Search
2023 Volume 3
Article Contents
REVIEW   Open Access    

Intercellular signaling across plasmodesmata in vegetable species

More Information
  • The formation of edible organs and stress adaption are two major focuses of the studies on vegetable species. The regulation of these two processes often involves cell-to-cell signaling. In most plants, including vegetable species, intercellular signaling can be delivered by mobile regulators that traffic through a channel called plasmodesmata connecting almost all cells. A large number of transcription factors and RNAs have been discovered to move across plasmodesmata (called the symplastic way) to travel a short-range or a long-distance. This symplastic transport of signaling molecules has emerged to be an important regulation of a wide range of developmental and physiological processes. Callose deposition to plasmodesmata is a key step controlling the plasmodesmata permeability in many cell types. Here we summarize the recent progress in our understanding of plasmodesmata-mediated signaling in plants.
  • 加载中
  • [1]

    Wilson EB. 1923. The physical basis of life. Science 57:277−86

    doi: 10.1126/science.57.1471.277

    CrossRef   Google Scholar

    [2]

    Sevilem I, Yadav SR, Helariutta Y. 2015. Plasmodesmata: channels for intercellular signaling during plant growth and development. Methods in Molecular Biology 1217:3−24

    doi: 10.1007/978-1-4939-1523-1_1

    CrossRef   Google Scholar

    [3]

    Vatén A, Dettmer J, Wu S, Stierhof YD, Miyashima S, et al. 2011. Callose biosynthesis regulates symplastic trafficking during root development. Developmental Cell 21:1144−55

    doi: 10.1016/j.devcel.2011.10.006

    CrossRef   Google Scholar

    [4]

    Terauchi M, Nagasato C, Motomura T. 2015. Plasmodesmata of brown algae. Journal of Plant Research 128:7−15

    doi: 10.1007/s10265-014-0677-4

    CrossRef   Google Scholar

    [5]

    Zambryski P, Crawford K. 2000. Plasmodesmata: gatekeepers for cell-to-cell transport of developmental signals in plants. Annual Review of Cell and Developmental Biology 16:393−421

    doi: 10.1146/annurev.cellbio.16.1.393

    CrossRef   Google Scholar

    [6]

    Lucas WJ, Lee JY. 2004. Plasmodesmata as a supracellular control network in plants. Nature Reviews Molecular Cell Biology 5:712−26

    doi: 10.1038/nrm1470

    CrossRef   Google Scholar

    [7]

    Brunkard JO, Runkel AM, Zambryski PC. 2015. The cytosol must flow: intercellular transport through plasmodesmata. Current Opinion in Cell Biology 35:13−20

    doi: 10.1016/j.ceb.2015.03.003

    CrossRef   Google Scholar

    [8]

    Burch-Smith TM, Zambryski PC. 2012. Plasmodesmata paradigm shift: regulation from without versus within. Annual Review of Plant Biology 63:239−60

    doi: 10.1146/annurev-arplant-042811-105453

    CrossRef   Google Scholar

    [9]

    Brunkard JO, Runkel AM, Zambryski PC. 2013. Plasmodesmata dynamics are coordinated by intracellular signaling pathways. Current Opinion in Plant Biology 16:614−20

    doi: 10.1016/j.pbi.2013.07.007

    CrossRef   Google Scholar

    [10]

    Zhang Y, He P, Ma X, Yang Z, Pang C, et al. 2019. Auxin-mediated statolith production for root gravitropism. New Phytologist 224:761−74

    doi: 10.1111/nph.15932

    CrossRef   Google Scholar

    [11]

    Sager R, Lee JY. 2014. Plasmodesmata in integrated cell signalling: insights from development and environmental signals and stresses. Journal of Experimental Botany 65:6337−58

    doi: 10.1093/jxb/eru365

    CrossRef   Google Scholar

    [12]

    Furuta K, Lichtenberger R, Helariutta Y. 2012. The role of mobile small RNA species during root growth and development. Current Opinion in Cell Biology 24:211−16

    doi: 10.1016/j.ceb.2011.12.005

    CrossRef   Google Scholar

    [13]

    Benkovics AH, Timmermans MCP. 2014. Developmental patterning by gradients of mobile small RNAs. Current Opinion in Genetics & Development 27:83−91

    doi: 10.1016/j.gde.2014.04.004

    CrossRef   Google Scholar

    [14]

    Gallagher KL, Sozzani R, Lee CM. 2014. Intercellular protein movement: deciphering the language of development. Annual Review of Cell and Developmental Biology 30:207−33

    doi: 10.1146/annurev-cellbio-100913-012915

    CrossRef   Google Scholar

    [15]

    Niehl A, Heinlein M. 2011. Cellular pathways for viral transport through plasmodesmata. Protoplasma 248:75−99

    doi: 10.1007/s00709-010-0246-1

    CrossRef   Google Scholar

    [16]

    Wu S, Gallagher KL. 2012. Transcription factors on the move. Current Opinion in Plant Biology 15:645−51

    doi: 10.1016/j.pbi.2012.09.010

    CrossRef   Google Scholar

    [17]

    Gömann J, Herrfurth C, Zienkiewicz A, Ischebeck T, Haslam TM, et al. 2021. Sphingolipid long-chain base hydroxylation influences plant growth and callose deposition in Physcomitrium patens. New Phytologist 231:297−314

    doi: 10.1111/nph.17345

    CrossRef   Google Scholar

    [18]

    Xu M, Cho E, Burch-Smith TM, Zambryski PC. 2012. Plasmodesmata formation and cell-to-cell transport are reduced in decreased size exclusion limit 1 during embryogenesis in Arabidopsis. Proceedings of the National Academy of Sciences of the United States of America 109:5098−103

    doi: 10.1073/pnas.1202919109

    CrossRef   Google Scholar

    [19]

    Ehlers K, Kollmann R. 2001. Primary and secondary plasmodesmata: structure, origin, and functioning. Protoplasma 216:1−30

    doi: 10.1007/BF02680127

    CrossRef   Google Scholar

    [20]

    Yadav SR, Yan D, Sevilem I, Helariutta Y. 2014. Plasmodesmata-mediated intercellular signaling during plant growth and development. Frontiers in Plant Science 5:44

    doi: 10.3389/fpls.2014.00044

    CrossRef   Google Scholar

    [21]

    Jackson D. 2015. Plasmodesmata spread their influence. F1000Prime Reports 7:25

    doi: 10.12703/p7-25

    CrossRef   Google Scholar

    [22]

    Wu S, Gallagher KL. 2011. Mobile protein signals in plant development. Current Opinion in Plant Biology 14:563−70

    doi: 10.1016/j.pbi.2011.06.006

    CrossRef   Google Scholar

    [23]

