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Apomixis and Its Agricultural Potential

Yıl 2025, Cilt: 8 Sayı: 3, 232 - 242, 23.12.2025
https://doi.org/10.38001/ijlsb.1682670

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Apomixis is a form of reproduction without fertilization in plants. It has been observed in more than 400 plant species, although it is absent in major crop plants. Apomixis is considered to be a powerful biotechnology tool for maintaining hybrid vigor across generations by producing seeds that are genetically identical to maternal plant. However, the molecular mechanisms underlying apomixis remain poorly understood. Numerous studies have been conducted with the aim of introducing apomict phenotypes into crop species. This review provides a brief overview of apomixis, its mechanisms and current applications.

Kaynakça

  • References 1. Barman, H.N., et al., Generation of a new thermo-sensitive genic male sterile rice line by targeted mutagenesis of TMS5 gene through CRISPR/Cas9 system. BMC Plant Biology, 2019. 19(1). https://doi.org/10.1186/s12870-019-1715-0
  • 2. Smith, J., XXXII. Notice of a Plant which produces perfect Seeds without any apparent Action of Pollen. Transactions of the Linnean Society of London, 1841. 18(4): p. 509-512. https://doi.org/10.1111/j.1095-8339.1838.tb00200.x
  • 3. Barcaccia, G., et al., A Reappraisal of the Evolutionary and Developmental Pathway of Apomixis and Its Genetic Control in Angiosperms. Genes, 2020. 11(8). https://doi.org/10.3390/genes11080859
  • 4. Drews, G.N., D. Lee, and C.A. Christensen, Genetic Analysis of Female Gametophyte Development and Function. The Plant Cell, 1998. 10(1): p. 5-17. DOI:10.1105/tpc.10.1.5
  • 5. Tucker, M.R. and A.M.G. Koltunow, Sexual and asexual (apomictic) seed development in flowering plants: molecular, morphological and evolutionary relationships. Functional Plant Biology, 2009. 36(6). https://doi.org/10.1071/FP09078
  • 6. Brewbaker, J.L., The Distribution and Phylogenetic Significance of Binucleate and Trinucleate Pollen Grains in the Angiosperms. American Journal of Botany, 1967. 54(9): p. 1069-1083. https://doi.org/10.1002/j.1537-2197.1967.tb10735.x
  • 7. Spillane, C., M.D. Curtis, and U. Grossniklaus, Apomixis technology development—virgin births in farmers' fields? Nature Biotechnology, 2004. 22(6): p. 687-691. https://doi.org/10.1038/nbt976
  • 8. Koltunow, A.M. and U. Grossniklaus, Apomixis: A Developmental Perspective. Annual Review of Plant Biology, 2003. 54(1): p. 547-574. https://doi.org/10.1146/annurev.arplant.54.110901.160842
  • 9. Bicknell, R.A., Understanding Apomixis: Recent Advances and Remaining Conundrums. The Plant Cell Online, 2004. 16(suppl_1): p. S228-S245. https://doi.org/10.1105/tpc.017921
  • 10. Koltunow, A.M., Apomixis: Embryo Sacs and Embryos Formed without Meiosis or Fertilization in Ovules. The Plant Cell, 1993: p. 1425-1437. https://doi.org/10.1105/tpc.5.10.1425
  • 11. Fei, X., et al., The steps from sexual reproduction to apomixis. Planta, 2019. 249(6): p. 1715-1730. https://doi.org/10.1007/s00425-019-03113-6
  • 12. Koltunow, A.M., et al., Anther, ovule, seed, and nucellar embryo development inCitrus sinensiscv. Valencia. Canadian Journal of Botany, 1995. 73(10): p. 1567-1582. https://doi.org/10.1139/b95-170
  • 13. Schmidt, A., Controlling Apomixis: Shared Features and Distinct Characteristics of Gene Regulation. Genes, 2020. 11(3). https://doi.org/10.3390/genes11030329
  • 14. Grimanelli, D., et al., Developmental genetics of gametophytic apomixis. Trends in Genetics, 2001. 17(10): p. 597-604. DOI: 10.1016/s0168-9525(01)02454-4
  • 15. Barrett, S.C.H., The Evolution, Maintenance, and Loss of Self-Incompatibility Systems, in Plant Reproductive Ecology. 1990. p. 98-124. https://doi.org/10.1093/oso/9780195063943.003.0005
  • 16. Boldrini, K.R., M.S. Pagliarini, and C.B. do Valle, Cell fusion and cytomixis during microsporogenesis in Brachiaria humidicola (Poaceae). South African Journal of Botany, 2006. 72(3): p. 478-481. https://doi.org/10.1016/j.sajb.2005.11.004
  • 17. Sharbel, T.F., et al., Molecular signatures of apomictic and sexual ovules in the Boechera holboellii complex. The Plant Journal, 2009. 58(5): p. 870-882. DOI: 10.1111/j.1365-313X.2009.03826.x
  • 18. Hörandl, E. and F. Hadacek, The oxidative damage initiation hypothesis for meiosis. Plant Reproduction, 2013. 26(4): p. 351-367. doi: 10.1007/s00497-013-0234-7
  • 19. Ozias-Akins, P. and P.J. van Dijk, Mendelian Genetics of Apomixis in Plants. Annual Review of Genetics, 2007. 41(1): p. 509-537. DOI: 10.1146/annurev.genet.40.110405.090511
  • 20. Pupilli, F. and G. Barcaccia, Cloning plants by seeds: Inheritance models and candidate genes to increase fundamental knowledge for engineering apomixis in sexual crops. Journal of Biotechnology, 2012. 159(4): p. 291-311. DOI: 10.1016/j.jbiotec.2011.08.028
  • 21. Barcaccia, G. and E. Albertini, Apomixis in plant reproduction: a novel perspective on an old dilemma. Plant Reproduction, 2013. 26(3): p. 159-179. doi: 10.1007/s00497-013-0222-y
  • 22. Hand, M.L. and A.M.G. Koltunow, The Genetic Control of Apomixis: Asexual Seed Formation. Genetics, 2014. 197(2): p. 441-450. DOI: 10.1534/genetics.114.163105
  • 23. Schallau, A., et al., Identification and genetic analysis of the APOSPORY locus in Hypericum perforatum L. The Plant Journal, 2010. 62(5): p. 773-784. DOI: 10.1111/j.1365-313X.2010.04188.x
  • 24. Noyes, R.D. and L.H. Rieseberg, Two Independent Loci Control Agamospermy (Apomixis) in the Triploid Flowering Plant Erigeron annuus. Genetics, 2000. 155(1): p. 379-390. DOI: 10.1093/genetics/155.1.379
  • 25. van Dijk, P.J. and J.M.T. Bakx-Schotman, Formation of Unreduced Megaspores (Diplospory) in Apomictic Dandelions (Taraxacum officinale, s.l.) Is Controlled by a Sex-Specific Dominant Locus. Genetics, 2004. 166(1): p. 483-492. DOI: 10.1534/genetics.166.1.483
  • 26. Vašut, R.J., et al., Fluorescent in situ hybridization shows DIPLOSPOROUS located on one of the NOR chromosomes in apomictic dandelions (Taraxacum) in the absence of a large hemizygous chromosomal region. Genome, 2014. 57(11/12): p. 609-620. DOI: 10.1139/gen-2014-0143
