Skip to content
naulon
Get started

No retraction, correction or concern found when last checked on Oct 17, 2026.

Most Caenorhabditis elegans microRNAs Are Individually Not Essential for Development or Viability

Eric A Miska, Ezequiel Alvarez-Saavedra, Allison L Abbott and 6 more

PLoS Genetics · 2007

doi:10.1371/journal.pgen.0030215

Abstract

MicroRNAs (miRNAs), a large class of short noncoding RNAs found in many plants and animals, often act to post-transcriptionally inhibit gene expression. We report the generation of deletion mutations in 87 miRNA genes in Caenorhabditis elegans, expanding the number of mutated miRNA genes to 95, or 83% of known C. elegans miRNAs. We find that the majority of miRNAs are not essential for the viability or development of C. elegans, and mutations in most miRNA genes do not result in grossly abnormal phenotypes. These observations are consistent with the hypothesis that there is significant functional redundancy among miRNAs or among gene pathways regulated by miRNAs. This study represents the first comprehensive genetic analysis of miRNA function in any organism and provides a unique, permanent resource for the systematic study of miRNAs.

Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 18, 2026. Paragraph numbers are ours, so an answer can point at one.

Introduction

MicroRNAs (miRNAs) were discovered in C. elegans during studies of the control of developmental timing. miRNAs are approximately 22-nucleotide noncoding RNAs that are thought to regulate gene expression through sequence-specific base-pairing with target mRNAs. miRNAs have been identified in organisms as diverse as roundworms, flies, fish, frogs, mammals, flowering plants, mosses, and even viruses, using genetics, molecular cloning, and predictions from bioinformatics. In C. elegans about 115 miRNA genes have been confidently identified.

In animals, miRNAs are transcribed as long RNA precursors (pri-miRNAs), which are processed in the nucleus by the RNase III enzyme complex Drosha-Pasha/DGCR8 to form the approximately 70-base pre-miRNAs or are derived directly from introns. Pre-miRNAs are exported from the nucleus by Exportin-5, processed by the RNase III enzyme Dicer, and incorporated into an Argonaute-containing RNA-induced silencing complex (RISC). Within the silencing complex, metazoan miRNAs pair to the mRNAs of protein-coding genes, usually through imperfect base-pairing with the 3′-UTR, thereby specifying the posttranscriptional repression of these target mRNAs. Binding of the silencing complex causes translational repression and/or mRNA destabilization, which is sometimes through direct mRNA cleavage, but usually through other mechanisms. Because many messages have been under selective pressure to preserve pairing to a 6mer in the 5′ region of the miRNA known as the miRNA seed (nucleotides 2–7), targets of metazoan miRNAs can be predicted above the background of false-positives by searching for conserved matches to the seed region. In nematodes, at least 10% of the protein-coding messages appear to be conserved targets of miRNAs.

The in vivo functions of a few miRNAs have been established. In C. elegans, the lin-4 miRNA and the let-7 family of miRNAs control the timing of aspects of larval development. For example, the lin-4 miRNA controls hypodermal cell-fate decisions during early larval development by negatively regulating the lin-14 and lin-28 mRNAs. The let-7 miRNA controls hypodermal cell-fate decisions during late-larval development by regulating the lin-41, hbl-1, daf-12, and pha-4 mRNAs. Three additional C. elegans let-7-like miRNAs, miR-48, miR-84, and miR-241, also act in the control of developmental timing and likely regulate the hbl-1 mRNA, but act earlier in development than the let-7 miRNA. The C. elegans lsy-6 miRNA acts in the asymmetric differentiation of the left and right ASE chemosensory neurons. Specifically, the lsy-6 miRNA targets the cog-1 mRNA, resulting in a shift of marker gene expression in the left ASE to resemble marker gene expression in the right ASE. The first miRNA studied functionally in Drosophila is encoded by the bantam locus, which had previously been identified in a screen for deregulated tissue growth. The bantam miRNA stimulates cell proliferation and reduces programmed cell death. bantam directly regulates the pro-apoptotic gene hid. A second Drosophila miRNA, miR-14, also reduces programmed cell death. The muscle-specific Drosophila miRNA miR-1 is required for larval development and cardiac differentiation. Dmir-7 regulates the transcription factor Yan. Finally, Drosophila miR-9a is required for sensory organ precursor specification, and Drosophila miR-278 is required for energy homeostasis. The first loss-of-function studies of miRNAs in the mouse have been reported demonstrating a role for miR-1 and miR-208 in cardiac growth in response to stress and miR-155/BIC in normal immune function.

miRNA function has also been inferred from studies in which miRNAs have been misexpressed in worms, flies, frogs, mice, and cultured mammalian cells. In addition, miRNA function has been explored by perturbing the functions of genes in the pathway for miRNA biogenesis and by reducing miRNA levels using antisense oligonucleotides. For example, mutants defective in Dicer, which is essential for miRNA biogenesis, have been studied for C. elegans, Drosophila, the zebrafish, and the mouse. In all cases, Dicer was found to be essential for normal development. In addition, members of the AGO subfamily of Argonaute proteins, which act in the miRNA pathway, are essential for normal C. elegans and mouse development.

In Drosophila, 2′ O-methyl antisense oligoribonucleotides have been used in miRNA depletion studies. This technique was initially described for human cells and C. elegans and appears to offer sequence-specific inhibition of small RNAs for a limited time span. Injection of individual 2′ O-methyl antisense oligoribonucleotides complementary to the 46 miRNAs known to be expressed in the fly embryo resulted in a total of 25 different abnormal phenotypes, including defects in patterning, morphogenesis, and cell survival. Knockdown of miRNAs using modified 2′ O-methyl antisense oligoribonucleotides also has been reported for the mouse. Very recently, a study reported the use of morpholinos to knockdown miRNA function in zebrafish and identified a role for miR-375 in pancreatic islet development.