    Kim JY. 2018. Symplasmic intercellular communication through plasmodesmata. Plants 7:23

    doi: 10.3390/plants7010023

    CrossRef   Google Scholar

    [24]

    Kragler F. 2015. Analysis of the conductivity of plasmodesmata by microinjection. Methods in Molecular Biology 1217:173−84

    doi: 10.1007/978-1-4939-1523-1_12

    CrossRef   Google Scholar

    [25]

    Christensen NM, Faulkner C, Oparka K. 2009. Evidence for unidirectional flow through plasmodesmata. Plant Physiology 150:96−104

    doi: 10.1104/pp.109.137083

    CrossRef   Google Scholar

    [26]

    Lucas WJ, Bouché-Pillon S, Jackson DP, Nguyen L, Baker L, et al. 1995. Selective trafficking of KNOTTED1 homeodomain protein and its mRNA through plasmodesmata. Science 270:1980−83

    doi: 10.1126/science.270.5244.1980

    CrossRef   Google Scholar

    [27]

    Xu XM, Wang J, Xuan Z, Goldshmidt A, Borrill PGM, et al. 2011. Chaperonins facilitate KNOTTED1 cell-to-cell trafficking and stem cell function. Science 333:1141−44

    doi: 10.1126/science.1205727

    CrossRef   Google Scholar

    [28]

    Yadav RK, Perales M, Gruel J, Girke T, Jönsson H, et al. 2011. WUSCHEL protein movement mediates stem cell homeostasis in the Arabidopsis shoot apex. Genes & Development 25:2025−30

    doi: 10.1101/gad.17258511

    CrossRef   Google Scholar

    [29]

    Daum G, Medzihradszky A, Suzaki T, Lohmann JU. 2014. A mechanistic framework for noncell autonomous stem cell induction in Arabidopsis. Proceedings of the National Academy of Sciences of the United States of America 111:14619−24

    doi: 10.1073/pnas.1406446111

    CrossRef   Google Scholar

    [30]

    Pi L, Aichinger E, van der Graaff E, Llavata-Peris CI, Weijers D, et al. 2015. Organizer-derived WOX5 signal maintains root Columella stem cells through chromatin-mediated repression of CDF4 expression. Developmental Cell 33:576−88

    doi: 10.1016/j.devcel.2015.04.024

    CrossRef   Google Scholar

    [31]

    Kim JY, Yuan Z, Jackson D. 2003. Developmental regulation and significance of KNOX protein trafficking in Arabidopsis. Development 130:4351−62

    doi: 10.1242/dev.00618

    CrossRef   Google Scholar

    [32]

    Kim JY, Rim Y, Wang J, Jackson D. 2005. A novel cell-to-cell trafficking assay indicates that the KNOX homeodomain is necessary and sufficient for intercellular protein and mRNA trafficking. Genes & Development 19:788−93

    doi: 10.1101/gad.332805

    CrossRef   Google Scholar

    [33]

    Rim Y, Jung JH, Chu H, Cho WK, Kim SW, et al. 2009. A non-cell-autonomous mechanism for the control of plant architecture and epidermal differentiation involves intercellular trafficking of BREVIPEDICELLUS protein. Functional Plant Biology 36:280−89

    doi: 10.1071/FP08243

    CrossRef   Google Scholar

    [34]

    Schlereth A, Möller B, Liu W, Kientz M, Flipse J, et al. 2010. MONOPTEROS controls embryonic root initiation by regulating a mobile transcription factor. Nature 464:913−16

    doi: 10.1038/nature08836

    CrossRef   Google Scholar

    [35]

    Lu KJ, De Rybel B, van Mourik H, Weijers D. 2018. Regulation of intercellular TARGET OF MONOPTEROS 7 protein transport in the Arabidopsis root. Development 145:dev152892

    doi: 10.1242/dev.152892

    CrossRef   Google Scholar

    [36]

    Rim Y, Huang L, Chu H, Han X, Cho WK, et al. 2011. Analysis of Arabidopsis transcription factor families revealed extensive capacity for cell-to-cell movement as well as discrete trafficking patterns. Molecules and Cells 32:519−26

    doi: 10.1007/s10059-011-0135-2

    CrossRef   Google Scholar

    [37]

    Zhou J, Wang X, Lee JY, Lee JY. 2013. Cell-to-cell movement of two interacting AT-hook factors in Arabidopsis root vascular tissue patterning. The Plant cell 25:187−201

    doi: 10.1105/tpc.112.102210

    CrossRef   Google Scholar

    [38]

    Long Y, Goedhart J, Schneijderberg M, Terpstra I, Shimotohno A, et al. 2015. SCARECROW-LIKE23 and SCARECROW jointly specify endodermal cell fate but distinctly control SHORT-ROOT movement. The Plant Journal 84:773−84

    doi: 10.1111/tpj.13038

    CrossRef   Google Scholar

    [39]

    Corbesier L, Vincent C, Jang S, Fornara F, Fan Q, et al. 2007. FT protein movement contributes to long-distance signaling in floral induction of Arabidopsis. Science 316:1030−33

    doi: 10.1126/science.1141752

    CrossRef   Google Scholar

    [40]

    Jaeger KE, Wigge PA. 2007. FT protein acts as a long-range signal in Arabidopsis. Current Biology 17:1050−54

    doi: 10.1016/j.cub.2007.05.008

    CrossRef   Google Scholar

    [41]

    Chen X, Yao Q, Gao X, Jiang C, Harberd NP, et al. 2016. Shoot-to-root mobile transcription factor HY5 coordinates plant carbon and nitrogen acquisition. Current Biology 26:640−46

    doi: 10.1016/j.cub.2015.12.066

    CrossRef   Google Scholar

    [42]

    Harries P, Ding B. 2011. Cellular factors in plant virus movement: at the leading edge of macromolecular trafficking in plants. Virology 411:237−43

    doi: 10.1016/j.virol.2010.12.021

    CrossRef   Google Scholar

    [43]

    Wu S, Gallagher KL. 2013. Intact microtubules are required for the intercellular movement of the SHORT-ROOT transcription factor. The Plant Journal 74:148−59

    doi: 10.1111/tpj.12112

    CrossRef   Google Scholar

    [44]

    Koizumi K, Wu S, MacRae-Crerar A, Gallagher KL. 2011. An essential protein that interacts with endosomes and promotes movement of the SHORT-ROOT transcription factor. Current Biology 21:1559−64

    doi: 10.1016/j.cub.2011.08.013

    CrossRef   Google Scholar

    [45]

    Carrington JC, Ambros V. 2003. Role of microRNAs in plant and animal development. Science 301:336−38

    doi: 10.1126/science.1085242

    CrossRef   Google Scholar

    [46]