  • 27. Akiyama, Y., W.W. Hanna, and P. Ozias-Akins, High-resolution physical mapping reveals that the apospory-specific genomic region (ASGR) in Cenchrus ciliaris is located on a heterochromatic and hemizygous region of a single chromosome. Theoretical and Applied Genetics, 2005. 111(6): p. 1042-1051. DOI: 10.1007/s00122-005-0020-5
  • 28. Conner, J.A., et al., Sequence Analysis of Bacterial Artificial Chromosome Clones from the Apospory-Specific Genomic Region of Pennisetumand cenchrus Plant Physiology, 2008. 147(3): p. 1396-1411. DOI: 10.1104/pp.108.119081
  • 29. Goel, S., et al., Comparative Physical Mapping of the Apospory-Specific Genomic Region in Two Apomictic Grasses: Pennisetum squamulatum and Cenchrus ciliaris. Genetics, 2006. 173(1): p. 389-400. DOI: 10.1534/genetics.105.054429
  • 30. Galla, G., et al., A Portion of the Apomixis Locus of Paspalum Simplex is Microsyntenic with an Unstable Chromosome Segment Highly Conserved Among Poaceae. Scientific Reports, 2019. 9(1). https://doi.org/10.1038/s41598-019-39649-6
  • 31. Catanach, A.S., et al., Deletion mapping of genetic regions associated with apomixis in Hieracium. Proceedings of the National Academy of Sciences, 2006. 103(49): p. 18650-18655. DOI: 10.1073/pnas.0605588103
  • 32. Kotani, Y., et al., The LOSS OF APOMEIOSIS (LOA) locus in Hieracium praealtum can function independently of the associated large‐scale repetitive chromosomal structure. New Phytologist, 2013. 201(3): p. 973-981. DOI: 10.1111/nph.12574
  • 33. Okada, T., et al., Chromosomes Carrying Meiotic Avoidance Loci in Three Apomictic Eudicot Hieracium Subgenus Pilosella Species Share Structural Features with Two Monocot Apomicts Plant Physiology, 2011. 157(3): p. 1327-1341. https://doi.org/10.1104/pp.111.181164
  • 34. Mancini, M., et al., The MAP3K-Coding QUI-GON JINN (QGJ) Gene Is Essential to the Formation of Unreduced Embryo Sacs in Paspalum. Frontiers in Plant Science, 2018. 9. doi: 10.3389/fpls.2018.01547
  • 35. Podio, M., et al., A methylation status analysis of the apomixis-specific region in Paspalum spp. suggests an epigenetic control of parthenogenesis. Journal of Experimental Botany, 2014. 65(22): p. 6411-6424. DOI: 10.1093/jxb/eru354
  • 36. Siena, L.A., et al., An apomixis-linked ORC3-like pseudogene is associated with silencing of its functional homolog in apomictic Paspalum simplex. Journal of Experimental Botany, 2016. 67(6): p. 1965-1978. https://doi.org/10.1093/jxb/erw018
  • 37. Galla, G., et al., Ovule Gene Expression Analysis in Sexual and Aposporous Apomictic Hypericum perforatum L. (Hypericaceae) Accessions. Frontiers in Plant Science, 2019. 10. DOI: 10.3389/fpls.2019.00654
  • 38. Corral, J.M., et al., A Conserved Apomixis-Specific Polymorphism Is Correlated with Exclusive Exonuclease Expression in Premeiotic Ovules of Apomictic Boechera Species. Plant Physiology, 2013. 163(4): p. 1660-1672. DOI: 10.1104/pp.113.222430
  • 39. Mau, M., et al., Hybrid apomicts trapped in the ecological niches of their sexual ancestors. Proceedings of the National Academy of Sciences, 2015. 112(18). https://doi.org/10.1073/pnas.1423447112
  • 40. Mateo de Arias, M., et al., Whether Gametophytes Are Reduced or Unreduced in Angiosperms Might Be Determined Metabolically. Genes, 2020. 11(12). DOI: 10.3390/genes11121449
  • 41. Selva, J.P., et al., Genes Modulating the Increase in Sexuality in the Facultative Diplosporous Grass Eragrostis curvula under Water Stress Conditions. Genes, 2020. 11(9). DOI: 10.3390/genes11090969
  • 42. Wyder, S., et al., Differential gene expression profiling of one- and two-dimensional apogamous gametophytes of the fern Dryopteris affinis ssp. affinis. Plant Physiology and Biochemistry, 2020. 148: p. 302-311. DOI: 10.1016/j.plaphy.2020.01.021
  • 43. Fei, X., et al., Small RNA sequencing provides candidate miRNA-target pairs for revealing the mechanism of apomixis in Zanthoxylum bungeanum. BMC Plant Biology, 2021. 21(1). https://doi.org/10.1186/s12870-021-02935-5
  • 44. Klatt, S., et al., Photoperiod Extension Enhances Sexual Megaspore Formation and Triggers Metabolic Reprogramming in Facultative Apomictic Ranunculus auricomus. Frontiers in Plant Science, 2016. 7. DOI: 10.3389/fpls.2016.00278
  • 45. Ulum, F.B., C. Costa Castro, and E. Hörandl, Ploidy-Dependent Effects of Light Stress on the Mode of Reproduction in the Ranunculus auricomus Complex (Ranunculaceae). Frontiers in Plant Science, 2020. 11. DOI: 10.3389/fpls.2020.00104
  • 46. Chuong, E.B., N.C. Elde, and C. Feschotte, Regulatory activities of transposable elements: from conflicts to benefits. Nature Reviews Genetics, 2016. 18(2): p. 71-86. https://doi.org/10.1038/nrg.2016.139
  • 47. Martin, A., et al., A transposon-induced epigenetic change leads to sex determination in melon. Nature, 2009. 461(7267): p. 1135-1138. DOI: 10.1038/nature08498
  • 48. Ong-Abdullah, M., et al., Loss of Karma transposon methylation underlies the mantled somaclonal variant of oil palm. Nature, 2015. 525(7570): p. 533-537. DOI: 10.1038/nature15365
  • 49. Hojsgaard, D., Apomixis Technology: Separating the Wheat from the Chaff. Genes, 2020. 11(4). https://doi.org/10.3390/genes11040411
  • 50. Fiaz, S., et al., Apomixis and strategies to induce apomixis to preserve hybrid vigor for multiple generations. GM Crops & Food, 2020. 12(1): p. 57-70. DOI: 10.1080/21645698.2020.1808423
  • 51. Rathore, P., et al., Retro-Element Gypsy-163 Is Differentially Methylated in Reproductive Tissues of Apomictic and Sexual Plants of Cenchrus ciliaris. Frontiers in Genetics, 2020. 11. https://doi.org/10.3389/fgene.2020.00795
  • 52. Nonomura, K.-I., et al., The MSP1 Gene Is Necessary to Restrict the Number of Cells Entering into Male and Female Sporogenesis and to Initiate Anther Wall Formation in Rice. The Plant Cell, 2003. 15(8): p. 1728-1739. DOI: 10.1105/tpc.012401
  • 53. Brukhin, V., Molecular and genetic regulation of apomixis. Russian Journal of Genetics, 2017. 53(9): p. 943-964. DOI:10.1134/S1022795417090046
  • 54. Ozias-Akins, P., D. Roche, and W.W. Hanna, Tight clustering and hemizygosity of apomixis-linked molecular markers in Pennisetum squamulatum implies genetic control of apospory by a divergent locus that may have no allelic form in sexual genotypes. Proceedings of the National Academy of Sciences, 1998. 95(9): p. 5127-5132. DOI: 10.1073/pnas.95.9.5127
