional proteomics reveals the biochemical niche of C. elegans DCR-1 in multiple small-RNA-mediated pathways. Cell 124, 343–354 (2006).
Tabara, H., Yigit, E., Siomi, H. & Mello, C. C. The dsRNA binding protein RDE-4 interacts with RDE-1, DCR-1, and a DExH-box helicase to direct RNAi in C. elegans. Cell 109, 861–871 (2002).
Jiang, H., Chen, K., Sandoval, L. E., Leung, C. & Wang, D. An evolutionarily conserved pathway essential for Orsay virus infection of Caenorhabditis elegans. MBio 8, e00940–17 (2017).
Tanguy, M. et al. An alternative STAT signaling pathway acts in viral immunity in Caenorhabditis elegans. MBio 8, e00924–17 (2017).
van Wolfswinkel, J. C. et al. CDE-1 affects chromosome segregation through uridylation of CSR-1-bound siRNAs. Cell 139, 135–148 (2009).
Olsen, A., Vantipalli, M. C. & Lithgow, G. J. Checkpoint proteins control survival of the postmitotic cells in Caenorhabditis elegans. Science 312, 1381–1385 (2006).
Kwak, J. E. & Wickens, M. A family of poly(U) polymerases. RNA 13, 860–867 (2007).
Heo, I. et al. TUT4 in concert with Lin28 suppresses microRNA biogenesis through pre-microRNA uridylation. Cell 138, 696–708 (2009).
Norbury, C. J. Cytoplasmic RNA: a case of the tail wagging the dog. Nat. Rev. Mol. Cell Biol. 14, 643–653 (2013).
Wickens, M. & Kwak, J. E. Molecular biology. A tail tale for U. Science 319, 1344–1345 (2008).
Lee, M., Kim, B. & Kim, V. N. Emerging roles of RNA modification: m6A and U-tail. Cell 158, 980–987 (2014).
Rissland, O. S. & Norbury, C. J. Decapping is preceded by 3′ uridylation in a novel pathway of bulk mRNA turnover. Nat. Struct. Mol. Biol. 16, 616–623 (2009).
Morgan, M. et al. mRNA 3′ uridylation and poly(A) tail length sculpt the mammalian maternal transcriptome. Nature 548, 347–351 (2017).
Lim, J. et al. Uridylation by TUT4 and TUT7 marks mRNA for degradation. Cell 159, 1365–1376 (2014).
Chang, H., Lim, J., Ha, M. & Kim, V. N. TAIL-seq: genome-wide determination of poly(A) tail length and 3′ end modifications. Mol. Cell 53, 1044–1052 (2014).
Miki, T. S., Rüegger, S., Gaidatzis, D., Stadler, M. B. & Großhans, H. Engineering of a conditional allele reveals multiple roles of XRN2 in Caenorhabditis elegans development and substrate specificity in microRNA turnover. Nucleic Acids Res. 42, 4056–4067 (2014).
Kamath, R. S. et al. Systematic functional analysis of the Caenorhabditis elegans genome using RNAi. Nature 421, 231–237 (2003).
Molleston, J. M. et al. A conserved virus-induced cytoplasmic TRAMP-like complex recruits the exosome to target viral RNA for degradation. Genes Dev. 30, 1658–1670 (2016).
Huo, Y. et al. Widespread 3′-end uridylation in eukaryotic RNA viruses. Sci. Rep. 6, 25454 (2016).
Samji, T. Influenza A: understanding the viral life cycle. Yale J. Biol. Med. 82, 153–159 (2009).
Rehwinkel, J. Is anti-viral defence the evolutionary origin of mRNA turnover? BioEssays 38, 817 (2016).
Hamid, F. M. & Makeyev, E. V. Exaptive origins of regulated mRNA decay in eukaryotes. BioEssays 38, 830–838 (2016).
Manokaran, G. et al. Dengue subgenomic RNA binds TRIM25 to inhibit interferon expression for epidemiological fitness. Science 350, 217–221 (2015).
Brenner, S. The genetics of Caenorhabditis elegans. Genetics 77, 71–94 (1974).
Fire, A. Integrative transformation of Caenorhabditis elegans. EMBO J. 5, 2673–2680 (1986).
Jorgensen, E. M. & Mango, S. E. The art and design of genetic screens: Caenorhabditis elegans. Nat. Rev. Genet. 3, 356–369 (2002).