    Schramke V, Luciano P, Brevet V, Guillot S, Corda Y, et al. 2004. RPA regulates telomerase action by providing Est1p access to chromosome ends. Nature Genetics 36:46−54

    doi: 10.1038/ng1284

    CrossRef   Google Scholar

    [47]

    Khraiwesh B, Arif MA, Seumel GI, Ossowski S, Weigel D, et al. 2010. Transcriptional control of gene expression by microRNAs. Cell 140:111−22

    doi: 10.1016/j.cell.2009.12.023

    CrossRef   Google Scholar

    [48]

    Loreti E, Perata P. 2022. Mobile plant microRNAs allow communication within and between organisms. New Phytologist 235:2176−82

    doi: 10.1111/nph.18360

    CrossRef   Google Scholar

    [49]

    Chen X, Rechavi O. 2022. Plant and animal small RNA communications between cells and organisms. Nature Reviews Molecular Cell Biology 23:185−203

    doi: 10.1038/s41580-021-00425-y

    CrossRef   Google Scholar

    [50]

    Brioudes F, Jay F, Sarazin A, Grentzinger T, Devers EA, et al. 2021. HASTY, the Arabidopsis EXPORTIN5 ortholog, regulates cell-to-cell and vascular microRNA movement. The EMBO Journal 40:e107455

    doi: 10.15252/embj.2020107455

    CrossRef   Google Scholar

    [51]

    Lin SI, Chiang SF, Lin WY, Chen JW, Tseng CY, et al. 2008. Regulatory network of microRNA399 and PHO2 by systemic signaling. Plant Physiology 147:732−46

    doi: 10.1104/pp.108.116269

    CrossRef   Google Scholar

    [52]

    Huen AK, Rodriguez-Medina C, Ho AYY, Atkins CA, Smith PMC. 2017. Long-distance movement of phosphate starvation-responsive microRNAs in Arabidopsis. Plant Biology 19:643−49

    doi: 10.1111/plb.12568

    CrossRef   Google Scholar

    [53]

    Tsikou D, Yan Z, Holt DB, Abel NB, Reid DE, et al. 2018. Systemic control of legume susceptibility to rhizobial infection by a mobile microRNA. Science 362:233−36

    doi: 10.1126/science.aat6907

    CrossRef   Google Scholar

    [54]

    Buhtz A, Pieritz J, Springer F, Kehr J. 2010. Phloem small RNAs, nutrient stress responses, and systemic mobility. BMC Plant Biology 10:64

    doi: 10.1186/1471-2229-10-64

    CrossRef   Google Scholar

    [55]

    Martin A, Adam H, Díaz-Mendoza M, Żurczak M, González-Schain ND, et al. 2009. Graft-transmissible induction of potato tuberization by the microRNA miR172. Development 136:2873−81

    doi: 10.1242/dev.031658

    CrossRef   Google Scholar

    [56]

    Wu G, Park MY, Conway SR, Wang JW, Weigel D, et al. 2009. The sequential action of miR156 and miR172 regulates developmental timing in Arabidopsis. Cell 138:750−59

    doi: 10.1016/j.cell.2009.06.031

    CrossRef   Google Scholar

    [57]

    Eviatar-Ribak T, Shalit-Kaneh A, Chappell-Maor L, Amsellem Z, Eshed Y, et al. 2013. A cytokinin-activating enzyme promotes tuber formation in tomato. Current Biology 23:1057−64

    doi: 10.1016/j.cub.2013.04.061

    CrossRef   Google Scholar

    [58]

    Carlsbecker A, Lee JY, Roberts CJ, Dettmer J, Lehesranta S, et al. 2010. Cell signalling by microRNA165/6 directs gene dose-dependent root cell fate. Nature 465:316−21

    doi: 10.1038/nature08977

    CrossRef   Google Scholar

    [59]

    Knauer S, Holt AL, Rubio-Somoza I, Tucker EJ, Hinze A, et al. 2013. A protodermal miR394 signal defines a region of stem cell competence in the Arabidopsis shoot meristem. Developmental Cell 24:125−32

    doi: 10.1016/j.devcel.2012.12.009

    CrossRef   Google Scholar

    [60]

    Lu KJ, Huang NC, Liu YS, Lu CA, Yu TS. 2012. Long-distance movement of Arabidopsis FLOWERING LOCUS T RNA participates in systemic floral regulation. RNA Biology 9:653−62

    doi: 10.4161/rna.19965

    CrossRef   Google Scholar

    [61]

    Notaguchi M, Wolf S, Lucas WJ. 2012. Phloem-mobile Aux/IAA transcripts target to the root tip and modify root architecture. Journal of Integrative Plant Biology 54:760−72

    doi: 10.1111/j.1744-7909.2012.01155.x

    CrossRef   Google Scholar

    [62]

    Xoconostle-Cázares B, Xiang Y, Ruiz-Medrano R, Wang HL, Monzer J, et al. 1999. Plant paralog to viral movement protein that potentiates transport of mRNA into the phloem. Science 283:94−98

    doi: 10.1126/science.283.5398.94

    CrossRef   Google Scholar

    [63]

    Doering-Saad C, Newbury HJ, Couldridge CE, Bale JS, Pritchard J. 2006. A phloem-enriched cDNA library from Ricinus: insights into phloem function. Journal of Experimental Botany 57:3183−93

    doi: 10.1093/jxb/erl082

    CrossRef   Google Scholar

    [64]

    Kanehira A, Yamada K, Iwaya T, Tsuwamoto R, Kasai A, et al. 2010. Apple phloem cells contain some mRNAs transported over long distances. Tree Genetics & Genomes 6:635−42

    doi: 10.1007/s11295-010-0279-9

    CrossRef   Google Scholar

    [65]

    Kim M, Canio W, Kessler S, Sinha N. 2001. Developmental changes due to long-distance movement of a homeobox fusion transcript in tomato. Science 293:287−89

    doi: 10.1126/science.1059805

    CrossRef   Google Scholar

    [66]

    Luo KR, Huang NC, Yu TS. 2018. Selective targeting of mobile mRNAs to plasmodesmata for cell-to-cell movement. Plant Physiology 177:604−14

    doi: 10.1104/pp.18.00107

    CrossRef   Google Scholar

    [67]

    Thieme CJ, Rojas-Triana M, Stecyk E, Schudoma C, Zhang W, et al. 2015. Endogenous Arabidopsis messenger RNAs transported to distant tissues. Nature Plants 1:15025

    doi: 10.1038/nplants.2015.25

    CrossRef   Google Scholar

    [68]

    Yang Y, Mao L, Jittayasothorn Y, Kang Y, Jiao C, et al. 2015. Messenger RNA exchange between scions and rootstocks in grafted grapevines. BMC Plant Biology 15:251

    doi: 10.1186/s12870-015-0626-y

    CrossRef   Google Scholar

    [69]