To gain a broader understanding of miRNA function, we generated a collection of deletion mutants of the majority of known miRNA genes in C. elegans. We found that mutations in most miRNA genes do not result in striking abnormalities, and therefore most miRNA genes likely have subtle or redundant roles. This permanent collection provides a resource for detailed studies of miRNA function not possible previously.

Results

The cloning of many miRNAs from C. elegans using molecular biological techniques prompted us to take a genetic approach to study miRNA function in vivo in C. elegans through the generation of loss-of-function mutants. We isolated deletion mutants using established C. elegans techniques. We made extensive use of the “poison” primer method, which increases the sensitivity of detection of small deletions. Most C. elegans miRNAs were cloned and verified in northern blot experiments. Some miRNAs were predicted based on pre-miRNA folds and verified using northern blotting or PCR with specific primers and cloned miRNA libraries. The public database for miRNAs, miRBase release 9.0, listed 114 C. elegans miRNAs. Of these 114, 96 miRNAs are confidently identified, based on expression and the likelihood of being derived from stem-loop precursors, whereas many of the others do not appear to be authentic miRNAs. Recently, two studies using high-throughput sequencing methods identified 21 additional miRNAs bringing the total number of miRNAs identified with high confidence in C. elegans to 115 and the total number of annotated miRNA candidates to 135.

We isolated knockout mutants covering 87 miRNA genes. We previously described our studies of knockouts of three additional miRNA genes, and deletions in two other miRNA genes had been obtained by the C. elegans knockout consortium (D. Moerman, personal communication). Three miRNA genes had been mutated in genetic screens, lin-4, let-7, and lsy-6. Thus, 95 C. elegans miRNAs can now be functionally analyzed using mutants (Table 1). Additional alleles for a subset of these miRNA genes were also isolated by the C. elegans knockout consortium (D. Moerman, personal communication).

The median size of the deletions we isolated was 911 bases with a range of 181–6,288 bases (Tables 1 and S1). Some deletions likely affect neighboring genes in the case of intergenic miRNA genes or host genes in the case of miRNA genes found in introns. For example, the lethality linked to mir-50(n4099) (Table 2) might be a consequence of a loss-of-function of mir-50 or of an effect on the predicted host gene Y71G12B.11a (Table 1).

We performed a broad phenotypic study of all available miRNA loss-of-function mutants, including mutants that had been reported earlier. We focused on phenotypic assays that are relatively rapid and that examine C. elegans morphology, growth, development, and behavior. The assays we performed are shown in Table 3 and the phenotypes we observed are summarized in Table 2. Our initial phenotypic analysis revealed a single new abnormality linked to miRNA loss-of-function: deletion of the mir-240 mir-797 miRNA cluster resulted in abnormal defecation cycle lengths. This defecation defect was rescued by the introduction of a transgene carrying the mir-240 mir-797 genomic locus (Table S2). In addition, we observed other abnormal phenotypes. Mutation of the mir-35–41 miRNA cluster resulted in temperature-sensitive embryonic and larval lethality; this lethality was rescued by the introduction of a transgene carrying the mir-35–41 genomic locus (unpublished data). We were unable to generate homozygotes for alleles of mir-50 and mir-353. mir-50 and mir-353 are in introns of genes that when inactivated by RNAi result in embryonic lethality and may explain why we could not isolate homozygotes for our new deletions. Indeed, the introduction of a transgene carrying the mir-50 genomic locus failed to rescue the lethality associated with the mir-50 allele (unpublished data). The number of times each of the deletion strains has been outcrossed is shown in Table 2. It is conceivable that some of the miRNA deletion strains harbor additional mutations that suppress abnormalities conferred by miRNA deletion alleles and that could be revealed by outcrossing. To uncover subtle abnormalities in the miRNA mutant strains will require more detailed analyses, as has been performed for lin-4, let-7, lsy-6, mir-48, mir-84, and mir-241. Nevertheless, we note one striking conclusion: the majority of miRNAs are not essential for C. elegans viability and development.

Discussion

Here we report the first large-scale collection of miRNA loss-of-function mutants for any organism. We isolated new deletion alleles for 87 miRNA genes. Together with two publicly available deletion mutants, three mutants that we described elsewhere, and three mutants generated in genetic screens, there are now mutants for 95 C. elegans miRNA genes. We hope that this collection will become a widely used resource for the study of miRNA function.

Loss-of-Function versus Misexpression Studies

The overexpression of the miRNAs miR-84 and miR-61 from transgenes in C. elegans affects vulval development. The overexpression of miR-61 leads to the expression in Pn.p cells that do not normally generate vulval cell fates of reporter genes indicative of vulval cell fates. We examined if miR-61 and the closely related miR-247 were required for the normal induction of primary or secondary vulval cell fates by the Pn.p cells. We found that Pn.p cell induction was normal in mir-61 mutants and in mir-61; mir-247 double mutants (Table S3), although we did not test the effects of combining these mutants with mutants of mir-44 and mir-45, which have the same seed and thus are predicted to target the same messages. Similarly, let-60 RAS has been suggested to be a target of miR-84, based on the observation that overexpression of miR-84 from a transgene suppresses the multivulva phenotype of let-60 RAS activation mutants. If let-60 RAS is a target of miR-84, loss of mir-84 might result in let-60 RAS overexpression and possibly a multivulva phenotype. However, as we reported previously, mir-84 single mutants or mir-48 mir-241; mir-84 triple mutants do not have a multivulva phenotype. Thus, for both miR-84 and miR-61, we were unable to confirm a role in vulval development based on loss-of-function alleles. We conclude that these miRNAs are not required for vulval development and suggest that either they act redundantly with other miRNAs or other pathways in vulval development or they do not normally act in vulval development at all.