  • 55. Grimanelli, D., et al., Mapping diplosporous apomixis in tetraploid Tripsacum: one gene or several genes? Heredity, 1998. 80(1): p. 33-39. DOI: 10.1046/j.1365-2540.1998.00263.x
  • 56. Ozias-Akins, P., Y. Akiyama, and W.W. Hanna, Molecular characterization of the genomic region linked with apomixis in Pennisetum/Cenchrus. Functional & Integrative Genomics, 2003. 3(3): p. 94-104. DOI: 10.1007/s10142-003-0084-8
  • 57. Zappacosta, D., et al., A High-Density Linkage Map of the Forage Grass Eragrostis curvula and Localization of the Diplospory Locus. Frontiers in Plant Science, 2019. 10. https://doi.org/10.3389/fpls.2019.00918
  • 58. Olmedo-Monfil, V., et al., Control of female gamete formation by a small RNA pathway in Arabidopsis. Nature, 2010. 464(7288): p. 628-632. doi: 10.1038/nature08828.
  • 59. Underwood, C.J., et al., A PARTHENOGENESIS allele from apomictic dandelion can induce egg cell division without fertilization in lettuce. Nature Genetics, 2022. 54(1): p. 84-93. DOI: 10.1038/s41588-021-00984-y
  • 60. Dan, J., Xia, Y., Wang, Y., Zhan, Y., Tian, J., Tang, N., et al., One-line hybrid rice with high-efficiency synthetic apomixis a nd near-normal fertility. Plant Cell Reports, 2024. 43: p. 79. https://doi.org/10.1007/s00299-024-03154-6
  • 61. Vernet, A., Meynard, D., Lian, Q., Mieulet, D., Gibert, O., Bissah, M., et al., High-frequency synthetic apomixis in hybrid rice. Nature Communications, 2022. 13(1): p. 7963. https://doi.org/10.1038/s41467-022-35679-3
  • 62. Chen, B., Maas, L., Figueiredo, D., Zhong, Y., Reis, R., Li, M., et al., BABY BOOM regulates early embryo and endosperm development. Proceedings of the National Academy of Sciences, 2022. 119: p. e2201761119. https://doi.org/10.1073/pnas.2201761119
  • 63. Wang, Y., Fuentes, R.R., Van Rengs, W.M.J., Effgen, S., Zaidan, M.W.A.M., Franzen, R., et al., Harnessing clonal gametes in hybrid crops to engineer polyploid genomes. Nature Genetics, 2024. 56: p. 1075–1079. https://doi.org/10.1038/s41588-024-01750-6
  • 64. Zhang, C., Yang, Z., Tang, D., Zhu, Y., Wang, P., Li, D., et al., Genome design of hybrid potato. Cell, 2021. 184: p. 3873–3883.e12. DOI: 10.1016/j.cell.2021.06.006
  • 65. Mao, Y., Nakel, T., Erbasol Serbes, I., Joshi, S., Tekleyohans, D.G., Baum, T., and Gross-Hardt, R., ECS1 and ECS2 suppress polyspermy and the formation of haploid plants by promoting double fertilization. Elife, 2023. 12: p. e85832. DOI: 10.7554/eLife.85832
  • 66. Ravi, M., and Chan, S.W.L., Haploid plants produced by centromere-mediated genome elimination. Nature, 2010. 464: p. 615–618. DOI: 10.1038/nature08842
  • 67. Gilles, L.M., Khaled, A., Laffaire, J.B., Chaignon, S., Gendrot, G., Laplaige, J., Bergès, H., Beydon, G., Bayle, V., Barret, P., et al., Loss of pollen-specific phospholipase NOT LIKE DAD triggers gynogenesis in maize. EMBO Journal, 2017. 36: p. 707–717. DOI: 10.15252/embj.201796603
  • 68. Zhong, Y., Liu, C., Qi, X., Jiao, Y., Wang, D., Wang, Y., Liu, Z., Chen, C., Chen, B., Tian, X., et al., Mutation of ZmDMP enhances haploid induction in maize. Nature Plants, 2019. 5: p. 575–580. DOI: 10.1038/s41477-019-0443-7
  • 69. Conner, J.A., Podio, M., and Ozias-Akins, P., Haploid embryo production in rice and maize induced by PsASGR-BBML transgenes. Plant Reproduction, 2017. 30: p. 41–52. DOI: 10.1007/s00497-017-0298-x
  • 70. Huang, Y., Liang, Y., Xie, Y., Rao, Y., Xiong, J., Liu, C., Wang, C., Wang, X., Qian, Q., and Wang, K., Efficient haploid induction via egg cell expression of dandelion PARTHENOGENESIS in foxtail millet (Setaria italica). Plant Biotechnology Journal, 2024. DOI: 10.1111/pbi.14302
  • 71. d’Erfurth, I., Jolivet, S., Froger, N., Catrice, O., Novatchkova, M., and Mercier, R., Turning meiosis into mitosis. PLoS Biology, 2009. 7: p. e1000124. DOI: 10.1371/journal.pbio.1000124
  • 72. Wang, C., Liu, Q., Shen, Y., Hua, Y., Wang, J., Lin, J., et al., Clonal seeds from hybrid rice by simultaneous genome engineering of meiosis and fertilization genes. Nature Biotechnology, 2019. 37: p. 283–286. DOI: 10.1038/s41587-018-0003-0
  • 74. Liu, C., He, Z., Zhang, Y., Hu, F., Li, M., Liu, Q., et al., Synthetic apomixis enables stable transgenerational transmission of heterotic phenotypes in hybrid rice. Plant Communications, 2023. 4: p. 100470. https://doi.org/10.1016/j.xplc.2022.100470
  • 75. Boateng, K.A., et al., SWI1 is required for meiotic chromosome remodeling events. Mol Plant, 2008. 1(4): p. 620-33. DOI: 10.1093/mp/ssn030
  • 76. Hartung, F., et al., The catalytically active tyrosine residues of both SPO11-1 and SPO11-2 are required for meiotic double-strand break induction in Arabidopsis. Plant Cell, 2007. 19(10): p. 3090-9. doi: 10.1105/tpc.107.054817
  • 77. Grelon, M., et al., AtSPO11-1 is necessary for efficient meiotic recombination in plants. EMBO J, 2001. 20(3): p. 589-600. DOI: 10.1093/emboj/20.3.589
  • 78. Vrielynck, N., et al., A DNA topoisomerase VI-like complex initiates meiotic recombination. Science, 2016. 351(6276): p. 939-43. DOI: 10.1126/science.aad5196
  • 79. Zhang, C., et al., The Arabidopsis thaliana DSB formation (AtDFO) gene is required for meiotic double-strand break formation. Plant J, 2012. 72(2): p. 271-81. DOI: 10.1111/j.1365-313X.2012.05075.x
  • 80. De Muyt, A., et al., AtPRD1 is required for meiotic double strand break formation in Arabidopsis thaliana. EMBO J, 2007. 26(18): p. 4126-37. DOI: 10.1038/sj.emboj.7601815
  • 81. Gherbi, H., et al., Homologous recombination in planta is stimulated in the absence of Rad50. EMBO Rep, 2001. 2(4): p. 287-91. DOI: 10.1093/embo-reports/kve069
  • 82. Vannier, J.B., et al., Two roles for Rad50 in telomere maintenance. EMBO J, 2006. 25(19): p. 4577-85. DOI: 10.1038/sj.emboj.7601345
  • 83. Couteau, F., et al., Random chromosome segregation without meiotic arrest in both male and female meiocytes of a dmc1 mutant of Arabidopsis. Plant Cell, 1999. 11(9): p. 1623-34. DOI: 10.1105/tpc.11.9.1623
  • 84. Higgins, J.D., et al., The Arabidopsis MutS homolog AtMSH4 functions at an early step in recombination: evidence for two classes of recombination in Arabidopsis. Genes Dev, 2004. 18(20): p. 2557-70. doi: 10.1101/gad.317504
  • 85. Caryl, A.P., et al., A homologue of the yeast HOP1 gene is inactivated in the Arabidopsis meiotic mutant asy1. Chromosoma, 2000. 109(1-2): p. 62-71. DOI: 10.1007/s004120050413