    Notaguchi M, Higashiyama T, Suzuki T. 2015. Identification of mRNAs that move over long distances using an RNA-seq analysis of Arabidopsis/Nicotiana benthamiana heterografts. Plant and Cell Physiology 56:311−21

    doi: 10.1093/pcp/pcu210

    CrossRef   Google Scholar

    [70]

    Zhang Z, Zheng Y, Ham BK, Chen J, Yoshida A, et al. 2016. Vascular-mediated signalling involved in early phosphate stress response in plants. Nature Plants 2:16033

    doi: 10.1038/nplants.2016.33

    CrossRef   Google Scholar

    [71]

    Xia C, Zheng Y, Huang J, Zhou X, Li R, et al. 2018. Elucidation of the mechanisms of long-distance mRNA movement in a Nicotiana benthamiana/tomato heterograft system. Plant Physiology 177:745−58

    doi: 10.1104/pp.17.01836

    CrossRef   Google Scholar

    [72]

    Liu L, Chen X. 2018. Intercellular and systemic trafficking of RNAs in plants. Nature Plants 4:869−78

    doi: 10.1038/s41477-018-0288-5

    CrossRef   Google Scholar

    [73]

    Petricka JJ, Winter CM, Benfey PN. 2012. Control of Arabidopsis root development. Annual Review of Plant Biology 63:563−90

    doi: 10.1146/annurev-arplant-042811-105501

    CrossRef   Google Scholar

    [74]

    Helariutta Y, Fukaki H, Wysocka-Diller J, Nakajima K, Jung J, et al. 2000. The SHORT-ROOT gene controls radial patterning of the Arabidopsis root through radial signaling. Cell 101:555−67

    doi: 10.1016/S0092-8674(00)80865-X

    CrossRef   Google Scholar

    [75]

    Cui H, Levesque MP, Vernoux T, Jung JW, Paquette AJ, et al. 2007. An evolutionarily conserved mechanism delimiting SHR movement defines a single layer of endodermis in plants. Science 316:421−25

    doi: 10.1126/science.1139531

    CrossRef   Google Scholar

    [76]

    Levesque MP, Vernoux T, Busch W, Cui H, Wang JY, et al. 2006. Whole-genome analysis of the SHORT-ROOT developmental pathway in Arabidopsis. PLoS Biology 4:e143

    doi: 10.1371/journal.pbio.0040143

    CrossRef   Google Scholar

    [77]

    Sozzani R, Cui H, Moreno-Risueno MA, Busch W, Van Norman JM, et al. 2010. Spatiotemporal regulation of cell-cycle genes by SHORTROOT links patterning and growth. Nature 466:128−32

    doi: 10.1038/nature09143

    CrossRef   Google Scholar

    [78]

    Nakajima K, Sena G, Nawy T, Benfey PN. 2001. Intercellular movement of the putative transcription factor SHR in root patterning. Nature 413:307−11

    doi: 10.1038/35095061

    CrossRef   Google Scholar

    [79]

    Gallagher KL, Paquette AJ, Nakajima K, Benfey PN. 2004. Mechanisms regulating SHORT-ROOT intercellular movement. Current Biology 14:1847−51

    doi: 10.1016/j.cub.2004.09.081

    CrossRef   Google Scholar

    [80]

    Wu S, Lee CM, Hayashi T, Price S, Divol F, et al. 2014. A plausible mechanism, based upon SHORT-ROOT movement, for regulating the number of cortex cell layers in roots. Proceedings of the National Academy of Sciences of the United States of America 111:16184−89

    doi: 10.1073/pnas.1407371111

    CrossRef   Google Scholar

    [81]

    Geldner N. 2013. The endodermis. Annual Review of Plant Biology 64:531−58

    doi: 10.1146/annurev-arplant-050312-120050

    CrossRef   Google Scholar

    [82]

    Yu Q, Li P, Liang N, Wang H, Xu M, et al. 2017. Cell-fate specification in Arabidopsis roots requires coordinative action of lineage instruction and positional reprogramming. Plant Physiology 175:816−27

    doi: 10.1104/pp.17.00814

    CrossRef   Google Scholar

    [83]

    Li P, Yu Q, Gu X, Xu C, Qi S, et al. 2018. Construction of a functional casparian strip in non-endodermal lineages is orchestrated by two parallel signaling systems in Arabidopsis thaliana. Current Biology 28:2777−2286.E2

    doi: 10.1016/j.cub.2018.07.028

    CrossRef   Google Scholar

    [84]

    Di Ruocco G, Bertolotti G, Pacifici E, Polverari L, Tsiantis M, et al. 2018. Differential spatial distribution of miR165/6 determines variability in plant root anatomy. Development 145:dev153858

    doi: 10.1242/dev.153858

    CrossRef   Google Scholar

    [85]

    Hashimoto K, Miyashima S, Sato-Nara K, Yamada T, Nakajima K. 2018. Functionally diversified members of the MIR165/6 gene family regulate ovule morphogenesis in Arabidopsis thaliana. Plant and Cell Physiology 59:1017−26

    doi: 10.1093/pcp/pcy042

    CrossRef   Google Scholar

    [86]

    Slewinski TL, Baker RF, Stubert A, Braun DM. 2012. Tie-dyed2 encodes a callose synthase that functions in vein development and affects symplastic trafficking within the phloem of maize leaves. Plant Physiology 160:1540−50

    doi: 10.1104/pp.112.202473

    CrossRef   Google Scholar

    [87]

    Hoffman G, McEvoy PB. 1985. Mechanical limitations on feeding by meadow spittlebugs Philaenus spumarius (Homoptera: Cercopidae) on wild and cultivated host plants. Ecological Entomology 10:415−26

    doi: 10.1111/j.1365-2311.1985.tb00739.x

    CrossRef   Google Scholar

    [88]

    Yang C, Gao Y, Gao S, Yu G, Xiong C, et al. 2015. Transcriptome profile analysis of cell proliferation molecular processes during multicellular trichome formation induced by tomato Wov gene in tobacco. BMC Genetics 16:868

    doi: 10.1186/s12864-015-2099-7

    CrossRef   Google Scholar

    [89]

    Balkunde R, Bouyer D, Hulskamp M. 2011. Nuclear trapping by GL3 controls intercellular transport and redistribution of TTG1 protein in Arabidopsis. Development 138:5039−48

    doi: 10.1242/dev.072454

    CrossRef   Google Scholar

    [90]

    Wester K, Digiuni S, Geier F, Timmer J, Fleck C, et al. 2009. Functional diversity of R3 single-repeat genes in trichome development. Development 136:1487−96

    doi: 10.1242/dev.021733

    CrossRef   Google Scholar

    [91]