Redundancy of miRNAs and Their Regulatory Pathways

One difference between most protein-coding genes and most miRNA genes in C. elegans is the number of paralogs. Whereas fewer than 25% of protein-coding genes have a recognizable paralog in the C. elegans genome, about 60% of miRNAs are members of a family of two to eight genes. A higher number of paralogs might be a consequence of smaller gene size, which could allow a greater opportunity for gene duplication. As a consequence, miRNAs might act redundantly with other miRNAs and mutation of all paralogs of a miRNA or a miRNA family might result in synthetic abnormal synthetic phenotypes. Alternatively, some nematode miRNAs might act in parallel with other regulatory pathways that can compensate gene expression when the miRNAs are lost. For example, genetic data indicate that Drosophila mir-7 directly regulates the transcriptional repressor Yan in the fly eye, but that loss of mir-7 does not appreciably alter eye development, probably because of redundant protein turnover mechanisms that can also downregulate Yan. In such a scenario, disruptions in the other mechanisms would be needed to reveal the miRNA function.

Roles for Evolutionary Conserved miRNAs

The discovery that the let-7 miRNA is conserved among bilateria, including such disparate organisms as C. elegans and humans, appears not to have been an exception: for 15 miRNA families, miRNAs with identical seeds have been found in C. elegans, flies, fish, and mammals, and several additional families are predicted to be conserved throughout these diverse lineages. The conservation is not only for primary miRNA sequences, but also, at least in some cases, for patterns of expression. For example, the miRNA miR-1 is expressed in muscles of Drosophila, the zebrafish, and the mouse. However, the predicted mRNA targets of miRNAs might not share the same degree of conservation as miRNA expression patterns—the spectrum of predicted mRNA targets varies significantly among metazoans. With several miRNA loss-of-function mutants of Drosophila now available, we can begin to compare miRNA functions between C. elegans and Drosophila. Among the microRNAs for which mutations exist for flies and worms, Dmir-1 and C. elegans miR-1 are the most similar in sequence. Whereas Dmir-1 loss-of-function mutant fly larvae display muscle degeneration and die, we found that C. elegans miR-1 loss-of-function mutant animals are fully viable. Despite these differences, the mir-1 miRNA family could have a conserved role in muscle homeostasis and function. For example, the severity of the muscle defect of C. elegans mir-1 mutants might depend on physiological conditions, as is the case for the Dmir-1 mutant phenotypes of Drosophila.

We expect that as additional miRNA mutants become available for flies and other animals there will be future comparative studies of the biological functions of miRNAs using the collection of C. elegans miRNA mutants we have generated. More generally, we believe that the functions of miRNA genes, like the functions of protein-coding genes, will often prove to be conserved among animals, and that the collection of miRNA mutants we have generated will help define, test, and analyze general biological roles of miRNAs.

Nematode methods.

C. elegans was grown using standard conditions. The wild-type strain was var. Bristol N2. Nematodes were grown at 25 °C, except where otherwise indicated. Details about the mutant alleles we generated are shown in Table S1. All strains generated in this study have been submitted to the Caenorhabditis Genetics Center. Deletion allele information can be accessed directly from WormBase (http://www.wormbase.org).

Generation of deletion mutants.

Deletion mutants were isolated from a frozen library of worms mutagenized with ethyl methanesulphonate (EMS), 1,2,3,4-diepoxybutane (DEB), or a combination of UV irradiation and thymidine monophosphate (UV-TMP). In most instances, to enhance the detection of deletions one or two “poison” primers were included in the first round of nested PCR reactions. These poison primers were designed to anneal close to the mature miRNA sequence. In the first round of PCR, the three primers in the reaction (external forward, external reverse, and poison primers) generated both a full-length (from external primers) and a shorter product (from external and poison) from the wild-type allele. The shorter product was amplified more efficiently and thereby out-competed the amplification of full-length product. A deletion allele that removed the miRNA sequence and therefore removed the poison primer-binding site generated a product only from the external primers. In the second round of PCR, two internal primers designed just inside of the external primers amplified the full-length product but not the shorter product from the wild-type allele and a single product from the deletion allele. Mutant strains were outcrossed with the wild-type strain as indicated (Table S1).

Phenotypic analysis.

The minimum number of individual animals scored in each assay is given as n in parentheses below. (1) Locomotion: Number of body bends during a 20-s period were counted 4 min after transferring 1-d-old adult animals to fresh plates containing food (n = 10). (2) Pharyngeal pumping: Frequency of grinder displacement was counted for 20 s by eye, but otherwise as described previously (n = 5). (3) Defecation: The time between defecation cycles marked by posterior body muscle contraction events was measured (n = 3, 5 events per animal). (4) Egg laying: 1-d-old adult animals were lysed in bleaching solution for 10 min in the well of a round-bottom 96-well plate, and eggs were counted (n = 20). (5) Chemosensory neurons: L2 or L3 larvae were stained with DiO dye (Invitrogen) and filling of the neurons ASI, ASJ, ASH, ASK, AWS, ADL, PHA, PHB was scored (n = 15). (6) Cell number/nuclear morphology: L1 larvae were fixed and stained with 4′,6-diamidino-2-phenylindole, dihydrochloride (DAPI) (Invitrogen) as described previously. Nuclei of the ventral cord and intestine were counted (n = 15). (7) Dauer development: To assay dauer larva entry, three L4 animals were incubated at 25 °C until the F2/F3 progeny had been starved for at least five days. Animals were washed from plates using 1% SDS in de-ionized H2O for 30 min. Dauer larvae were identified by observing their thrashing and re-plated onto plates containing food to assay dauer exit. Constitutive dauer entry was scored by testing animals from plates with food for the presence of dauer larvae isolated after SDS treatment as described above (n = 50).