  • 86. Watanabe, Y. and P. Nurse, Cohesin Rec8 is required for reductional chromosome segregation at meiosis. Nature, 1999. 400(6743): p. 461-4. DOI: 10.1038/22774
  • 87. Chelysheva, L., et al., AtREC8 and AtSCC3 are essential to the monopolar orientation of the kinetochores during meiosis. J Cell Sci, 2005. 118(Pt 20): p. 4621-32. DOI: 10.1242/jcs.02583
  • 88. Schommer, C., et al., AHP2 is required for bivalent formation and for segregation of homologous chromosomes in Arabidopsis meiosis. Plant J, 2003. 36(1): p. 1-11. DOI: 10.1046/j.1365-313x.2003.01850.x
  • 89. Cromer, L., et al., OSD1 promotes meiotic progression via APC/C inhibition and forms a regulatory network with TDM and CYCA1;2/TAM. PLoS Genet, 2012. 8(7): p. e1002865. DOI: 10.1371/journal.pgen.1002865
  • 90. Wang, Y., et al., Progression through meiosis I and meiosis II in Arabidopsis anthers is regulated by an A-type cyclin predominately expressed in prophase I. Plant Physiol, 2004. 136(4): p. 4127-35. DOI: 10.1104/pp.104.051201
  • 91. Cifuentes, M., et al., TDM1 Regulation Determines the Number of Meiotic Divisions. PLoS Genet, 2016. 12(2): p. e1005856. DOI: 10.1371/journal.pgen.1005856
  • 92. Guitton, A.E. and F. Berger, Loss of function of MULTICOPY SUPPRESSOR OF IRA 1 produces nonviable parthenogenetic embryos in Arabidopsis. Curr Biol, 2005. 15(8): p. 750-4. https://doi.org/10.1016/j.cub.2005.02.066
  • 93. Ravi, M. and S.W. Chan, Haploid plants produced by centromere-mediated genome elimination. Nature, 2010. 464(7288): p. 615-8. DOI: 10.1038/nature08842
  • 94. Ohad, N., et al., Mutations in FIE, a WD polycomb group gene, allow endosperm development without fertilization. Plant Cell, 1999. 11(3): p. 407-16. DOI: 10.1105/tpc.11.3.407
  • 95. Chaudhury, A.M., et al., Fertilization-independent seed development in Arabidopsis thaliana. Proc Natl Acad Sci U S A, 1997. 94(8): p. 4223-8. https://doi.org/10.1073/pnas.94.8.4223
  • 96. Bierzychudek, P., Patterns in plant parthenogenesis. Experientia, 1985. 41(10): p. 1255-1264. DOI: 10.1007/978-3-0348-6273-8_9
  • 97. Nygren, A., Apomixis in the angiosperms. II. The Botanical Review, 1954. 20(10): p. 577-649. 10.1007/978-3-642-95000-1_21
  • 98. Conner, J.A., et al., A parthenogenesis gene of apomict origin elicits embryo formation from unfertilized eggs in a sexual plant. Proc Natl Acad Sci U S A, 2015. 112(36): p. 11205-10. https://doi.org/10.1073/pnas.1505856112
  • 99. Khanday, I., et al., A male-expressed rice embryogenic trigger redirected for asexual propagation through seeds. Nature, 2018. 565(7737): p. 91-95. DOI: 10.1038/s41586-018-0785-8
  • 100. Vijverberg, K., et al., Genetic fine-mapping of DIPLOSPOROUS in Taraxacum (dandelion; Asteraceae) indicates a duplicated DIP-gene. BMC Plant Biology, 2010. 10(1). doi: 10.1186/1471-2229-10-154
  • 101. Bicknell, R., et al., Genetic mapping of the LOSS OF PARTHENOGENESIS locus in Pilosella piloselloides and the evolution of apomixis in the Lactuceae. Frontiers in Plant Science, 2023. 14. DOI: 10.3389/fpls.2023.1239191
  • 102. Brukhin, V., et al., The Boechera Genus as a Resource for Apomixis Research. Frontiers in Plant Science, 2019. 10. doi: 10.3389/fpls.2019.00392
  • 103. Gao, J., et al., Trimethylguanosine Synthase1 (TGS1) Is Essential for Chilling Tolerance. Plant Physiology, 2017. 174(3): p. 1713-1727. DOI: 10.1104/pp.17.00340
  • 104. Ortiz, J.P.A., et al., Small RNA-seq reveals novel regulatory components for apomixis in Paspalum notatum. BMC Genomics, 2019. 20(1). https://doi.org/10.1186/s12864-019-5881-0
  • 105. Henikoff, S. and Y. Dalal, Centromeric chromatin: what makes it unique? Current Opinion in Genetics & Development, 2005. 15(2): p. 177-184. DOI: 10.1016/j.gde.2005.01.004
  • 106. Nogler, G.A., Gametophytic Apomixis, in Embryology of Angiosperms. 1984. p. 475-518. doi:10.1007/978-3-642-69302-1_10
  • 107. Pellino, M., et al., Asexual genome evolution in the apomictic Ranunculus auricomus complex: examining the effects of hybridization and mutation accumulation. Molecular Ecology, 2013. 22(23): p. 5908-5921. DOI: 10.1111/mec.12533
  • 108. Kondrashov, A.S., Deleterious mutations and the evolution of sexual reproduction. Nature, 1988. 336(6198): p. 435-440. https://doi.org/10.1038/336435a0
  • 109. Muller, H.J., The relation of recombination to mutational advance. Mutation Research/Fundamental and Molecular Mechanisms of Mutagenesis, 1964. 1(1): p. 2-9. https://doi.org/10.1016/0027-5107(64)90047-8
  • 110. Kimura, M., T. Maruyama, and J.F. Crow, The Mutation Load in Small Populations. Genetics, 1963. 48(10): p. 1303-1312. doi: 10.1093/genetics/48.10.1303
  • 111. Felsenstein, J., The Evolutionary Advantage of Recombination. Genetics, 1974. 78(2): p. 737-756. DOI: 10.1093/genetics/78.2.737
  • 112. Hill, W.G. and A. Robertson, The effect of linkage on limits to artificial selection. Genetical Research, 2009. 8(3): p. 269-294. DOI: https://doi.org/10.1017/S0016672300010156
  • 113. Maynard Smith, J., The evolution of sex. 1978, Cambridge Eng. ; New York: Cambridge University Press. x, 222 p. ISBN: 9780521293020
  • 114. Bell, G., The masterpiece of nature : the evolution and genetics of sexuality. 1982, Berkeley: University of California Press. 635 p. https://doi.org/10.4324/9780429322884
  • 115. Trivers, R., The Evolution of SexThe Masterpiece of Nature: The Evolution and Genetics of Sexuality.Graham Bell. The Quarterly Review of Biology, 1983. 58(1): p. 62-67. DOI:10.1086/413059
  • 116. Paun, O., et al., Patterns, sources and ecological implications of clonal diversity in apomictic Ranunculus carpaticola (Ranunculus auricomus complex, Ranunculaceae). Molecular Ecology, 2006. 15(4): p. 897-910. DOI: 10.1111/j.1365-294X.2006.02800.x
  • 117. Koch, M.A., Multiple Hybrid Formation in Natural Populations: Concerted Evolution of the Internal Transcribed Spacer of Nuclear Ribosomal DNA (ITS) in North American Arabis divaricarpa (Brassicaceae). Molecular Biology and Evolution, 2003. 20(3): p. 338-350. DOI: 10.1093/molbev/msg046
  • 118. Otto, S.P. and J. Whitton, Polyploid Incidence and Evolution. Annual Review of Genetics, 2000. 34(1): p. 401-437. DOI: 10.1146/annurev.genet.34.1.401
  • 119. Hojsgaard, D. and E. HÃrandl, A little bit of sex matters for genome evolution in asexual plants. Frontiers in Plant Science, 2015. 6. DOI: 10.3389/fpls.2015.00082