    Kang YH, Song SK, Schiefelbein J, Lee MM. 2013. Nuclear trapping controls the position-dependent localization of CAPRICE in the root epidermis of Arabidopsis. Plant Physiology 163:193−204

    doi: 10.1104/pp.113.221028

    CrossRef   Google Scholar

    [92]

    Cnops G, Wang X, Linstead P, Montagu MV, Van Lijsebettens M, et al. 2000. TORNADO1 and TORNADO2 are required for the specification of radial and circumferential pattern in the Arabidopsis root. Development 127:3385−94

    doi: 10.1242/dev.127.15.3385

    CrossRef   Google Scholar

    [93]

    Kwak SH, Song SK, Lee MM, Schiefelbein J. 2015. TORNADO1 regulates root epidermal patterning through the WEREWOLF pathway in Arabidopsis thaliana. Plant Signaling & Behavior 10:e1103407

    doi: 10.1080/15592324.2015.1103407

    CrossRef   Google Scholar

    [94]

    Fan Y, Lin S, Li T, Shi F, Shan G, et al. 2022. The plasmodesmata-Located β-1, 3-Glucanase enzyme PdBG4 regulates trichomes growth in Arabidopsis thaliana. Cells 11:2856

    doi: 10.3390/cells11182856

    CrossRef   Google Scholar

    [95]

    Pillitteri LJ, Torii KU. 2012. Mechanisms of stomatal development. Annual Review of Plant Biology 63:591−614

    doi: 10.1146/annurev-arplant-042811-105451

    CrossRef   Google Scholar

    [96]

    Guseman JM, Lee JS, Bogenschutz NL, Peterson KM, Virata RE, et al. 2010. Dysregulation of cell-to-cell connectivity and stomatal patterning by loss-of-function mutation in Arabidopsis CHORUS (GLUCAN SYNTHASE-LIKE 8). Development 137:1731−41

    doi: 10.1242/dev.049197

    CrossRef   Google Scholar

    [97]

    Raissig MT, Matos JL, Anleu Gil MX, Kornfeld A, Bettadapur A, et al. 2017. Mobile MUTE specifies subsidiary cells to build physiologically improved grass stomata. Science 355:1215−18

    doi: 10.1126/science.aal3254

    CrossRef   Google Scholar

    [98]

    Bilska A, Sowiński P. 2010. Closure of plasmodesmata in maize (Zea mays) at low temperature: a new mechanism for inhibition of photosynthesis. Annals of Botany 106:675−86

    doi: 10.1093/aob/mcq169

    CrossRef   Google Scholar

    [99]

    Wu J, Sun W, Sun C, Xu C, Li S, et al. 2022. Cold stress induces malformed tomato fruits by breaking the feedback loops of stem cell regulation in floral meristem. New Phytologist 237:2268−83

    doi: 10.1111/nph.18699

    CrossRef   Google Scholar

    [100]

    Xie B, Wang X, Zhu M, Zhang Z, Hong Z. 2011. CalS7 encodes a callose synthase responsible for callose deposition in the phloem. The Plant Journal 65:1−14

    doi: 10.1111/j.1365-313X.2010.04399.x

    CrossRef   Google Scholar

    [101]

    Cui W, Lee JY. 2016. Arabidopsis callose synthases CalS1/8 regulate plasmodesmal permeability during stress. Nature Plants 2:16034

    doi: 10.1038/nplants.2016.34

    CrossRef   Google Scholar

    [102]

    Rinne PLH, van den Boogaard R, Mensink MGJ, Kopperud C, Kormelink R, et al. 2005. Tobacco plants respond to the constitutive expression of the tospovirus movement protein NSM with a heat-reversible sealing of plasmodesmata that impairs development. The Plant Journal 43:688−707

    doi: 10.1111/j.1365-313X.2005.02489.x

    CrossRef   Google Scholar

    [103]

    Liu J, Liu Y, Wang S, Cui Y, Yan D. 2022. Heat stress reduces root meristem size via induction of plasmodesmal callose accumulation inhibiting phloem unloading in Arabidopsis. International Journal of Molecular Sciences 23:2063

    doi: 10.3390/ijms23042063

    CrossRef   Google Scholar

    [104]

    Sivaguru M, Fujiwara T, Šamaj J, Baluška F, Yang Z, et al. 2000. Aluminum-induced 1→3-beta-D-glucan inhibits cell-to-cell trafficking of molecules through plasmodesmata. A new mechanism of aluminum toxicity in plants. Plant Physiology 124:991−1006

    doi: 10.1104/pp.124.3.991

    CrossRef   Google Scholar

    [105]

    Ueki S, Citovsky V. 2002. The systemic movement of a tobamovirus is inhibited by a cadmium-ion-induced glycine-rich protein. Nature Cell Biology 4:478−86

    doi: 10.1038/ncb806

    CrossRef   Google Scholar

    [106]

    Ueki S, Citovsky V. 2005. Identification of an interactor of cadmium ion-induced glycine-rich protein involved in regulation of callose levels in plant vasculature. Proceedings of the National Academy of Sciences of the United States of America 102:12089−12094

    doi: 10.1073/pnas.0505927102

    CrossRef   Google Scholar

    [107]

    O'Lexy R, Kasai K, Clark N, Fujiwara T, Sozzani R, et al. 2018. Exposure to heavy metal stress triggers changes in plasmodesmatal permeability via deposition and breakdown of callose. Journal of Experimental Botany 69:3715−28

    doi: 10.1093/jxb/ery171

    CrossRef   Google Scholar

    [108]

    Faulkner C, Petutschnig E, Benitez-Alfonso Y, Beck M, Robatzek S, et al. 2013. LYM2-dependent chitin perception limits molecular flux via plasmodesmata. Proceedings of the National Academy of Sciences of the United States of America 110:9166−70

    doi: 10.1073/pnas.1203458110

    CrossRef   Google Scholar

    [109]

    Wang X, Sager R, Cui W, Zhang C, Lu H, et al. 2013. Salicylic acid regulates Plasmodesmata closure during innate immune responses in Arabidopsis. The Plant Cell 25:2315−29

    doi: 10.1105/tpc.113.110676

    CrossRef   Google Scholar

    [110]

    Ding B, Haudenshield JS, Hull RJ, Wolf S, Beachy RN, et al. 1992. Secondary plasmodesmata are specific sites of localization of the tobacco mosaic virus movement protein in transgenic tobacco plants. The Plant Cell 4:915−28

    doi: 10.1105/tpc.4.8.915

    CrossRef   Google Scholar

    [111]