Accession Numbers

The miRNA sequences discussed in this paper can be found in the miRNA Registry (http://www.sanger.ac.uk/Software/Rfam/mirna/index.shtml). The C. elegans miRNA genes, their genomic location and deletion allele information can de accessed directly from WormBase (http://www.wormbase.org).

References 107

  1. 1.

    Ambros V, Horvitz HR. Heterochronic mutants of the nematode Caenorhabditis elegans. Science. 1984;226:409-416.

    Not in our library10.1126/science.6494891

  2. 2.

    Lee RC, Feinbaum RL, Ambros V. The C. elegans heterochronic gene lin-4 encodes small RNAs with antisense complementarity to lin-14. Cell. 1993;75:843-854.

    Not in our library10.1016/0092-8674(93)90529-y

  3. 3.

    Wightman B, Ha I, Ruvkun G. Posttranscriptional regulation of the heterochronic gene lin-14 by lin-4 mediates temporal pattern formation in C. elegans. Cell. 1993;75:855-862.

    Not in our library10.1016/0092-8674(93)90530-4

  4. 4.

    Reinhart BJ, Slack FJ, Basson M, Pasquinelli AE, Bettinger JC, et al. The 21-nucleotide let-7 RNA regulates developmental timing in Caenorhabditis elegans. Nature. 2000;403:901-906.

    Not in our library10.1038/35002607

  5. 5.

    Chalfie M, Horvitz HR, Sulston JE. Mutations that lead to reiterations in the cell lineages of C. elegans. Cell. 1981;24:59-69.

    Not in our library10.1016/0092-8674(81)90501-8

  6. 6.

    Bartel DP. MicroRNAs: genomics, biogenesis, mechanism, and function. Cell. 2004;116:281-297.

    Not in our library10.1016/s0092-8674(04)00045-5

  7. 7.

    Arazi T, Talmor-Neiman M, Stav R, Riese M, Huijser P, et al. Cloning and characterization of micro-RNAs from moss. Plant J. 2005;43:837-848.

    Not in our library10.1111/j.1365-313x.2005.02499.x

  8. 8.

    Axtell MJ, Bartel DP. Antiquity of microRNAs and their targets in land plants. Plant Cell. 2005;17:1658-1673.

    Not in our library10.1105/tpc.105.032185

  9. 9.

    Lagos-Quintana M, Rauhut R, Lendeckel W, Tuschl T. Identification of novel genes coding for small expressed RNAs. Science. 2001;294:853-858.

    Not in our library10.1126/science.1064921

  10. 10.

    Lau NC, Lim LP, Weinstein EG, Bartel DP. An abundant class of tiny RNAs with probable regulatory roles in Caenorhabditis elegans. Science. 2001;294:858-862.

    Not in our library10.1126/science.1065062

  11. 11.

    Lee RC, Ambros V. An extensive class of small RNAs in Caenorhabditis elegans. Science. 2001;294:862-864.

    Not in our library10.1126/science.1065329

  12. 12.

    Lim LP, Glasner ME, Yekta S, Burge CB, Bartel DP. Vertebrate microRNA genes. Science. 2003;299:1540.

    Not in our library10.1126/science.1080372

  13. 13.

    Llave C, Kasschau KD, Rector MA, Carrington JC. Endogenous and silencing-associated small RNAs in plants. Plant Cell. 2002;14:1605-1619.

    Not in our library10.1105/tpc.003210

  14. 14.

    Reinhart BJ, Weinstein EG, Rhoades MW, Bartel B, Bartel DP. MicroRNAs in plants. Genes Dev. 2002;16:1616-1626.

    Not in our library10.1101/gad.1004402

  15. 15.

    Watanabe T, Takeda A, Mise K, Okuno T, Suzuki T, et al. Stage-specific expression of microRNAs during Xenopus development. FEBS Lett. 2005;579:318-324.

    Not in our library10.1016/j.febslet.2004.11.067

  16. 16.

    Pfeffer S, Zavolan M, Grasser FA, Chien M, Russo JJ, et al. Identification of virus-encoded microRNAs. Science. 2004;304:734-736.

    Not in our library10.1126/science.1096781

  17. 17.

    Lim LP, Lau NC, Weinstein EG, Abdelhakim A, Yekta S, et al. The microRNAs of Caenorhabditis elegans. Genes Dev. 2003;17:991-1008.

    Not in our library10.1101/gad.1074403

  18. 18.

    Ohler U, Yekta S, Lim LP, Bartel DP, Burge CB. Patterns of flanking sequence conservation and a characteristic upstream motif for microRNA gene identification. RNA. 2004;10:1309-1322.

    Not in our library10.1261/rna.5206304

  19. 19.

    Ruby JG, Jan C, Player C, Axtell MJ, Lee W, et al. Large-scale sequencing reveals 21U-RNAs and additional microRNAs and endogenous siRNAs in C. elegans. Cell. 2006;127:1193-1207.

    Not in our library10.1016/j.cell.2006.10.040

  20. 20.