Apomiksi ve apomiksinin tarımsal potansiyali

Yıl 2025, Cilt: 8 Sayı: 3, 232 - 242, 23.12.2025
https://doi.org/10.38001/ijlsb.1682670

Öz

Apomiksi, bitkilerde döllenme olmadan gerçekleşen bir üreme biçimidir. 400'den fazla bitki türünde gözlemlenmiştir, ancak önemli mahsul bitkilerinde yoktur. Apomiksi, genetik olarak ana bitkiyle aynı olan tohumlar üreterek nesiller boyunca melez canlılığı sürdürmek için güçlü bir biyoteknoloji aracı olarak kabul edilir. Ancak, apomiksinin altında yatan moleküler mekanizmalar yeterince anlaşılmamıştır. Mahsul türlerine apomiktik fenotipleri tanıtmak amacıyla çok sayıda çalışma yürütülmüştür. Bu inceleme, apomiksi, mekanizmaları ve güncel uygulamaları hakkında kısa bir genel bakış sunmaktadır.

Kaynakça

  • References 1. Barman, H.N., et al., Generation of a new thermo-sensitive genic male sterile rice line by targeted mutagenesis of TMS5 gene through CRISPR/Cas9 system. BMC Plant Biology, 2019. 19(1). https://doi.org/10.1186/s12870-019-1715-0
  • 2. Smith, J., XXXII. Notice of a Plant which produces perfect Seeds without any apparent Action of Pollen. Transactions of the Linnean Society of London, 1841. 18(4): p. 509-512. https://doi.org/10.1111/j.1095-8339.1838.tb00200.x
  • 3. Barcaccia, G., et al., A Reappraisal of the Evolutionary and Developmental Pathway of Apomixis and Its Genetic Control in Angiosperms. Genes, 2020. 11(8). https://doi.org/10.3390/genes11080859
  • 4. Drews, G.N., D. Lee, and C.A. Christensen, Genetic Analysis of Female Gametophyte Development and Function. The Plant Cell, 1998. 10(1): p. 5-17. DOI:10.1105/tpc.10.1.5
  • 5. Tucker, M.R. and A.M.G. Koltunow, Sexual and asexual (apomictic) seed development in flowering plants: molecular, morphological and evolutionary relationships. Functional Plant Biology, 2009. 36(6). https://doi.org/10.1071/FP09078
  • 6. Brewbaker, J.L., The Distribution and Phylogenetic Significance of Binucleate and Trinucleate Pollen Grains in the Angiosperms. American Journal of Botany, 1967. 54(9): p. 1069-1083. https://doi.org/10.1002/j.1537-2197.1967.tb10735.x
  • 7. Spillane, C., M.D. Curtis, and U. Grossniklaus, Apomixis technology development—virgin births in farmers' fields? Nature Biotechnology, 2004. 22(6): p. 687-691. https://doi.org/10.1038/nbt976
  • 8. Koltunow, A.M. and U. Grossniklaus, Apomixis: A Developmental Perspective. Annual Review of Plant Biology, 2003. 54(1): p. 547-574. https://doi.org/10.1146/annurev.arplant.54.110901.160842
  • 9. Bicknell, R.A., Understanding Apomixis: Recent Advances and Remaining Conundrums. The Plant Cell Online, 2004. 16(suppl_1): p. S228-S245. https://doi.org/10.1105/tpc.017921
  • 10. Koltunow, A.M., Apomixis: Embryo Sacs and Embryos Formed without Meiosis or Fertilization in Ovules. The Plant Cell, 1993: p. 1425-1437. https://doi.org/10.1105/tpc.5.10.1425
  • 11. Fei, X., et al., The steps from sexual reproduction to apomixis. Planta, 2019. 249(6): p. 1715-1730. https://doi.org/10.1007/s00425-019-03113-6
  • 12. Koltunow, A.M., et al., Anther, ovule, seed, and nucellar embryo development inCitrus sinensiscv. Valencia. Canadian Journal of Botany, 1995. 73(10): p. 1567-1582. https://doi.org/10.1139/b95-170
  • 13. Schmidt, A., Controlling Apomixis: Shared Features and Distinct Characteristics of Gene Regulation. Genes, 2020. 11(3). https://doi.org/10.3390/genes11030329
  • 14. Grimanelli, D., et al., Developmental genetics of gametophytic apomixis. Trends in Genetics, 2001. 17(10): p. 597-604. DOI: 10.1016/s0168-9525(01)02454-4
  • 15. Barrett, S.C.H., The Evolution, Maintenance, and Loss of Self-Incompatibility Systems, in Plant Reproductive Ecology. 1990. p. 98-124. https://doi.org/10.1093/oso/9780195063943.003.0005
  • 16. Boldrini, K.R., M.S. Pagliarini, and C.B. do Valle, Cell fusion and cytomixis during microsporogenesis in Brachiaria humidicola (Poaceae). South African Journal of Botany, 2006. 72(3): p. 478-481. https://doi.org/10.1016/j.sajb.2005.11.004
  • 17. Sharbel, T.F., et al., Molecular signatures of apomictic and sexual ovules in the Boechera holboellii complex. The Plant Journal, 2009. 58(5): p. 870-882. DOI: 10.1111/j.1365-313X.2009.03826.x
  • 18. Hörandl, E. and F. Hadacek, The oxidative damage initiation hypothesis for meiosis. Plant Reproduction, 2013. 26(4): p. 351-367. doi: 10.1007/s00497-013-0234-7
  • 19. Ozias-Akins, P. and P.J. van Dijk, Mendelian Genetics of Apomixis in Plants. Annual Review of Genetics, 2007. 41(1): p. 509-537. DOI: 10.1146/annurev.genet.40.110405.090511
  • 20. Pupilli, F. and G. Barcaccia, Cloning plants by seeds: Inheritance models and candidate genes to increase fundamental knowledge for engineering apomixis in sexual crops. Journal of Biotechnology, 2012. 159(4): p. 291-311. DOI: 10.1016/j.jbiotec.2011.08.028
  • 21. Barcaccia, G. and E. Albertini, Apomixis in plant reproduction: a novel perspective on an old dilemma. Plant Reproduction, 2013. 26(3): p. 159-179. doi: 10.1007/s00497-013-0222-y
  • 22. Hand, M.L. and A.M.G. Koltunow, The Genetic Control of Apomixis: Asexual Seed Formation. Genetics, 2014. 197(2): p. 441-450. DOI: 10.1534/genetics.114.163105
  • 23. Schallau, A., et al., Identification and genetic analysis of the APOSPORY locus in Hypericum perforatum L. The Plant Journal, 2010. 62(5): p. 773-784. DOI: 10.1111/j.1365-313X.2010.04188.x
  • 24. Noyes, R.D. and L.H. Rieseberg, Two Independent Loci Control Agamospermy (Apomixis) in the Triploid Flowering Plant Erigeron annuus. Genetics, 2000. 155(1): p. 379-390. DOI: 10.1093/genetics/155.1.379
  • 25. van Dijk, P.J. and J.M.T. Bakx-Schotman, Formation of Unreduced Megaspores (Diplospory) in Apomictic Dandelions (Taraxacum officinale, s.l.) Is Controlled by a Sex-Specific Dominant Locus. Genetics, 2004. 166(1): p. 483-492. DOI: 10.1534/genetics.166.1.483
  • 26. Vašut, R.J., et al., Fluorescent in situ hybridization shows DIPLOSPOROUS located on one of the NOR chromosomes in apomictic dandelions (Taraxacum) in the absence of a large hemizygous chromosomal region. Genome, 2014. 57(11/12): p. 609-620. DOI: 10.1139/gen-2014-0143
  • 27. Akiyama, Y., W.W. Hanna, and P. Ozias-Akins, High-resolution physical mapping reveals that the apospory-specific genomic region (ASGR) in Cenchrus ciliaris is located on a heterochromatic and hemizygous region of a single chromosome. Theoretical and Applied Genetics, 2005. 111(6): p. 1042-1051. DOI: 10.1007/s00122-005-0020-5
