Meloidogyne incognita genes involved in the repellent behavior in response to ascr#9.
Meloidogyne incognita
Ascaroside
Dispersal
RNAi
Journal
Scientific reports
ISSN: 2045-2322
Titre abrégé: Sci Rep
Pays: England
ID NLM: 101563288
Informations de publication
Date de publication:
28 Oct 2024
28 Oct 2024
Historique:
received:
04
02
2024
accepted:
14
10
2024
medline:
28
10
2024
pubmed:
28
10
2024
entrez:
28
10
2024
Statut:
epublish
Résumé
Meloidogyne incognita is one of the globally serious plant parasitic nematodes. New control measure is urgently needed to replace the common chemical control method. Ascarosides are pheromones regulating the nematodes' aggregation, avoidance, mating, dispersal and dauer recovery and formation. Ascr#9, one of the ascarosides, exhibits the potential to repel M. incognita. However, the nematode genes involved in the perception of ascr# 9 signal are totally unknown. In this study, the transcriptome of ascr#9-treated second stage M. incognita juveniles (J2s) was analyzed, 44 pathways were significantly affected, multiple ligand-receptor and mucin type O-glycan were induced, and olfactory transduction was disturbed. A total of 11 highly differentially expressed genes involved in neuroactive ligand-receptor interaction and FMRFamide-like peptide related process were identified and knocked down by RNAi. The dispersal rates of M. incognita with three knocked-down genes (flp-14, mgl-1 and ADOR-1) significantly decreased, respectively, when ascr#9 was present. The results demonstrate that flp-14, mgl-1, and ADOR-1 are involved in the dispersal behavior of M. incognita nematodes responding to ascr#9, which promotes the interaction study between ascarosides and M. incognita, and provides new ideas for the prevention and control of M. incognita by using pheromone ascarosides.
Identifiants
pubmed: 39465253
doi: 10.1038/s41598-024-76370-5
pii: 10.1038/s41598-024-76370-5
doi:
Substances chimiques
Pheromones
0
Helminth Proteins
0
Types de publication
Journal Article
Langues
eng
Sous-ensembles de citation
IM
Pagination
25706Subventions
Organisme : Guangdong Provincial Science & Technology Project
ID : 2022A0505050051
Organisme : GDAS Special Project of Science and Technology Development
ID : 2022GDASZH-2022010101
Organisme : GDAS Special Project of Science and Technology Development
ID : 2022GDASZH-2022010101
Informations de copyright
© 2024. The Author(s).
Références
Jones, J. T. et al. Top 10 plant-parasitic nematodes in molecular plant pathology. Mol. Plant Pathol.14(9), 946–961. https://doi.org/10.1111/mpp.12057 (2013).
doi: 10.1111/mpp.12057
pubmed: 23809086
pmcid: 6638764
Coyne, D. L. et al. Plant-parasitic nematodes and food security in sub-saharan africa. Annu. Rev. Phytopathol.56, 381–403. https://doi.org/10.1146/annurev-phyto-080417-045833 (2018).
doi: 10.1146/annurev-phyto-080417-045833
pubmed: 29958072
pmcid: 7340484
Favery, B. et al. Gall-forming root-knot nematodes hijack key plant cellular functions to induce multinucleate and hypertrophied feeding cells. J. Insect. Physiol.84, 60–69. https://doi.org/10.1016/j.jinsphys.2015.07.013 (2016).
doi: 10.1016/j.jinsphys.2015.07.013
pubmed: 26211599
Juvale, P. S. & Baum, T. J. Cyst-ained research into Heterodera parasitism. PLoS Pathogens 14(2), e1006791 (2018). https://doi.org/10.1371/journal.ppat.1006791
Castagnone-Sereno, P. Genetic variability in parthenogenetic root-knot nematodes, Meloidogyne spp. and their ability to overcome plant resistance genes. Nematology. 4(5), 605–608. https://doi.org/10.1163/15685410260438872 (2002).