    Johnston RJ, Hobert O. A microRNA controlling left/right neuronal asymmetry in Caenorhabditis elegans. Nature. 2003;426:845-849.

    Not in our library10.1038/nature02255

  21. 21.

    Denli AM, Tops BB, Plasterk RH, Ketting RF, Hannon GJ. Processing of primary microRNAs by the Microprocessor complex. Nature. 2004;432:231-235.

    Not in our library10.1038/nature03049

  22. 22.

    Gregory RI, Yan KP, Amuthan G, Chendrimada T, Doratotaj B, et al. The Microprocessor complex mediates the genesis of microRNAs. Nature. 2004;432:235-240.

    Not in our library10.1038/nature03120

  23. 23.

    Han J, Lee Y, Yeom KH, Kim YK, Jin H, et al. The Drosha-DGCR8 complex in primary microRNA processing. Genes Dev. 2004;18:3016-3027.

    Not in our library10.1101/gad.1262504

  24. 24.

    Landthaler M, Yalcin A, Tuschl T. The human DiGeorge syndrome critical region gene 8 and its D. melanogaster homolog are required for miRNA biogenesis. Curr Biol. 2004;14:2162-2167.

    Not in our library10.1016/j.cub.2004.11.001

  25. 25.

    Lee Y, Ahn C, Han J, Choi H, Kim J, et al. The nuclear RNase III Drosha initiates microRNA processing. Nature. 2003;425:415-419.

    Not in our library10.1038/nature01957

  26. 26.

    Ruby JG, Jan CH, Bartel DP. Intronic microRNA precursors that bypass Drosha processing. Nature. 2007;448:83-86.

    Not in our library10.1038/nature05983

  27. 27.

    Okamura K, Hagen JW, Duan H, Tyler DM, Lai EC. The mirtron pathway generates microRNA-class regulatory RNAs in Drosophila. Cell. 2007;130:89-100.

    Not in our library10.1016/j.cell.2007.06.028

  28. 28.

    Lund E, Guttinger S, Calado A, Dahlberg JE, Kutay U. Nuclear export of microRNA precursors. Science. 2003;303:95-98.

    Not in our library10.1126/science.1090599

  29. 29.

    Du T, Zamore PD. microPrimer: the biogenesis and function of microRNA. Development. 2005;132:4645-4652.

    Not in our library10.1242/dev.02070

  30. 30.

    Pillai RS. MicroRNA function: multiple mechanisms for a tiny RNA? RNA. 2005;11:1753-1761.

    Not in our library10.1261/rna.2248605

  31. 31.

    Olsen PH, Ambros V. The lin-4 regulatory RNA controls developmental timing in Caenorhabditis elegans by blocking LIN-14 protein synthesis after the initiation of translation. Dev Biol. 1999;216:671-680.

    Not in our library10.1006/dbio.1999.9523

  32. 32.

    Petersen CP, Bordeleau ME, Pelletier J, Sharp PA. Short RNAs repress translation after initiation in mammalian cells. Mol Cell. 2006;21:533-542.

    Not in our library10.1016/j.molcel.2006.01.031

  33. 33.

    Seggerson K, Tang L, Moss EG. Two genetic circuits repress the Caenorhabditis elegans heterochronic gene lin-28 after translation initiation. Dev Biol. 2002;243:215-225.

    Not in our library10.1006/dbio.2001.0563

  34. 34.

    Yekta S, Shih IH, Bartel DP. MicroRNA-directed cleavage of HOXB8 mRNA. Science. 2004;304:594-596.

    Not in our library10.1126/science.1097434

  35. 35.

    Mansfield JH, Harfe BD, Nissen R, Obenauer J, Srineel J, et al. MicroRNA-responsive “sensor” transgenes uncover Hox-like and other developmentally regulated patterns of vertebrate microRNA expression. Nat Genet. 2004;36:1079-1083.

    Not in our library10.1038/ng1421

  36. 36.

    Bagga S, Bracht J, Hunter S, Massirer K, Holtz J, et al. Regulation by let-7 and lin-4 miRNAs results in target mRNA degradation. Cell. 2005;122:553-563.

    Not in our library10.1016/j.cell.2005.07.031

  37. 37.

    Jing Q, Huang S, Guth S, Zarubin T, Motoyama A, et al. Involvement of microRNA in AU-rich element-mediated mRNA instability. Cell. 2005;120:623-634.

    Not in our library10.1016/j.cell.2004.12.038

  38. 38.

    Giraldez AJ, Mishima Y, Rihel J, Grocock RJ, Van Dongen S, et al. Zebrafish MiR-430 promotes deadenylation and clearance of maternal mRNAs. Science. 2006;312:75-79.

    Not in our library10.1126/science.1122689

  39. 39.

    Lim LP, Lau NC, Garrett-Engele P, Grimson A, Schelter JM, et al. Microarray analysis shows that some microRNAs downregulate large numbers of target mRNAs. Nature. 2005;433:769-773.

    Not in our library10.1038/nature03315

  40. 40.

    Wu L, Fan J, Belasco JG. MicroRNAs direct rapid deadenylation of mRNA. Proc Natl Acad Sci U S A. 2006;103:4034-4039.

    Not in our library10.1073/pnas.0510928103

  41. 41.

    Lewis BP, Burge CB, Bartel DP. Conserved seed pairing, often flanked by adenosines, indicates that thousands of human genes are microRNA targets. Cell. 2005;120:15-20.

    Not in our library10.1016/j.cell.2004.12.035

  42. 42.

    Lewis BP, Shih IH, Jones-Rhoades MW, Bartel DP, Burge CB. Prediction of mammalian microRNA targets. Cell. 2003;115:787-798.