  • 28. Conner, J.A., et al., Sequence Analysis of Bacterial Artificial Chromosome Clones from the Apospory-Specific Genomic Region of Pennisetumand cenchrus Plant Physiology, 2008. 147(3): p. 1396-1411. DOI: 10.1104/pp.108.119081
  • 29. Goel, S., et al., Comparative Physical Mapping of the Apospory-Specific Genomic Region in Two Apomictic Grasses: Pennisetum squamulatum and Cenchrus ciliaris. Genetics, 2006. 173(1): p. 389-400. DOI: 10.1534/genetics.105.054429
  • 30. Galla, G., et al., A Portion of the Apomixis Locus of Paspalum Simplex is Microsyntenic with an Unstable Chromosome Segment Highly Conserved Among Poaceae. Scientific Reports, 2019. 9(1). https://doi.org/10.1038/s41598-019-39649-6
  • 31. Catanach, A.S., et al., Deletion mapping of genetic regions associated with apomixis in Hieracium. Proceedings of the National Academy of Sciences, 2006. 103(49): p. 18650-18655. DOI: 10.1073/pnas.0605588103
  • 32. Kotani, Y., et al., The LOSS OF APOMEIOSIS (LOA) locus in Hieracium praealtum can function independently of the associated large‐scale repetitive chromosomal structure. New Phytologist, 2013. 201(3): p. 973-981. DOI: 10.1111/nph.12574
  • 33. Okada, T., et al., Chromosomes Carrying Meiotic Avoidance Loci in Three Apomictic Eudicot Hieracium Subgenus Pilosella Species Share Structural Features with Two Monocot Apomicts Plant Physiology, 2011. 157(3): p. 1327-1341. https://doi.org/10.1104/pp.111.181164
  • 34. Mancini, M., et al., The MAP3K-Coding QUI-GON JINN (QGJ) Gene Is Essential to the Formation of Unreduced Embryo Sacs in Paspalum. Frontiers in Plant Science, 2018. 9. doi: 10.3389/fpls.2018.01547
  • 35. Podio, M., et al., A methylation status analysis of the apomixis-specific region in Paspalum spp. suggests an epigenetic control of parthenogenesis. Journal of Experimental Botany, 2014. 65(22): p. 6411-6424. DOI: 10.1093/jxb/eru354
  • 36. Siena, L.A., et al., An apomixis-linked ORC3-like pseudogene is associated with silencing of its functional homolog in apomictic Paspalum simplex. Journal of Experimental Botany, 2016. 67(6): p. 1965-1978. https://doi.org/10.1093/jxb/erw018
  • 37. Galla, G., et al., Ovule Gene Expression Analysis in Sexual and Aposporous Apomictic Hypericum perforatum L. (Hypericaceae) Accessions. Frontiers in Plant Science, 2019. 10. DOI: 10.3389/fpls.2019.00654
  • 38. Corral, J.M., et al., A Conserved Apomixis-Specific Polymorphism Is Correlated with Exclusive Exonuclease Expression in Premeiotic Ovules of Apomictic Boechera Species. Plant Physiology, 2013. 163(4): p. 1660-1672. DOI: 10.1104/pp.113.222430
  • 39. Mau, M., et al., Hybrid apomicts trapped in the ecological niches of their sexual ancestors. Proceedings of the National Academy of Sciences, 2015. 112(18). https://doi.org/10.1073/pnas.1423447112
  • 40. Mateo de Arias, M., et al., Whether Gametophytes Are Reduced or Unreduced in Angiosperms Might Be Determined Metabolically. Genes, 2020. 11(12). DOI: 10.3390/genes11121449
  • 41. Selva, J.P., et al., Genes Modulating the Increase in Sexuality in the Facultative Diplosporous Grass Eragrostis curvula under Water Stress Conditions. Genes, 2020. 11(9). DOI: 10.3390/genes11090969
  • 42. Wyder, S., et al., Differential gene expression profiling of one- and two-dimensional apogamous gametophytes of the fern Dryopteris affinis ssp. affinis. Plant Physiology and Biochemistry, 2020. 148: p. 302-311. DOI: 10.1016/j.plaphy.2020.01.021
  • 43. Fei, X., et al., Small RNA sequencing provides candidate miRNA-target pairs for revealing the mechanism of apomixis in Zanthoxylum bungeanum. BMC Plant Biology, 2021. 21(1). https://doi.org/10.1186/s12870-021-02935-5
  • 44. Klatt, S., et al., Photoperiod Extension Enhances Sexual Megaspore Formation and Triggers Metabolic Reprogramming in Facultative Apomictic Ranunculus auricomus. Frontiers in Plant Science, 2016. 7. DOI: 10.3389/fpls.2016.00278
  • 45. Ulum, F.B., C. Costa Castro, and E. Hörandl, Ploidy-Dependent Effects of Light Stress on the Mode of Reproduction in the Ranunculus auricomus Complex (Ranunculaceae). Frontiers in Plant Science, 2020. 11. DOI: 10.3389/fpls.2020.00104
  • 46. Chuong, E.B., N.C. Elde, and C. Feschotte, Regulatory activities of transposable elements: from conflicts to benefits. Nature Reviews Genetics, 2016. 18(2): p. 71-86. https://doi.org/10.1038/nrg.2016.139
  • 47. Martin, A., et al., A transposon-induced epigenetic change leads to sex determination in melon. Nature, 2009. 461(7267): p. 1135-1138. DOI: 10.1038/nature08498
  • 48. Ong-Abdullah, M., et al., Loss of Karma transposon methylation underlies the mantled somaclonal variant of oil palm. Nature, 2015. 525(7570): p. 533-537. DOI: 10.1038/nature15365
  • 49. Hojsgaard, D., Apomixis Technology: Separating the Wheat from the Chaff. Genes, 2020. 11(4). https://doi.org/10.3390/genes11040411
  • 50. Fiaz, S., et al., Apomixis and strategies to induce apomixis to preserve hybrid vigor for multiple generations. GM Crops & Food, 2020. 12(1): p. 57-70. DOI: 10.1080/21645698.2020.1808423
  • 51. Rathore, P., et al., Retro-Element Gypsy-163 Is Differentially Methylated in Reproductive Tissues of Apomictic and Sexual Plants of Cenchrus ciliaris. Frontiers in Genetics, 2020. 11. https://doi.org/10.3389/fgene.2020.00795
  • 52. Nonomura, K.-I., et al., The MSP1 Gene Is Necessary to Restrict the Number of Cells Entering into Male and Female Sporogenesis and to Initiate Anther Wall Formation in Rice. The Plant Cell, 2003. 15(8): p. 1728-1739. DOI: 10.1105/tpc.012401
  • 53. Brukhin, V., Molecular and genetic regulation of apomixis. Russian Journal of Genetics, 2017. 53(9): p. 943-964. DOI:10.1134/S1022795417090046
  • 54. Ozias-Akins, P., D. Roche, and W.W. Hanna, Tight clustering and hemizygosity of apomixis-linked molecular markers in Pennisetum squamulatum implies genetic control of apospory by a divergent locus that may have no allelic form in sexual genotypes. Proceedings of the National Academy of Sciences, 1998. 95(9): p. 5127-5132. DOI: 10.1073/pnas.95.9.5127
  • 55. Grimanelli, D., et al., Mapping diplosporous apomixis in tetraploid Tripsacum: one gene or several genes? Heredity, 1998. 80(1): p. 33-39. DOI: 10.1046/j.1365-2540.1998.00263.x
  • 56. Ozias-Akins, P., Y. Akiyama, and W.W. Hanna, Molecular characterization of the genomic region linked with apomixis in Pennisetum/Cenchrus. Functional & Integrative Genomics, 2003. 3(3): p. 94-104. DOI: 10.1007/s10142-003-0084-8