    Wang H, Cao S, Li T, Zhang L, Yao J, et al. 2021. Classification and expression analysis of cucumber (Cucumis sativus L.) callose synthase (CalS) family genes and localization of CsCalS4, a protein involved in pollen development. Biotechnology and Biotechnological Equipment 35:1992−2004

    doi: 10.1080/13102818.2022.2038670

    CrossRef   Google Scholar

    [112]

    Parr R, Gomez-Jimenez MC. 2020. Spatio–temporal immunolocalization of extensin protein and hemicellulose polysaccharides during olive fruit abscission. Planta 252:32

    doi: 10.1007/s00425-020-03403-4

    CrossRef   Google Scholar

    [113]

    Zhang J, Liu N, Yan A, Sun T, Yao G, et al. 2022. Callose deposited at soybean sieve element inhibits long-distance transport of Soybean mosaic virus. AMB Express 12:66

    doi: 10.1186/s13568-022-01402-0

    CrossRef   Google Scholar

    [114]

    Rinne PLH, Kaikuranta PM, van der Schoot C. 2001. The shoot apical meristem restores its symplasmic organization during chilling-induced release from dormancy. The Plant Journal 26:249−64

    doi: 10.1046/j.1365-313X.2001.01022.x

    CrossRef   Google Scholar

    [115]

    Iswanto ABB, Vu MH, Pike S, Lee J, Kang H, et al. 2022. Pathogen effectors: what do they do at plasmodesmata? Molecular Plant Pathology 23:795−804

    doi: 10.1111/mpp.13142

    CrossRef   Google Scholar

    [116]

    Mbiza NIT, Hu Z, Zhang H, Zhang Y, Luo X, et al. 2022. GhCalS5 is involved in cotton response to aphid attack through mediating callose formation. Frontiers in Plant Science 13:892630

    doi: 10.3389/fpls.2022.892630

    CrossRef   Google Scholar

    [117]

    Niu Q, Zhang P, Su S, Jiang B, Liu X, et al. 2022. Characterization and expression analyses of callose synthase enzyme (Cals) family genes in maize (Zea mays L.). Biochemical Genetics 60:351−69

    doi: 10.1007/s10528-021-10103-5

    CrossRef   Google Scholar

    [118]

    Rinne PLH, Welling A, Vahala J, Ripel L, Ruonala R, et al. 2011. Chilling of dormant buds hyperinduces FLOWERING LOCUS T and recruits GA-inducible 1, 3-β-glucanases to reopen signal conduits and release dormancy in Populus. The Plant Cell 23:130−46

    doi: 10.1105/tpc.110.081307

    CrossRef   Google Scholar

    [119]

    Tylewicz S, Petterle A, Marttila S, Miskolczi P, Azeez A, et al. 2018. Photoperiodic control of seasonal growth is mediated by ABA acting on cell-cell communication. Science 360:212−15

    doi: 10.1126/science.aan8576

    CrossRef   Google Scholar

    [120]

    Leubner-Metzger G, Meins F Jr. 2000. Sense transformation reveals a novel role for class I β-1, 3-glucanase in tobacco seed germination. The Plant Journal 23:215−21

    doi: 10.1046/j.1365-313x.2000.00773.x

    CrossRef   Google Scholar

    [121]

    Wu S, O'Lexy R, Xu M, Sang Y, Chen X, et al. 2016. Symplastic signaling instructs cell division, cell expansion, and cell polarity in the ground tissue of Arabidopsis thaliana roots. Proceedings of the National Academy of Sciences of the United States of America 113:11621−26

    doi: 10.1073/pnas.1610358113

    CrossRef   Google Scholar

    [122]

    Barratt DHP, Kölling K, Graf A, Pike M, Calder G, et al. 2011. Callose synthase GSL7 is necessary for normal phloem transport and inflorescence growth in Arabidopsis. Plant Physiology 155:328−41

    doi: 10.1104/pp.110.166330

    CrossRef   Google Scholar

    [123]

    De Storme N, De Schrijver J, Van Criekinge W, Wewer V, Dörmann P, et al. 2013. GLUCAN SYNTHASE-LIKE8 and STEROL METHYLTRANSFERASE2 are required for ploidy consistency of the sexual reproduction system in Arabidopsis. The Plant Cell 25:387−403

    doi: 10.1105/tpc.112.106278

    CrossRef   Google Scholar

    [124]

    Sevilem I, Miyashima S, Helariutta Y. 2013. Cell-to-cell communication via plasmodesmata in vascular plants. Cell Adhesion & Migration 7:27−32

    doi: 10.4161/cam.22126

    CrossRef   Google Scholar

    [125]

    Zuo J, Niu QW, Chua NH. 2000. Technical advance: An estrogen receptor-based transactivator XVE mediates highly inducible gene expression in transgenic plants. The Plant Journal 24:265−73

    doi: 10.1046/j.1365-313x.2000.00868.x

    CrossRef   Google Scholar

    [126]

    Benitez-Alfonso Y, Faulkner C, Pendle A, Miyashima S, Helariutta Y, et al. 2013. Symplastic intercellular connectivity regulates lateral root patterning. Developmental Cell 26:136−47

    doi: 10.1016/j.devcel.2013.06.010

    CrossRef   Google Scholar

    [127]

    van den Berg C, Willemsen V, Hage W, Weisbeek P, Scheres B. 1995. Cell fate in the Arabidopsis root meristem determined by directional signalling. Nature 378:62−65

    doi: 10.1038/378062a0

    CrossRef   Google Scholar

    [128]

    Liu Y, Xu M, Liang N, Zheng Y, Yu Q, et al. 2017. Symplastic communication spatially directs local auxin biosynthesis to maintain root stem cell niche in Arabidopsis. Proceedings of the National Academy of Sciences of the United States of America 114:4005−10

    doi: 10.1073/pnas.1616387114

    CrossRef   Google Scholar

    [129]

    Li M, Wang M, Lin Q, Wang M, Niu X, et al. 2022. Symplastic communication in the root cap directs auxin distribution to modulate root development. Journal of Integrative Plant Biology 64:859−70

    doi: 10.1111/jipb.13237

    CrossRef   Google Scholar

    [130]

    Chambaud C, Cookson SJ, Ollat N, Bayer E, Brocard L. 2022. A correlative light electron microscopy approach reveals plasmodesmata ultrastructure at the graft interface. Plant Physiology 188:44−55

    doi: 10.1093/plphys/kiab485

    CrossRef   Google Scholar

    [131]

    Band LR. 2021. Auxin fluxes through plasmodesmata. New Phytologist 231:1686−92

    doi: 10.1111/nph.17517

    CrossRef   Google Scholar

    [132]

    Botha CEJ, Murugan N. 2021. Changes in structure and dimension of plasmodesmata in the phloem loading pathway in Tecoma capensis (Bignoniaceae) - locating the polymer trap. South African Journal of Botany 140:76−86

    doi: 10.1016/j.sajb.2021.03.032

    CrossRef   Google Scholar

    [133]