    Not in our library10.1016/s0092-8674(03)01018-3

  43. 43.

    Krek A, Grun D, Poy MN, Wolf R, Rosenberg L, et al. Combinatorial microRNA target predictions. Nat Genet. 2005;37:495-500.

    Not in our library10.1038/ng1536

  44. 44.
    Principles of MicroRNA–Target Recognition · 2005
    No notice
  45. 45.

    Xie X, Lu J, Kulbokas EJ, Golub TR, Mootha V, et al. Systematic discovery of regulatory motifs in human promoters and 3′ UTRs by comparison of several mammals. Nature. 2005;434:338-345.

    Not in our library10.1038/nature03441

  46. 46.

    Lall S, Grun D, Krek A, Chen K, Wang YL, et al. A genome-wide map of conserved microRNA targets in C. elegans. Curr Biol. 2006;16:460-471.

    Not in our library10.1016/j.cub.2006.01.050

  47. 47.

    Moss EG, Lee RC, Ambros V. The cold shock domain protein LIN-28 controls developmental timing in C. elegans and is regulated by the lin-4 RNA. Cell. 1997;88:637-646.

    Not in our library10.1016/s0092-8674(00)81906-6

  48. 48.

    Slack FJ, Basson M, Liu Z, Ambros V, Horvitz HR, et al. The lin-41 RBCC gene acts in the C. elegans heterochronic pathway between the let-7 regulatory RNA and the LIN-29 transcription factor. Mol Cell. 2000;5:659-669.

    Not in our library10.1016/s1097-2765(00)80245-2

  49. 49.

    Abrahante JE, Daul AL, Li M, Volk ML, Tennessen JM, et al. The Caenorhabditis elegans hunchback-like gene lin-57/hbl-1 controls developmental time and is regulated by microRNAs. Dev Cell. 2003;4:625-637.

    Not in our library10.1016/s1534-5807(03)00127-8

  50. 50.

    Grosshans H, Johnson T, Reinert KL, Gerstein M, Slack FJ. The temporal patterning microRNA let-7 regulates several transcription factors at the larval to adult transition in C. elegans. Dev Cell. 2005;8:321-330.

    Not in our library10.1016/j.devcel.2004.12.019

  51. 51.

    Lin SY, Johnson SM, Abraham M, Vella MC, Pasquinelli A, et al. The C. elegans hunchback homolog, hbl-1, controls temporal patterning and is a probable microRNA target. Dev Cell. 2003;4:639-650.

    Not in our library10.1016/s1534-5807(03)00124-2

  52. 52.

    Abbott AL, Alvarez-Saavedra E, Miska EA, Lau NC, Bartel DP, et al. The let-7 microRNA family members mir-48, mir-84, and mir-241 function together to regulate developmental timing in Caenorhabditis elegans. Dev Cell. 2005;9:403-414.

    Not in our library10.1016/j.devcel.2005.07.009

  53. 53.

    Li M, Jones-Rhoades MW, Lau NC, Bartel DP, Rougvie AE. Regulatory mutations of mir-48, a C. elegans let-7 family MicroRNA, cause developmental timing defects. Dev Cell. 2005;9:415-422.

    Not in our library10.1016/j.devcel.2005.08.002

  54. 54.

    Brennecke J, Hipfner DR, Stark A, Russell RB, Cohen SM. bantam encodes a developmentally regulated microRNA that controls cell proliferation and regulates the proapoptotic gene hid in Drosophila. Cell. 2003;113:25-36.

    Not in our library10.1016/s0092-8674(03)00231-9

  55. 55.

    Xu P, Vernooy SY, Guo M, Hay BA. The Drosophila microRNA mir-14 suppresses cell death and is required for normal fat metabolism. Curr Biol. 2003;13:790-795.

    Not in our library10.1016/s0960-9822(03)00250-1

  56. 56.

    Sokol NS, Ambros V. Mesodermally expressed Drosophila microRNA-1 is regulated by Twist and is required in muscles during larval growth. Genes Dev. 2005;19:2343-2354.

    Not in our library10.1101/gad.1356105

  57. 57.

    Kwon C, Han Z, Olson EN, Srivastava D. MicroRNA1 influences cardiac differentiation in Drosophila and regulates Notch signaling. Proc Natl Acad Sci U S A. 2005;102:18986-18991.

    Not in our library10.1073/pnas.0509535102

  58. 58.

    Li X, Carthew RW. A microRNA mediates EGF receptor signaling and promotes photoreceptor differentiation in the Drosophila eye. Cell. 2005;123:1267-1277.

    Not in our library10.1016/j.cell.2005.10.040

  59. 59.

    Li Y, Wang F, Lee JA, Gao FB. MicroRNA-9a ensures the precise specification of sensory organ precursors in Drosophila. Genes Dev. 2006;20:2793-2805.

    Not in our library10.1101/gad.1466306

  60. 60.

    Teleman AA, Maitra S, Cohen SM. Drosophila lacking microRNA miR-278 are defective in energy homeostasis. Genes Dev. 2006;20:417-422.

    Not in our library10.1101/gad.374406

  61. 61.

    van Rooij E, Sutherland LB, Qi X, Richardson JA, Hill J, et al. Control of stress-dependent cardiac growth and gene expression by a microRNA. Science. 2007;316:575-579.

    Not in our library10.1126/science.1139089

  62. 62.

    Zhao Y, Ransom JF, Li A, Vedantham V, von Drehle M, et al. Dysregulation of cardiogenesis, cardiac conduction, and cell cycle in mice lacking miRNA-1–2. Cell. 2007;129:303-317.