doi: 10.1163/15685410260438872
Khanal, C. et al. Identification and haplotype designation of Meloidogyne spp. of Arkansas using molecular diagnostics. Nematropica. 46(2), 261–270 (2016). https://www.researchgate.net/publication/314089197
Kepenekci, I. et al. Application methods of Steinernema feltiae, Xenorhabdus bovienii and purpureocillium lilacinum to control root-knot nematodes in greenhouse tomato systems. Crop Prot.108, 31–38. https://doi.org/10.1016/j.cropro.2018.02.009 (2018).
doi: 10.1016/j.cropro.2018.02.009
Ntalli, N. G. & Caboni, P. Botanical nematicides: A review. J. Agric. Food Chem.60, 9929–9940. https://doi.org/10.1021/jf303107j (2012).
doi: 10.1021/jf303107j
pubmed: 22973877
Abdel-Rahman, F. H. et al. Nematicidal activity of terpenoids. J. Environ. Sci. Health Part. B. 48(1), 16–22. https://doi.org/10.1080/03601234.2012.716686 (2013).
doi: 10.1080/03601234.2012.716686
Maleita, C. et al. Naphthoquinones from walnut husk residues show strong nematicidal activities against the root-knot nematode Meloidogyne hispanica. ACS Sustain. Chem. Eng.5(4), 3390–3398. https://doi.org/10.1021/acssuschemeng.7b00039 (2017).
doi: 10.1021/acssuschemeng.7b00039
Oota, M. et al. Identification of naturally occurring polyamines as root-knot nematode attractants. Mol. Plant. 13(4), 658–665. https://doi.org/10.1016/j.molp.2019.12.010 (2020).
doi: 10.1016/j.molp.2019.12.010
pubmed: 31891776
Brennan, P. A. & Zufall, F. Pheromonal communication in vertebrates. Nature. 444(7117), 308–315. https://doi.org/10.1038/nature05404 (2006).
doi: 10.1038/nature05404
pubmed: 17108955
Jeong, P. Y. et al. Chemical structure and biological activity of the Caenorhabditis elegans dauer-inducing pheromone. Nature. 433(7025), 541–545. https://doi.org/10.1038/nature03201 (2005).
doi: 10.1038/nature03201
pubmed: 15690045
Kim, B. et al. Root exudation by aphid leaf infestation recruits root-associated Paenibacillus spp. to lead plant insect susceptibility. J. Microbiol. Biotechnol.26(3), 549–557. https://doi.org/10.4014/jmb.1511.11058 (2016).
doi: 10.4014/jmb.1511.11058
pubmed: 26699743
Butcher, R. A. Decoding chemical communication in nematodes. Nat. Prod. Rep.34(5), 472–477. https://doi.org/10.1039/c7np00007c (2017).
doi: 10.1039/c7np00007c
pubmed: 28386618
pmcid: 5800508
von Reuss, S. H. Exploring modular glycolipids involved in nematode chemical communication. Chimia. 72, 297–303. https://doi.org/10.2533/chimia.2018.297 (2018).
doi: 10.2533/chimia.2018.297
Hartley, C. J. et al. Infective juveniles of entomopathogenic nematodes (Steinernema and Heterorhabditis) secrete ascarosides and respond to interspecific dispersal signals. J. Invertebr. Pathol.168 https://doi.org/10.1016/j.jip.2019.107257 (2019).
Oliveira-Hofman, C. et al. Pheromone extracts act as boosters for entomopathogenic nematodes efficacy. J. Invertebr. Pathol.164, 38–42. https://doi.org/10.1016/j.jip.2019.04.008 (2019).
doi: 10.1016/j.jip.2019.04.008
pubmed: 31034842
Wang, J. et al. Influence of the ascarosides on the recovery, yield and dispersal of entomopathogenic nematodes. J. Invertebr. Pathol.188 https://doi.org/10.1016/j.jip.2022.107717 (2022).
Dai, K. et al. Influence of entomopathogenic nematodes, symbiotic bacteriaand ascarosides on the dispersal behaviour of Meloidogyne incognita. Nematology. 0, 1–11. https://doi.org/10.1163/15685411-bja10184 (2022).
doi: 10.1163/15685411-bja10184
Kaplan, F. et al. Interspecific nematode signals regulate dispersal behavior. PloS One. 7(6). e38735 (2012).