Sarov, M. et al. A genome-scale resource for in vivo tag-based protein function exploration in C. elegans. Cell 150, 855–866 (2012).
Dobin, A. et al. STAR: ultrafast universal RNA-seq aligner. Bioinformatics 29, 15–21 (2013).
Liao, Y., Smyth, G. K. & Shi, W. featureCounts: an efficient general purpose program for assigning sequence reads to genomic features. Bioinformatics 30, 923–930 (2014).
Love, M. I., Huber, W. & Anders, S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 15, 550 (2014).
Paix, A., Folkmann, A., Rasoloson, D. & Seydoux, G. High efficiency, homology-directed genome editing in Caenorhabditis elegans using CRISPR-Cas9 ribonucleoprotein complexes. Genetics 201, 47–54 (2015).
Martin, M. Cutadapt removes adapter sequences from high-throughput sequencing reads. EMBnet J 17, 10 (2011).
Harris, T. W. et al. WormBase 2014: new views of curated biology. Nucleic Acids Res. 42, D789–D793 (2014).
Quinlan, A. R. & Hall, I. M. BEDTools: a flexible suite of utilities for comparing genomic features. Bioinformatics 26, 841–842 (2010).
Robinson, M. D., McCarthy, D. J. & Smyth, G. K. edgeR: a Bioconductor package for differential expression analysis of digital gene expression data. Bioinformatics 26, 139–140 (2010).
Zhang, J., Kobert, K., Flouri, T. & Stamatakis, A. PEAR: a fast and accurate Illumina Paired-End reAd mergeR. Bioinformatics 30, 614–620 (2014).
Kawakami, E. et al. Strand-specific real-time RT-PCR for distinguishing influenza vRNA, cRNA, and mRNA. J. Virol. Methods 173, 1–6 (2011).
We thank M. Tanguy (Gurdon Institute, University of Cambridge) for OrV viral filtrates, L. Frézal for help with OrV RNA fluorescence in situ hybridization, I. Wilkinson for support with the genetic screen, N. J. Lehrbach for help with microinjections and M. Ridyard for lab management. We thank K. Harnish, F. Braukmann and S. Moss for high-throughput-sequencing support. We are grateful to V. N. Kim and H. Chang for sharing information on TAIL-seq and A. C. M. Boon (Washington University School of Medicine) for providing IAV. We thank A. Ashe and P. Sarkies for theoretical input on the screen design. We thank the International C. elegans Gene Knockout Consortium, the TransgeneOme project, J. Ahringer and M.-A. Félix for providing reagents. We thank V. Benes and the EMBL genome core for sequencing support. We thank G. Allen and C. Bradshaw for core bioinformatics support. We thank R. Medhi and D. Zijlmans for help with TUT western blots. This work was supported by the following grants to E.A.M.: Cancer Research UK (C13474/A18583 and C6946/A14492), the Wellcome Trust (104640/Z/14/Z and 092096/Z/10/Z) and The European Research Council (ERC grant 260688). J.L.P. was supported by the Wellcome Trust (093970/Z/10/Z). A.K. is supported by the Wellcome trust (102452/Z/13/Z). D.W. is supported as an Investigator through the Pathogenesis of Infectious Disease Award from the Burroughs Wellcome Fund.
The authors declare no competing interests.
Publisher’s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Integrated supplementary information
Supplementary Figure 1 The viral stress sensor (lys-3p::GFP) is constitutively active in some tissues but is induced in the intestine after severe viral infection.