    Not in our library10.1016/j.cell.2007.03.030

  63. 63.

    Rodriguez A, Vigorito E, Clare S, Warren MV, Couttet P, et al. Requirement of bic/microRNA-155 for normal immune function. Science. 2007;316:608-611.

    Not in our library10.1126/science.1139253

  64. 64.

    Thai TH, Calado DP, Casola S, Ansel KM, Xiao C, et al. Regulation of the germinal center response by microRNA-155. Science. 2007;316:604-608.

    Not in our library10.1126/science.1141229

  65. 65.

    Miska EA. How microRNAs control cell division, differentiation, and death. Curr Opin Genet Dev. 2005;15:563-568.

    Not in our library10.1016/j.gde.2005.08.005

  66. 66.

    Knight SW, Bass BL. A role for the RNase III enzyme DCR-1 in RNA interference and germ line development in Caenorhabditis elegans. Science. 2001;293:2269-2271.

    Not in our library10.1126/science.1062039

  67. 67.

    Grishok A, Pasquinelli AE, Conte D, Li N, Parrish S, et al. Genes and mechanisms related to RNA interference regulate expression of the small temporal RNAs that control C. elegans developmental timing. Cell. 2001;106:23-34.

    Not in our library10.1016/s0092-8674(01)00431-7

  68. 68.

    Lee YS, Nakahara K, Pham JW, Kim K, He Z, et al. Distinct roles for Drosophila Dicer-1 and Dicer-2 in the siRNA/miRNA silencing pathways. Cell. 2004;117:69-81.

    Not in our library10.1016/s0092-8674(04)00261-2

  69. 69.

    Hatfield SD, Shcherbata HR, Fischer KA, Nakahara K, Carthew RW, et al. Stem cell division is regulated by the microRNA pathway. Nature. 2005;435:974-978.

    Not in our library10.1038/nature03816

  70. 70.

    Wienholds E, Koudijs MJ, van Eeden FJ, Cuppen E, Plasterk RH. The microRNA-producing enzyme Dicer1 is essential for zebrafish development. Nat Genet. 2003;35:217-218.

    Not in our library10.1038/ng1251

  71. 71.

    Giraldez AJ, Cinalli RM, Glasner ME, Enright AJ, Thomson JM, et al. MicroRNAs regulate brain morphogenesis in zebrafish. Science. 2005;308:833-838.

    Not in our library10.1126/science.1109020

  72. 72.

    Bernstein E, Kim SY, Carmell MA, Murchison EP, Alcorn H, et al. Dicer is essential for mouse development. Nat Genet. 2003;35:215-217.

    Not in our library10.1038/ng1253

  73. 73.

    Harfe BD, McManus MT, Mansfield JH, Hornstein E, Tabin CJ. The RNaseIII enzyme Dicer is required for morphogenesis but not patterning of the vertebrate limb. Proc Natl Acad Sci U S A. 2005;102:10898-10903.

    Not in our library10.1073/pnas.0504834102

  74. 74.

    Yang WJ, Yang DD, Na S, Sandusky GE, Zhang Q, et al. Dicer is required for embryonic angiogenesis during mouse development. J Biol Chem. 2005;280:9330-9335.

    Not in our library10.1074/jbc.m413394200

  75. 75.

    Harris KS, Zhang Z, McManus MT, Harfe BD, Sun X. Dicer function is essential for lung epithelium morphogenesis. Proc Natl Acad Sci U S A. 2006;103:2208-2213.

    Not in our library10.1073/pnas.0510839103

  76. 76.

    Liu J, Carmell MA, Rivas FV, Marsden CG, Thomson JM, et al. Argonaute2 is the catalytic engine of mammalian RNAi. Science. 2004;305:1437-1441.

    Not in our library10.1126/science.1102513

  77. 77.

    Leaman D, Chen PY, Fak J, Yalcin A, Pearce M, et al. Antisense-mediated depletion reveals essential and specific functions of microRNAs in Drosophila development. Cell. 2005;121:1097-1108.

    Not in our library10.1016/j.cell.2005.04.016

  78. 78.
    Sequence-Specific Inhibition of Small RNA Function · 2004
    No notice
  79. 79.

    Meister G, Landthaler M, Dorsett Y, Tuschl T. Sequence-specific inhibition of microRNA- and siRNA-induced RNA silencing. RNA. 2004;10:544-550.

    Not in our library10.1261/rna.5235104

  80. 80.

    Krutzfeldt J, Rajewsky N, Braich R, Rajeev KG, Tuschl T, et al. Silencing of microRNAs in vivo with “antagomirs.”. Nature. 2005;438:685-689.

    Not in our library10.1038/nature04303

  81. 81.
    Targeted Inhibition of miRNA Maturation with Morpholinos Reveals a Role for miR-375 in Pancreatic Islet Development · 2007
    Expression of concern
  82. 82.

    Jansen G, Hazendonk E, Thijssen KL, Plasterk RH. Reverse genetics by chemical mutagenesis in Caenorhabditis elegans. Nat Genet. 1997;17:119-121.

    Not in our library10.1038/ng0997-119

  83. 83.

    Liu LX, Spoerke JM, Mulligan EL, Chen J, Reardon B, et al. High-throughput isolation of Caenorhabditis elegans deletion mutants. Genome Res. 1999;9:859-867.

    Not in our library10.1101/gr.9.9.859

  84. 84.

    Edgley M, D'Souza A, Moulder G, McKay S, Shen B, et al. Improved detection of small deletions in complex pools of DNA. Nucleic Acids Res. 2002;30:e52.