Choe, A. et al. Ascaroside signaling is widely conserved among nematodes. Curr. Biol.22(9), 772–780. https://doi.org/10.1016/j.cub.2012.03.024 (2012).
doi: 10.1016/j.cub.2012.03.024
pubmed: 22503501
pmcid: 3360977
Manohar, M. et al. Plant metabolism of nematode pheromones mediates plant-nematode interactions. Nat. Commun.11(1), 208. https://doi.org/10.1038/s41467-019-14104-2 (2020).
doi: 10.1038/s41467-019-14104-2
pubmed: 31924834
pmcid: 6954178
Urwin, P. E., Lilley, C. J. & Atkinson, H. J. Ingestion of double-stranded RNA by preparasitic juvenile cyst nematodes leads to RNA interference. Mol. Plant-Microbe Int.15, 747–752. https://doi.org/10.1094/MPMI.2002.15.8.747 (2002).
doi: 10.1094/MPMI.2002.15.8.747
Chen, Q., Rehman, S., Smant, G. & Jones, J. T. Functional analysis of pathogenicity proteins of the potato cyst nematode Globodera rostochiensis using RNAi. Mol. Plant. Microbe Int.18, 621–625. https://doi.org/10.1094/mpmi-18-0621 (2005).
doi: 10.1094/mpmi-18-0621
Dong, L. et al. Lauric acid in crown daisy root exudate potently regulates root-knot nematode chemotaxis and disrupts Mi-flp-18 expression to block infection. J. Exp. Bot.65(1), 131–141. https://doi.org/10.1093/jxb/ert356 (2014).
doi: 10.1093/jxb/ert356
pubmed: 24170741
Fosu-Nyarko, J., Iqbal, S. & Jones, M. G. K. Targeting nematode genes by RNA silencing. Plant. Gene Silencing Mech. Appl.5, 176–192. https://doi.org/10.1079/9781780647678.0176 (2017).
doi: 10.1079/9781780647678.0176
Iqbal, S., Fosu-Nyarko, J. & Jones, M. G. K. Attempt to silence genes of the RNAi pathways of the Root-Knot Nematode, Meloidogyne incognita results in diverse responses including increase and no change in expression of some genes. Front. Plant. Sci.11, 328 (2020). Published 2020 Mar 24.
doi: 10.3389/fpls.2020.00328
pubmed: 32265973
pmcid: 7105803
Choi, I. et al. RNA-Seq of plant-parasitic nematode Meloidogyne incognita at various stages of its development. Front. Genet.8 https://doi.org/10.3389/fgene.2017.00190 (2017).
Meng, Q. P. et al. PCR assays for rapid and sensitive identification of three major root-knot nematodes, Meloidogyne incognita, M. Javanica and M. arenaria. Acta Phytopathologica Sinica. 34(3), 204–210 (2004).
Haas, B. J. et al. De novo transcript sequence reconstruction from RNA-seq using the Trinity platform for reference generation and analysis. Nat. Protoc.8, 1494–1512. https://doi.org/10.1038/nprot.2013.084 (2013).
doi: 10.1038/nprot.2013.084
pubmed: 23845962
Ashburner, M. et al. Gene Ontology: tool for the unification of biology. The Gene Ontology Consortium. Nat. Genet.25, 25–29. https://doi.org/10.1038/75556 (2000).
doi: 10.1038/75556
pubmed: 10802651
pmcid: 3037419
Kanehisa, M. & Goto, S. K. E. G. G. Kyoto Encyclopedia of genes and genomes. Nucleic Acids Res.28, 27–30. https://doi.org/10.1093/nar/28.1.27 (2000).
doi: 10.1093/nar/28.1.27
pubmed: 10592173
Li, B., Dewey, C. N. RSEM accurate transcript quantification from RNA-Seq data with or without a reference genome. BMC Bioinform.12, 323. https://doi.org/10.1186/1471-2105-12-323 (2011).