    Ross-Elliott TJ, Jensen KH, Haaning KS, Wager BM, Knoblauch J, et al. 2017. Phloem unloading in Arabidopsis roots is convective and regulated by the phloem-pole pericycle. eLife 6:e24125

    doi: 10.7554/eLife.24125

    CrossRef   Google Scholar

    [134]

    Holdaway-Clarke TL, Walker NA, Hepler PK, Overall RL. 1999. Physiological elevations in cytoplasmic free calcium by cold or ion injection result in transient closure of higher plant plasmodesmata. Planta 210:329−35

    doi: 10.1007/PL00008141

    CrossRef   Google Scholar

    [135]

    Stonebloom S, Brunkard JO, Cheung AC, Jiang K, Feldman L, et al. 2012. Redox states of plastids and mitochondria differentially regulate intercellular transport via plasmodesmata. Plant Physiology 158:190−99

    doi: 10.1104/pp.111.186130

    CrossRef   Google Scholar

    [136]

    Waigmann E, Chen MH, Bachmaier R, Ghoshroy S, Citovsky V. 2000. Regulation of plasmodesmal transport by phosphorylation of tobacco mosaic virus cell-to-cell movement protein. The EMBO Journal 19:4875−84

    doi: 10.1093/emboj/19.18.4875

    CrossRef   Google Scholar

    [137]

    Huang X, Zhang Q, Wang G, Guo X, Li Z. 2019. Medical image super-resolution based on the generative adversarial network. Proceedings of 2019 Chinese Intelligent Systems Conference. pp. 243–53. https://link.springer.com/chapter/10.1007/978-981-32-9686-2_29

    [138]

    Linh NM, Scarpella E. 2022. Leaf vein patterning is regulated by the aperture of plasmodesmata intercellular channels. PLoS Biology 20:e3001781

    doi: 10.1371/journal.pbio.3001781

    CrossRef   Google Scholar

    [139]

    Kameoka H, Dun EA, Lopez-Obando M, Brewer PB, de Saint Germain A, et al. 2016. Phloem transport of the receptor DWARF14 protein is required for full function of strigolactones. Plant Physiology 172:1844−52

    doi: 10.1104/pp.16.01212

    CrossRef   Google Scholar

    [140]

    Kawade K, Horiguchi G, Usami T, Hirai MY, Tsukaya H. 2013. ANGUSTIFOLIA3 signaling coordinates proliferation between clonally distinct cells in leaves. Current Biology 23:788−92

    doi: 10.1016/j.cub.2013.03.044

    CrossRef   Google Scholar

    [141]

    Mähönen AP, Ten Tusscher K, Siligato R, Smetana O, Díaz-Triviño S, et al. 2014. PLETHORA gradient formation mechanism separates auxin responses. Nature 515:125−29

    doi: 10.1038/nature13663

    CrossRef   Google Scholar

    [142]

    Galinha C, Hofhuis H, Luijten M, Willemsen V, Blilou I, et al. 2007. PLETHORA proteins as dose-dependent master regulators of Arabidopsis root development. Nature 449:1053−57

    doi: 10.1038/nature06206

    CrossRef   Google Scholar

    [143]

    Spiegelman Z, Ham BK, Zhang Z, Toal TW, Brady SM, et al. 2015. A tomato phloem-mobile protein regulates the shoot-to-root ratio by mediating the auxin response in distant organs. The Plant Journal 83:853−63

    doi: 10.1111/tpj.12932

    CrossRef   Google Scholar

    [144]

    Tsukagoshi H, Busch W, Benfey PN. 2010. Transcriptional regulation of ROS controls transition from proliferation to differentiation in the root. Cell 143:606−16

    doi: 10.1016/j.cell.2010.10.020

    CrossRef   Google Scholar

    [145]

    Kühn C, Barker L, Bürkle L, Frommer WB. 1999. Update on sucrose transport in higher plants. Journal of Experimental Botany 50:935−53

    doi: 10.1093/jxb/50.Special_Issue.935

    CrossRef   Google Scholar

    [146]

    Ruiz-Medrano R, Xoconostle-Cázares B, Lucas WJ. 1999. Phloem long-distance transport of CmNACP mRNA: implications for supracellular regulation in plants. Development 126:4405−19

    doi: 10.1242/dev.126.20.4405

    CrossRef   Google Scholar

    [147]

    Banerjee AK, Lin T, Hannapel DJ. 2009. Untranslated regions of a mobile transcript mediate RNA metabolism. Plant Physiology 151:1831−43

    doi: 10.1104/pp.109.144428

    CrossRef   Google Scholar

    [148]

    Mahajan A, Bhogale S, Kang IH, Hannapel DJ, Banerjee AK. 2012. The mRNA of a Knotted1-like transcription factor of potato is phloem mobile. Plant Molecular Biology 79:595−608

    doi: 10.1007/s11103-012-9931-0

    CrossRef   Google Scholar

    [149]

    Zhang H, Yu P, Zhao J, Jiang H, Wang H, et al. 2018. Expression of tomato prosystemin gene in Arabidopsis reveals systemic translocation of its mRNA and confers necrotrophic fungal resistance. New Phytologist 217:799−812

    doi: 10.1111/nph.14858

    CrossRef   Google Scholar

    [150]

    Roney JK, Khatibi PA, Westwood JH. 2007. Cross-species translocation of mRNA from host plants into the parasitic plant dodder. Plant Physiology 143:1037−43

    doi: 10.1104/pp.106.088369

    CrossRef   Google Scholar

    [151]

    David-Schwartz R, Runo S, Townsley B, Machuka J, Sinha N. 2008. Long-distance transport of mRNA via parenchyma cells and phloem across the host-parasite junction in Cuscuta. New Phytologist 179:1133−41

    doi: 10.1111/j.1469-8137.2008.02540.x

    CrossRef   Google Scholar

    [152]

    Huang NC, Jane WN, Chen J, Yu TS. 2012. Arabidopsis thaliana CENTRORADIALIS homologue (ATC) acts systemically to inhibit floral initiation in Arabidopsis. The Plant Journal 72:175−84

    doi: 10.1111/j.1365-313X.2012.05076.x

    CrossRef   Google Scholar

    [153]

    Yang HW, Yu TS. 2010. Arabidopsis floral regulators FVE and AGL24 are phloem-mobile RNAs. Botanical Studies 51:17−26

    Google Scholar

    [154]

    Chitwood DH, Nogueira FTS, Howell MD, Montgomery TA, Carrington JC, et al. 2009. Pattern formation via small RNA mobility. Genes & Development 23:549−54

    doi: 10.1101/gad.1770009

    CrossRef   Google Scholar

    [155]