  • 57. Zappacosta, D., et al., A High-Density Linkage Map of the Forage Grass Eragrostis curvula and Localization of the Diplospory Locus. Frontiers in Plant Science, 2019. 10. https://doi.org/10.3389/fpls.2019.00918
  • 58. Olmedo-Monfil, V., et al., Control of female gamete formation by a small RNA pathway in Arabidopsis. Nature, 2010. 464(7288): p. 628-632. doi: 10.1038/nature08828.
  • 59. Underwood, C.J., et al., A PARTHENOGENESIS allele from apomictic dandelion can induce egg cell division without fertilization in lettuce. Nature Genetics, 2022. 54(1): p. 84-93. DOI: 10.1038/s41588-021-00984-y
  • 60. Dan, J., Xia, Y., Wang, Y., Zhan, Y., Tian, J., Tang, N., et al., One-line hybrid rice with high-efficiency synthetic apomixis a nd near-normal fertility. Plant Cell Reports, 2024. 43: p. 79. https://doi.org/10.1007/s00299-024-03154-6
  • 61. Vernet, A., Meynard, D., Lian, Q., Mieulet, D., Gibert, O., Bissah, M., et al., High-frequency synthetic apomixis in hybrid rice. Nature Communications, 2022. 13(1): p. 7963. https://doi.org/10.1038/s41467-022-35679-3
  • 62. Chen, B., Maas, L., Figueiredo, D., Zhong, Y., Reis, R., Li, M., et al., BABY BOOM regulates early embryo and endosperm development. Proceedings of the National Academy of Sciences, 2022. 119: p. e2201761119. https://doi.org/10.1073/pnas.2201761119
  • 63. Wang, Y., Fuentes, R.R., Van Rengs, W.M.J., Effgen, S., Zaidan, M.W.A.M., Franzen, R., et al., Harnessing clonal gametes in hybrid crops to engineer polyploid genomes. Nature Genetics, 2024. 56: p. 1075–1079. https://doi.org/10.1038/s41588-024-01750-6
  • 64. Zhang, C., Yang, Z., Tang, D., Zhu, Y., Wang, P., Li, D., et al., Genome design of hybrid potato. Cell, 2021. 184: p. 3873–3883.e12. DOI: 10.1016/j.cell.2021.06.006
  • 65. Mao, Y., Nakel, T., Erbasol Serbes, I., Joshi, S., Tekleyohans, D.G., Baum, T., and Gross-Hardt, R., ECS1 and ECS2 suppress polyspermy and the formation of haploid plants by promoting double fertilization. Elife, 2023. 12: p. e85832. DOI: 10.7554/eLife.85832
  • 66. Ravi, M., and Chan, S.W.L., Haploid plants produced by centromere-mediated genome elimination. Nature, 2010. 464: p. 615–618. DOI: 10.1038/nature08842
  • 67. Gilles, L.M., Khaled, A., Laffaire, J.B., Chaignon, S., Gendrot, G., Laplaige, J., Bergès, H., Beydon, G., Bayle, V., Barret, P., et al., Loss of pollen-specific phospholipase NOT LIKE DAD triggers gynogenesis in maize. EMBO Journal, 2017. 36: p. 707–717. DOI: 10.15252/embj.201796603
  • 68. Zhong, Y., Liu, C., Qi, X., Jiao, Y., Wang, D., Wang, Y., Liu, Z., Chen, C., Chen, B., Tian, X., et al., Mutation of ZmDMP enhances haploid induction in maize. Nature Plants, 2019. 5: p. 575–580. DOI: 10.1038/s41477-019-0443-7
  • 69. Conner, J.A., Podio, M., and Ozias-Akins, P., Haploid embryo production in rice and maize induced by PsASGR-BBML transgenes. Plant Reproduction, 2017. 30: p. 41–52. DOI: 10.1007/s00497-017-0298-x
  • 70. Huang, Y., Liang, Y., Xie, Y., Rao, Y., Xiong, J., Liu, C., Wang, C., Wang, X., Qian, Q., and Wang, K., Efficient haploid induction via egg cell expression of dandelion PARTHENOGENESIS in foxtail millet (Setaria italica). Plant Biotechnology Journal, 2024. DOI: 10.1111/pbi.14302
  • 71. d’Erfurth, I., Jolivet, S., Froger, N., Catrice, O., Novatchkova, M., and Mercier, R., Turning meiosis into mitosis. PLoS Biology, 2009. 7: p. e1000124. DOI: 10.1371/journal.pbio.1000124
  • 72. Wang, C., Liu, Q., Shen, Y., Hua, Y., Wang, J., Lin, J., et al., Clonal seeds from hybrid rice by simultaneous genome engineering of meiosis and fertilization genes. Nature Biotechnology, 2019. 37: p. 283–286. DOI: 10.1038/s41587-018-0003-0
  • 74. Liu, C., He, Z., Zhang, Y., Hu, F., Li, M., Liu, Q., et al., Synthetic apomixis enables stable transgenerational transmission of heterotic phenotypes in hybrid rice. Plant Communications, 2023. 4: p. 100470. https://doi.org/10.1016/j.xplc.2022.100470
  • 75. Boateng, K.A., et al., SWI1 is required for meiotic chromosome remodeling events. Mol Plant, 2008. 1(4): p. 620-33. DOI: 10.1093/mp/ssn030
  • 76. Hartung, F., et al., The catalytically active tyrosine residues of both SPO11-1 and SPO11-2 are required for meiotic double-strand break induction in Arabidopsis. Plant Cell, 2007. 19(10): p. 3090-9. doi: 10.1105/tpc.107.054817
  • 77. Grelon, M., et al., AtSPO11-1 is necessary for efficient meiotic recombination in plants. EMBO J, 2001. 20(3): p. 589-600. DOI: 10.1093/emboj/20.3.589
  • 78. Vrielynck, N., et al., A DNA topoisomerase VI-like complex initiates meiotic recombination. Science, 2016. 351(6276): p. 939-43. DOI: 10.1126/science.aad5196
  • 79. Zhang, C., et al., The Arabidopsis thaliana DSB formation (AtDFO) gene is required for meiotic double-strand break formation. Plant J, 2012. 72(2): p. 271-81. DOI: 10.1111/j.1365-313X.2012.05075.x
  • 80. De Muyt, A., et al., AtPRD1 is required for meiotic double strand break formation in Arabidopsis thaliana. EMBO J, 2007. 26(18): p. 4126-37. DOI: 10.1038/sj.emboj.7601815
  • 81. Gherbi, H., et al., Homologous recombination in planta is stimulated in the absence of Rad50. EMBO Rep, 2001. 2(4): p. 287-91. DOI: 10.1093/embo-reports/kve069
  • 82. Vannier, J.B., et al., Two roles for Rad50 in telomere maintenance. EMBO J, 2006. 25(19): p. 4577-85. DOI: 10.1038/sj.emboj.7601345
  • 83. Couteau, F., et al., Random chromosome segregation without meiotic arrest in both male and female meiocytes of a dmc1 mutant of Arabidopsis. Plant Cell, 1999. 11(9): p. 1623-34. DOI: 10.1105/tpc.11.9.1623
  • 84. Higgins, J.D., et al., The Arabidopsis MutS homolog AtMSH4 functions at an early step in recombination: evidence for two classes of recombination in Arabidopsis. Genes Dev, 2004. 18(20): p. 2557-70. doi: 10.1101/gad.317504
  • 85. Caryl, A.P., et al., A homologue of the yeast HOP1 gene is inactivated in the Arabidopsis meiotic mutant asy1. Chromosoma, 2000. 109(1-2): p. 62-71. DOI: 10.1007/s004120050413
  • 86. Watanabe, Y. and P. Nurse, Cohesin Rec8 is required for reductional chromosome segregation at meiosis. Nature, 1999. 400(6743): p. 461-4. DOI: 10.1038/22774
  • 87. Chelysheva, L., et al., AtREC8 and AtSCC3 are essential to the monopolar orientation of the kinetochores during meiosis. J Cell Sci, 2005. 118(Pt 20): p. 4621-32. DOI: 10.1242/jcs.02583
  • 88. Schommer, C., et al., AHP2 is required for bivalent formation and for segregation of homologous chromosomes in Arabidopsis meiosis. Plant J, 2003. 36(1): p. 1-11. DOI: 10.1046/j.1365-313x.2003.01850.x