Robinson, M. D. et al. EdgeR: a bioconductor package for differential expression analysis of digital gene expression data. Bioinformatics. 26, 139–140. https://doi.org/10.1093/bioinformatics/btp616 (2010).
doi: 10.1093/bioinformatics/btp616
pubmed: 19910308
Conesa, A. et al. Blast2GO: a universal tool for annotation, visualization and analysis in functional genomics research. Bioinformatic. 21(18), 3674–3676. https://doi.org/10.1093/bioinformatics/bti610 (2005).
doi: 10.1093/bioinformatics/bti610
Xie, C. et al. KOBAS 2.0: a web server for annotation and identification of enriched pathways and diseases. Nucleic Acids Res.39, W316–W322. https://doi.org/10.1093/nar/gkr483 (2011).
doi: 10.1093/nar/gkr483
pubmed: 21715386
pmcid: 3125809
Papolu, P. K. et al. Utility of host delivered RNAi of two FMRF amide like peptides, flp-14 and flp-18, for the management of root knot nematode, Meloidogyne incognita. PLoS ONE. 8(11), e80603. https://doi.org/10.1371/journal.pone.0080603 (2013).
doi: 10.1371/journal.pone.0080603
pubmed: 24223228
pmcid: 3819290
Livak, K. J. & Schmittgen, T. D. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) method. Methods. 25, 402–408. https://doi.org/10.1006/meth.2001.1262 (2001).
doi: 10.1006/meth.2001.1262
pubmed: 11846609
Vandesompele, J. et al. Accurate normalization of real-time quantitative RT-PCR data by geometric averaging of multiple internal control genes. Genome Biol.3, 7, RESEARCH0034. https://doi.org/10.1186/gb-2002-3-7-research0034 (2002).
doi: 10.1186/gb-2002-3-7-research0034
pubmed: 12184808
pmcid: 126239
Dong, L. et al. Lauric acid in crown daisy root exudate potently regulates root-knot nematode chemotaxis and disrupts Mi-flp-18 expression to block infection. J. Exp. Bot.65(1), 131–141. https://doi.org/10.1093/jxb/ert356 (2014).
doi: 10.1093/jxb/ert356
pubmed: 24170741
Pierce, K. L. et al. Seven-transmembrane receptors. Nat. Rev. Mol. Cell Biol.3(9), 639–650. https://doi.org/10.1038/nrm908 (2002).
doi: 10.1038/nrm908
pubmed: 12209124
Chen, S. A. et al. Prey sensing and response in a nematode-trapping fungus is governed by the MAPK pheromone response pathway. Genetics. 217(2), iyaa008. https://doi.org/10.1093/genetics/iyaa008 (2021).
doi: 10.1093/genetics/iyaa008
pubmed: 33724405
Ferkey, D. M. et al. Chemosensory signal transduction in Caenorhabditis elegans. Genetics. 217(3), iyab004 (2021).
Macosko, E. Z. et al. A hub-and-spoke circuit drives pheromone attraction and social behaviour in C. Elegans. Nature. 458, 1171–1175. https://doi.org/10.1038/nature07886 (2009).
doi: 10.1038/nature07886
pubmed: 19349961
pmcid: 2760495
Jang, H. et al. Neuromodulatory state and sex specify alternative behaviors through antagonistic synaptic pathways in C. Elegans. Neuron. 75, 585–592. https://doi.org/10.1016/j.neuron.2012.06.034 (2012).
doi: 10.1016/j.neuron.2012.06.034
pubmed: 22920251
pmcid: 3462069
Park, D. et al. A conserved neuronal DAF-16/FoxO plays an important role in conveying pheromone signals to elicit repulsion behavior in Caenorhabditis elegans. Sci. Rep.7(1), 7260. https://doi.org/10.1038/s41598-017-07313-6 (2017).
doi: 10.1038/s41598-017-07313-6
pubmed: 28775361
pmcid: 5543152
Artyukhin, A. B. et al. Succinylated octopamine ascarosides and a new pathway of biogenic amine metabolism in Caenorhabditis elegans. J. Biol. Chem.288(26), 18778–18783. https://doi.org/10.1074/jbc.C113.477000 (2013).