a, Comparison of viral load and the lys-3 and sdz-6 mRNA expression after two days of infection by qRT-PCR, strains as indicated. Dots: independent infection. Samples as in Fig. 3d,e. b, Representative confocal sections (10 × or 20 × magnification, as specified) of the viral stress sensor in wild type and drh-1 mutants without infection. The viral stress sensor exhibited constitutive activity in uninfected individuals, which was restricted to specific tissues. GFP was observed at all developmental stages in the pharynx and the rectum of hermaphrodites. Additionally, hermaphrodites at the L4 larval stage would show a strong GFP signal around the vulva and gravid adults exhibited the GFP in the uterine lumen. In males, GFP was observed in the pharynx and the tail. GFP expression was comparable in wild type and drh-1 mutants and independent of viral infection. Thus, the gene lys-3 is constitutively active in tissues neighboring openings exposed to the environment, the most likely entry points of potential bacterial pathogens. c, Representative confocal sections (20 × magnification) of young adults (strains as indicated) carrying the viral stress sensor. Animals were uninfected (mock) or infected with OrV for four days. The viral stress sensor was strongly induced in the intestine after infection of drh-1 mutants, which is in agreement with the tropism of OrV. Intestinal GFP was most often visible around the collar of the nematodes, in the anterior region of the intestine in young adults. Some infected individuals exhibited a strong GFP signal throughout their entire body (data not shown), suggesting that the induction of the viral stress sensor can spread from cell to cell, like an inflammation process. The viral stress sensor offers an opportunity to easily monitor viral infections in living animals. Source data
a, Workflow of cde-1/ovid-9 (mj414) × cde-1 (tm1021) F8 recombinant family generation. A similar strategy was used to construct the cde-1 (mj414) × drh-1 (ok3495) F8 recombinant families. All animals were homozygous for the viral stress sensor (mjIs228). b-c, Number of families that activated the viral stress sensor in more than 20% of individuals after four days of infection with OrV. Approximately 50 individuals scored per family. Bars: number of families meeting criteria as indicated. Source data
Workflow and data monitoring the inter-individual transmission of OrV infection (in strains as indicated) using the viral stress sensor. Source data
a, Representative confocal sections (20 × magnification) of OrV in vivo RNA FISH. b, Representative confocal section (10 × magnification) of a C. elegans L4 larva expressing cde-1::GFP. As two previous reports disagreed about the expression pattern of CDE-1 (Olsen, A. et al. Science 312, 1381–1385, 2006; van Wolfswinkel, J. C. et al. Cell 139, 135–148, 2009), we used fosmid-recombineering to generate transgenic animals driving GFP expression from an endogenous genomic context. c, Diagram of the cde-1 rescue transgene, using the intestine-specific promoter of the gene vha-6. This transgene was injected in cde-1 null mutants. d, Viral load as measured by qRT-PCR of OrV RNA1 genome in adults two days after infection. Bars: average value; error: SEM; five independent infections. One-tailed student’s t-test: *** p < 0.001, **p < 0.01. e, Incidence of male in the progeny of hermaphrodites left to self-fertilize at 25 °C, in strains as indicated. Source data
a, Non-templated nucleotides at the 3ʹ end of the different classes of endogenous and antiviral small RNAs as indicated. RNA was isolated from young adults after two days of infection with OrV. b, miRNA expression in cde-1 (tm1021) mutants as compared to wild type, samples as in a. Dots: individual miRNA; reads per million averaging two independent C. elegans culture plates. c, piRNAs and endogenous 22G-RNAs abundance in cde-1 (tm1021) mutants as compared to wild type, normalised to library size. Dots: independent C. elegans culture plate. Samples as in a.
Supplementary Figure 6 CDE-1-depleted animals show high expression of stress-response genes during OrV infection.
a, Fold change in the length of poly(A) tails (measured by TAIL-seq) in cde-1 mutants compared to wild type. RNA was isolated from young adults after two days of OrV infection. Turkey boxplot. b, Differential mRNA expression in cde-1 (tm1021) compared to wild type, two days of OrV infection (mRNA-seq). Turkey boxplot; dots: outliers.
a, Simplified workflow of 3ʹ RACE-seq of OrV RNA1 and OrV RNA2. b-c, Comparison between the viral load and the fraction of non-templated mono(U) tails at the 3ʹ end of OrV RNA1 and OrV RNA2, respectively, in strains as indicated. Dots: independent infection. Samples as in Figs. 3d,e and 4e. Source data
a, Protein level of the IAV NP measured by immunofluorescence (FACS). Error: SEM in three independent infections. b, Level of expression of the IAV NP mRNA normalized to Gapdh in MEF cells of different genotypes as indicated. Bars: average value; error: SEM; three independent infections. Source data
Supplementary Figures 1–8 and Supplementary Tables 1–3
About this article
Cite this article
Le Pen, J., Jiang, H., Di Domenico, T. et al. Terminal uridylyltransferases target RNA viruses as part of the innate immune system. Nat Struct Mol Biol 25, 778–786 (2018). https://doi.org/10.1038/s41594-018-0106-9
RNA stabilization by a poly(A) tail 3′-end binding pocket and other modes of poly(A)-RNA interaction
Biological Chemistry (2021)
WIREs RNA (2021)
Molecular Microbiology (2021)
Remembering your enemies: mechanisms of within‐generation and multigenerational immune priming in Caenorhabditis elegans
The FEBS Journal (2021)