  • 89. Cromer, L., et al., OSD1 promotes meiotic progression via APC/C inhibition and forms a regulatory network with TDM and CYCA1;2/TAM. PLoS Genet, 2012. 8(7): p. e1002865. DOI: 10.1371/journal.pgen.1002865
  • 90. Wang, Y., et al., Progression through meiosis I and meiosis II in Arabidopsis anthers is regulated by an A-type cyclin predominately expressed in prophase I. Plant Physiol, 2004. 136(4): p. 4127-35. DOI: 10.1104/pp.104.051201
  • 91. Cifuentes, M., et al., TDM1 Regulation Determines the Number of Meiotic Divisions. PLoS Genet, 2016. 12(2): p. e1005856. DOI: 10.1371/journal.pgen.1005856
  • 92. Guitton, A.E. and F. Berger, Loss of function of MULTICOPY SUPPRESSOR OF IRA 1 produces nonviable parthenogenetic embryos in Arabidopsis. Curr Biol, 2005. 15(8): p. 750-4. https://doi.org/10.1016/j.cub.2005.02.066
  • 93. Ravi, M. and S.W. Chan, Haploid plants produced by centromere-mediated genome elimination. Nature, 2010. 464(7288): p. 615-8. DOI: 10.1038/nature08842
  • 94. Ohad, N., et al., Mutations in FIE, a WD polycomb group gene, allow endosperm development without fertilization. Plant Cell, 1999. 11(3): p. 407-16. DOI: 10.1105/tpc.11.3.407
  • 95. Chaudhury, A.M., et al., Fertilization-independent seed development in Arabidopsis thaliana. Proc Natl Acad Sci U S A, 1997. 94(8): p. 4223-8. https://doi.org/10.1073/pnas.94.8.4223
  • 96. Bierzychudek, P., Patterns in plant parthenogenesis. Experientia, 1985. 41(10): p. 1255-1264. DOI: 10.1007/978-3-0348-6273-8_9
  • 97. Nygren, A., Apomixis in the angiosperms. II. The Botanical Review, 1954. 20(10): p. 577-649. 10.1007/978-3-642-95000-1_21
  • 98. Conner, J.A., et al., A parthenogenesis gene of apomict origin elicits embryo formation from unfertilized eggs in a sexual plant. Proc Natl Acad Sci U S A, 2015. 112(36): p. 11205-10. https://doi.org/10.1073/pnas.1505856112
  • 99. Khanday, I., et al., A male-expressed rice embryogenic trigger redirected for asexual propagation through seeds. Nature, 2018. 565(7737): p. 91-95. DOI: 10.1038/s41586-018-0785-8
  • 100. Vijverberg, K., et al., Genetic fine-mapping of DIPLOSPOROUS in Taraxacum (dandelion; Asteraceae) indicates a duplicated DIP-gene. BMC Plant Biology, 2010. 10(1). doi: 10.1186/1471-2229-10-154
  • 101. Bicknell, R., et al., Genetic mapping of the LOSS OF PARTHENOGENESIS locus in Pilosella piloselloides and the evolution of apomixis in the Lactuceae. Frontiers in Plant Science, 2023. 14. DOI: 10.3389/fpls.2023.1239191
  • 102. Brukhin, V., et al., The Boechera Genus as a Resource for Apomixis Research. Frontiers in Plant Science, 2019. 10. doi: 10.3389/fpls.2019.00392
  • 103. Gao, J., et al., Trimethylguanosine Synthase1 (TGS1) Is Essential for Chilling Tolerance. Plant Physiology, 2017. 174(3): p. 1713-1727. DOI: 10.1104/pp.17.00340
  • 104. Ortiz, J.P.A., et al., Small RNA-seq reveals novel regulatory components for apomixis in Paspalum notatum. BMC Genomics, 2019. 20(1). https://doi.org/10.1186/s12864-019-5881-0
  • 105. Henikoff, S. and Y. Dalal, Centromeric chromatin: what makes it unique? Current Opinion in Genetics & Development, 2005. 15(2): p. 177-184. DOI: 10.1016/j.gde.2005.01.004
  • 106. Nogler, G.A., Gametophytic Apomixis, in Embryology of Angiosperms. 1984. p. 475-518. doi:10.1007/978-3-642-69302-1_10
  • 107. Pellino, M., et al., Asexual genome evolution in the apomictic Ranunculus auricomus complex: examining the effects of hybridization and mutation accumulation. Molecular Ecology, 2013. 22(23): p. 5908-5921. DOI: 10.1111/mec.12533
  • 108. Kondrashov, A.S., Deleterious mutations and the evolution of sexual reproduction. Nature, 1988. 336(6198): p. 435-440. https://doi.org/10.1038/336435a0
  • 109. Muller, H.J., The relation of recombination to mutational advance. Mutation Research/Fundamental and Molecular Mechanisms of Mutagenesis, 1964. 1(1): p. 2-9. https://doi.org/10.1016/0027-5107(64)90047-8
  • 110. Kimura, M., T. Maruyama, and J.F. Crow, The Mutation Load in Small Populations. Genetics, 1963. 48(10): p. 1303-1312. doi: 10.1093/genetics/48.10.1303
  • 111. Felsenstein, J., The Evolutionary Advantage of Recombination. Genetics, 1974. 78(2): p. 737-756. DOI: 10.1093/genetics/78.2.737
  • 112. Hill, W.G. and A. Robertson, The effect of linkage on limits to artificial selection. Genetical Research, 2009. 8(3): p. 269-294. DOI: https://doi.org/10.1017/S0016672300010156
  • 113. Maynard Smith, J., The evolution of sex. 1978, Cambridge Eng. ; New York: Cambridge University Press. x, 222 p. ISBN: 9780521293020
  • 114. Bell, G., The masterpiece of nature : the evolution and genetics of sexuality. 1982, Berkeley: University of California Press. 635 p. https://doi.org/10.4324/9780429322884
  • 115. Trivers, R., The Evolution of SexThe Masterpiece of Nature: The Evolution and Genetics of Sexuality.Graham Bell. The Quarterly Review of Biology, 1983. 58(1): p. 62-67. DOI:10.1086/413059
  • 116. Paun, O., et al., Patterns, sources and ecological implications of clonal diversity in apomictic Ranunculus carpaticola (Ranunculus auricomus complex, Ranunculaceae). Molecular Ecology, 2006. 15(4): p. 897-910. DOI: 10.1111/j.1365-294X.2006.02800.x
  • 117. Koch, M.A., Multiple Hybrid Formation in Natural Populations: Concerted Evolution of the Internal Transcribed Spacer of Nuclear Ribosomal DNA (ITS) in North American Arabis divaricarpa (Brassicaceae). Molecular Biology and Evolution, 2003. 20(3): p. 338-350. DOI: 10.1093/molbev/msg046
  • 118. Otto, S.P. and J. Whitton, Polyploid Incidence and Evolution. Annual Review of Genetics, 2000. 34(1): p. 401-437. DOI: 10.1146/annurev.genet.34.1.401
  • 119. Hojsgaard, D. and E. HÃrandl, A little bit of sex matters for genome evolution in asexual plants. Frontiers in Plant Science, 2015. 6. DOI: 10.3389/fpls.2015.00082
Toplam 118 adet kaynakça vardır.

Ayrıntılar

Birincil Dil İngilizce
Konular Bitki Biyoteknolojisi
Bölüm Derleme
Yazarlar

Gözde Yüzbaşıoğlu 0009-0001-5770-7360

Gönderilme Tarihi 24 Nisan 2025
Kabul Tarihi 18 Ağustos 2025
Erken Görünüm Tarihi 15 Aralık 2025
Yayımlanma Tarihi 23 Aralık 2025
Yayımlandığı Sayı Yıl 2025 Cilt: 8 Sayı: 3

Kaynak Göster

EndNote Yüzbaşıoğlu G (01 Aralık 2025) Apomixis and Its Agricultural Potential. International Journal of Life Sciences and Biotechnology 8 3 232–242.


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