doi: 10.1074/jbc.C113.477000
pubmed: 23689506
pmcid: 3696653
Chute, C. D. et al. Co-option of neurotransmitter signaling for inter-organismal communication in C. Elegans. Nat. Commun.10(1), 3186. https://doi.org/10.1038/s41467-019-11240-7 (2019).
doi: 10.1038/s41467-019-11240-7
pubmed: 31320626
pmcid: 6639374
Rex, E. et al. TYRA-2 (F01E11.5): A Caenorhabditis elegans tyramine receptor expressed in the MC and NSM pharyngeal neurons. J. Neurochem.94(1), 181–191. https://doi.org/10.1111/j.1471-4159.2005.03180.x (2005).
doi: 10.1111/j.1471-4159.2005.03180.x
pubmed: 15953361
Roderick, H. et al. Rational design of biosafe crop resistance to a range of nematodes using RNA interference. Plant Biotechnol. J.16(2), 520–529. https://doi.org/10.1111/pbi.12792 (2018).
doi: 10.1111/pbi.12792
pubmed: 28703405
Dalzell, J. J. et al. Non-nematode-derived double-stranded RNAs induce profound phenotypic changes in Meloidogyne incognita and Globodera pallida infective juveniles. Int. J. Parasitol.39, 1503–1516. https://doi.org/10.1016/j.ijpara.2009.05.006 (2009).
doi: 10.1016/j.ijpara.2009.05.006
pubmed: 19482028
Dalzell, J. J. et al. Short interfering RNA-mediated gene silencing in Globodera Pallida and Meloidogyne incognita infective stage juveniles. Int. J. Parasitol.40, 91–100. https://doi.org/10.1016/j.ijpara.2009.07.003 (2010).
doi: 10.1016/j.ijpara.2009.07.003
pubmed: 19651131
Lim, M. A. et al. Neuroendocrine modulation sustains the C. Elegans forward motor state. Elife. 5, e19887. https://doi.org/10.7554/eLife.19887 (2016).
doi: 10.7554/eLife.19887
pubmed: 27855782
pmcid: 5120884
Gershkovich, M. M. et al. Pharmacological and functional similarities of the human neuropeptide Y system in C. Elegans challenges phylogenetic views on the FLP/NPR system. Cell. Communication Signal.17(1), 123. https://doi.org/10.1186/s12964-019-0436-1 (2019).
doi: 10.1186/s12964-019-0436-1
Kamath, R. S. et al. Systematic functional analysis of the Caenorhabditis elegans genome using RNAi. Nature. 421, 231–237. https://doi.org/10.1038/nature01278 (2003).
doi: 10.1038/nature01278
pubmed: 12529635
Greer, E. R. et al. Neural and molecular dissection of a C. Elegans sensory circuit that regulates fat and feeding. Cell Metabol.8, 118–131. https://doi.org/10.1016/j.cmet.2008.06.005 (2008).
doi: 10.1016/j.cmet.2008.06.005
Luo, Z. et al. Obesogenic effect of erythromycin on Caenorhabditis elegans through over-eating and lipid metabolism disturbances. Environ. Pollut.294, 118615. https://doi.org/10.1016/j.envpol.2021.118615 (2022).
doi: 10.1016/j.envpol.2021.118615
pubmed: 34863891
Kang, C. & Avery, L. Systemic regulation of starvation response in Caenorhabditis elegans. Genes Dev.23(1), 12–17. https://doi.org/10.1101/gad.1723409 (2009).
doi: 10.1101/gad.1723409
pubmed: 19136622
pmcid: 2632168
Dillon, J. et al. Metabotropic glutamate receptors: modulators of context-dependent feeding behaviour in C. Elegans. J. Biol. Chem.290(24), 15052–15065. https://doi.org/10.1074/jbc.M114.606608 (2015).
doi: 10.1074/jbc.M114.606608
pubmed: 25869139
pmcid: 4463449
Ahmadi, M. & Roy, R. AMPK acts as a molecular trigger to coordinate glutamatergic signals and adaptive behaviours during acute starvation. Elife. 5, e16349. https://doi.org/10.7554/eLife.16349 (2016).