    Not in our library10.1093/nar/gnf051

  85. 85.

    Ambros V, Lee RC, Lavanway A, Williams PT, Jewell D. MicroRNAs and other tiny endogenous RNAs in C. elegans. Curr Biol. 2003;13:807-818.

    Not in our library10.1016/s0960-9822(03)00287-2

  86. 86.

    Grad Y, Aach J, Hayes GD, Reinhart BJ, Church GM, et al. Computational and experimental identification of C. elegans microRNAs. Mol Cell. 2003;11:1253-1263.

    Not in our library10.1016/s1097-2765(03)00153-9

  87. 87.

    Griffiths-Jones S. The microRNA Registry. Nucleic Acids Res. 2004;32:D109-D111.

    Not in our library10.1093/nar/gkh023

  88. 88.

    Griffiths-Jones S, Grocock RJ, van Dongen S, Bateman A, Enright AJ. miRBase: microRNA sequences, targets, and gene nomenclature. Nucleic Acids Res. 2006;34:D140-D144.

    Not in our library10.1093/nar/gkj112

  89. 89.

    Yoo AS, Greenwald I. LIN-12/Notch activation leads to microRNA-mediated downregulation of Vav in C. elegans. Science. 2005;310:1330-1333.

    Not in our library10.1126/science.1119481

  90. 90.

    Johnson SM, Grosshans H, Shingara J, Byrom M, Jarvis R, et al. RAS is regulated by the let-7 microRNA family. Cell. 2005;120:635-647.

    Not in our library10.1016/j.cell.2005.01.014

  91. 91.

    Beitel GJ, Clark SG, Horvitz HR. Caenorhabditis elegans ras gene let-60 acts as a switch in the pathway of vulval induction. Nature. 1990;348:503-509.

    Not in our library10.1038/348503a0

  92. 92.

    Han M, Aroian RV, Sternberg PW. The let-60 locus controls the switch between vulval and nonvulval cell fates in Caenorhabditis elegans. Genetics. 1990;126:899-913.

    Not in our library10.1093/genetics/126.4.899

  93. 93.

    Genome sequence of the nematode C. elegans: a platform for investigating biology. Science. 1998;282:2012-2018.

    Not in our library10.1126/science.282.5396.2012

  94. 94.

    Pasquinelli AE, Reinhart BJ, Slack F, Martindale MQ, Kuroda MI, et al. Conservation of the sequence and temporal expression of let-7 heterochronic regulatory RNA. Nature. 2000;408:86-89.

    Not in our library10.1038/35040556

  95. 95.

    Ruby JG, Stark A, Johnston W, Kellis M, Bartel DP, et al. Biogenesis, expression, and target predictions for an expanded set of microRNA genes in Drosophila. Genome Res. 2007.

    Not in our library10.1101/gr.6597907

  96. 96.

    Sempere LF, Cole CN, McPeek MA, Peterson KJ. The phylogenetic distribution of metazoan microRNAs: insights into evolutionary complexity and constraint. J Exp Zoolog B Mol Dev Evol. 2006;306:575-588.

    Not in our library10.1002/jez.b.21118

  97. 97.

    Prochnik SE, Rokhsar DS, Aboobaker AA. Evidence for a microRNA expansion in the bilaterian ancestor. Dev Genes Evol. 2007;217:73-77.

    Not in our library10.1007/s00427-006-0116-1

  98. 98.

    Wienholds E, Kloosterman WP, Miska E, Alvarez-Saavedra E, Berezikov E, et al. MicroRNA expression in zebrafish embryonic development. Science. 2005;309:310-311.

    Not in our library10.1126/science.1114519

  99. 99.
    microRNA Target Predictions across Seven Drosophila Species and Comparison to Mammalian Targets · 2005
    No notice
  100. 100.

    Wood WB, Community of C. elegans Researchers. The nematode Caenorhabditis elegans. 1988.

    Not in our library

  101. 101.

    Brenner S. The genetics of Caenorhabditis elegans. Genetics. 1974;77:71-94.

    Not in our library10.1093/genetics/77.1.71

  102. 102.

    Avery L, Horvitz HR. A cell that dies during wild-type C. elegans development can function as a neuron in a ced-3 mutant. Cell. 1987;51:1071-1078.

    Not in our library10.1016/0092-8674(87)90593-9

  103. 103.

    Thomas JH. Genetic analysis of defecation in Caenorhabditis elegans. Genetics. 1990;124:855-872.

    Not in our library10.1093/genetics/124.4.855

  104. 104.

    Sawin ER. Genetic and cellular analysis of modulated behaviors in Caenorhabditis elegans. 1996.

    Not in our library

  105. 105.

    Fixsen WD. The genetic control of hypodermal cell lineages during nematode development. 1985.

    Not in our library

  106. 106.

    Horvitz HR, Sulston JE. Isolation and genetic characterization of cell-lineage mutants of the nematode Caenorhabditis elegans. Genetics. 1980;96:435-454.

    Not in our library10.1093/genetics/96.2.435

  107. 107.

    Schwarz EM, Antoshechkin I, Bastiani C, Bieri T, Blasiar D, et al. WormBase: better software, richer content. Nucleic Acids Res. 2006;34:D475-D478.

    Not in our library10.1093/nar/gkj061

Cited by papers we hold 13

Reads

Read by people on this page

None yet.

Checked or read by AI through naulon

None yet.

A person counts once a day per paper. We keep no cookie and no address: a reader is a hash that changes every day. Any month under five reads shows no number.

Are you an author?

Is this your paper?

Claim it to add your note and be paid when agents cite it. Free.