doi: 10.7554/eLife.16349
pubmed: 27642785
pmcid: 5028190
Jeong, H. & Paik, Y. K. MGL-1 on AIY neurons translates starvation to reproductive plasticity via neuropeptide signaling in Caenorhabditis elegans. Dev. Biol.430(1), 80–89. https://doi.org/10.1016/j.ydbio.2017.08.014 (2017).
doi: 10.1016/j.ydbio.2017.08.014
pubmed: 28807780
López-Cruz, A. et al. Parallel multimodal circuits control an innate foraging behavior. Neuron. 102(2), 407–419. https://doi.org/10.1016/j.neuron.2019.01.053 (2019).
doi: 10.1016/j.neuron.2019.01.053
pubmed: 30824353
pmcid: 9161785
De la Parra-Guerra, A. et al. Intergenerational toxicity of nonylphenol ethoxylate (NP-9) in Caenorhabditis elegans. Ecotoxicol. Environ. Saf.197 https://doi.org/10.1016/j.ecoenv.2020.110588 (2020).
Chai, C. M. et al. Interneuron control of C. Elegans developmental decision-making. Curr. Biol.32(10), 2316–2324. https://doi.org/10.1016/j.cub.2022.03.077 (2022).
doi: 10.1016/j.cub.2022.03.077
pubmed: 35447086
pmcid: 9270850
Ribeiro, J. A. Adenosine A2A receptor interactions with receptors for other neurotransmitters and neuromodulators. Eur. J. Pharmacol.375(1–3), 101–113. https://doi.org/10.1016/s0014-2999(99)00230-7 (1999).
doi: 10.1016/s0014-2999(99)00230-7
pubmed: 10443568
Friedman, B. et al. Adenosine A2A receptor signaling promotes FoxO associated autophagy in chondrocytes. Sci. Rep.11(1), 968. https://doi.org/10.1038/s41598-020-80244-x (2021).
doi: 10.1038/s41598-020-80244-x
pubmed: 33441836
pmcid: 7806643
Ling, C. et al. AdoR-1 (Adenosine Receptor) contributes to protection against paraquat-induced oxidative stress in Caenorhabditis elegans. Oxidative Med. Cell. Longev.1759009 https://doi.org/10.1155/2022/1759009 (2022).
da Silva, T. C. et al. Exogenous adenosine modulates behaviors and stress response in Caenorhabditis elegans. Neurochem. Res.48(1), 117–130. https://doi.org/10.1007/s11064-022-03727-5 (2023).
doi: 10.1007/s11064-022-03727-5
pubmed: 36018438
Bridi, J. C. et al. Lifespan extension induced by caffeine in Caenorhabditis elegans is partially dependent on adenosine signaling. Front. Aging Neurosci.7, 220. https://doi.org/10.3389/fnagi.2015.00220 (2015).
doi: 10.3389/fnagi.2015.00220
pubmed: 26696878
pmcid: 4672644
Arantes, L. P. et al. Mechanisms involved in anti-aging effects of guarana (Paullinia cupana) in Caenorhabditis elegans. Braz. J. Med. Biol. Res.51(9), e7552. https://doi.org/10.1590/1414-431X20187552 (2018).
doi: 10.1590/1414-431X20187552
pubmed: 29972429
pmcid: 6040867
Machado, M. L. et al. Ilex paraguariensis modulates fat metabolism in Caenorhabditis elegans through purinergic system (ADOR-1) and nuclear hormone receptor (NHR-49) pathways. PLoS ONE. 13(9), e0204023. https://doi.org/10.1371/journal.pone.0204023 (2018).
doi: 10.1371/journal.pone.0204023
pubmed: 30252861
pmcid: 6155532
Di Rocco, M. et al. Phenotypic assessment of pathogenic variants in GNAO1 and response to caffeine in C. Elegans models of the disease. Genes. 14(2), 319. https://doi.org/10.3390/genes14020319 (2023).
doi: 10.3390/genes14020319
pubmed: 36833246
pmcid: 9957173