    Baldrich P, San Segundo B. 2016. MicroRNAs in rice innate immunity. Rice 9:6

    doi: 10.1186/s12284-016-0078-5

    CrossRef   Google Scholar

    [156]

    Pant BD, Buhtz A, Kehr J, Scheible WR. 2008. MicroRNA399 is a long-distance signal for the regulation of plant phosphate homeostasis. The Plant Journal 53:731−38

    doi: 10.1111/j.1365-313X.2007.03363.x

    CrossRef   Google Scholar

  • Cite this article

    Li M, Niu X, Li S, Li Q, Fu S, et al. 2023. Intercellular signaling across plasmodesmata in vegetable species. Vegetable Research 3:22 doi: 10.48130/VR-2023-0022
    Li M, Niu X, Li S, Li Q, Fu S, et al. 2023. Intercellular signaling across plasmodesmata in vegetable species. Vegetable Research 3:22 doi: 10.48130/VR-2023-0022

Figures(2)  /  Tables(2)

Article Metrics

Article views(4054) PDF downloads(503)

REVIEW   Open Access    

Intercellular signaling across plasmodesmata in vegetable species

Vegetable Research  3 Article number: 22  (2023)  |  Cite this article

Abstract: The formation of edible organs and stress adaption are two major focuses of the studies on vegetable species. The regulation of these two processes often involves cell-to-cell signaling. In most plants, including vegetable species, intercellular signaling can be delivered by mobile regulators that traffic through a channel called plasmodesmata connecting almost all cells. A large number of transcription factors and RNAs have been discovered to move across plasmodesmata (called the symplastic way) to travel a short-range or a long-distance. This symplastic transport of signaling molecules has emerged to be an important regulation of a wide range of developmental and physiological processes. Callose deposition to plasmodesmata is a key step controlling the plasmodesmata permeability in many cell types. Here we summarize the recent progress in our understanding of plasmodesmata-mediated signaling in plants.

    • First visualized by Robert Hooke in 1665, cells had long been regarded as individual units of a whole organism. Whether the cell represents an autonomous entity was a question that had been a subject of debate in 19th Century. The observation of intercellular bridges and plasmodesmata supports the idea that the cellular structure forms the protoplasmic continuity, highlighting the importance of reciprocal interaction of cells within a multicellular organism. As a pioneering cell biologist, Wilson wrote in 1923, "it is the 'organism as a whole' and a 'property of the system as such' "[1], almost all plant cells are connected by the intercellular channel called plasmodesmata (PD)[2].

      Primary PD is a straight channel-like structure, as small as 30-50 nm in diameter, connecting two neighboring plant cells[3,4]. A major component of this channel is an endoplasmic reticulum (ER) derived central membranous strands called desmotubles, which form presumably through trapping ER strands in the cell plate during cytokinesis[5,6]. In between the desmotubule and flanking plasma membrane is the cytosolic space called cytoplasmic sleeve[7,8]. Components including cytoskeletons, a GPI-anchor protein and PD localizing proteins (PDLP) have been suggested to participate in the organization and function of plasmodesmata[9, 10].

      More recently, sphingolipids were found to affect the pore size of plasmodesmata[11]. Interestingly, analysis of Physcomitrium patens plasmodesmata proteome suggested the enrichment of cell-wall located proteins including EXORDIUM-family members and xyloglucan transglycosylases in plasmodesmata[12]. In particular, this study identified callose-degrading glycolyl hydrolase family 17 (GHL17) proteins as an abundant PD protein family[12], suggesting the potentially conserved plasmodesmata regulation by callose (will be further discussed later in this review) over the evolution.

      Smaller molecules, ions and metabolic substance can all pass through PD by diffusion. Other micro-molecules including proteins and RNAs are thought to transverse PD via active transport[1115]. Mobile molecules can move across PD via either the cytoplasmic sleeve, or through the desmotubule (in lumen or lateral diffusion in the desmotubule membrane), or via diffusion in the flanking plasma membrane[16,17]. In support of these hypotheses, it was found that the interference of the membrane structure affected PD permeability[17]. In old tissues, plant cells further produce secondary PD that is normally branched and complex in shape. Localized cell wall modification could be involved in secondary PD formation, and the complexity of this type of PD is correlated with reduced PD permeability[18,19]. Nevertheless,the detailed mechanism and the exact roles of secondary PD during development are still far from clear. Interestingly, multiple types of PD were found at grafted wounds, suggesting that different PD types could have distinct functions[20]. In this review, we focus on our current understanding of cell-to-cell signaling across plasmodesmata.

    • The observation of cell-to-cell movement of large molecules initially arose from the micro-injection of fluorescent dye in plant tissues[2124]. The first endogenous protein exhibiting the intercellular mobility is KNOTTED1 (KN1), a homeodomain protein essential for maintenance of the shoot apical meristem (SAM) in maize[25,26]. Recently, the ribosomal RNA-processing protein 44A (AtRRP44A) was shown to mediate the cell-to-cell trafficking of KN1[27]. Since then, a large number of transcription factors were identified in plants that can move between tissues and cells to provide positional instruction during plant development[21]. These mobile regulators can traffic across just a few cell layers to function locally or over a long distance to affect global developmental change.

      One of the central questions in organogenesis is how to spatiotemporally maintain stem cells and specify cell fates. In SAM, WUSCHEL (WUS) is expressed in the organizing center of shoot apical meristem, but the protein moves to the layer1 and 2 (L1 & 2) of shoot apical meristem where WUS triggers CLAVATA 3 (CLV3) expression, which in turn inhibits WUS transcription in L1 and L2 layer[28,29]. With this WUS-CLV3 feedback loop, plants can maintain the stem cell population in proper size in SAM. With the similar strategy, plants maintain the root stem cell niche via WOX5-CLE40 loop, in which WOX5 traffics from quiescent center (QC) to columella stem cell (CSC) to repress the cell differentiation[30]. In Arabidopsis, SHOOT MERISTEMLESS (STM) and ARABIDOPSIS KNOTTED-LIKE (KNAT1)/BREVIPEDICELLUS (BP) are two homologs of the KN1 gene, previously described to be mobile in maize SAM. When driven by an L1 specific promoter, STM and KNAT1 were observed to move from the L1 layer into the inner cell layers of the SAM[31,32]. In addition, KNAT1 was able to pass the interface between cortex and epidermis in Arabidopsis when mis-expressed by a mesophyll specific promoter[33].

      In embryogenesis, TARGET OF MONOPTEROS 7 (TMO7), encoding a bHLH transcription factor, is essential for hypophysis, the founder cell for forming root apex during post-embryonic growth. TMO7 is transcribed in embryonic cells while the TMO7-GFP fusion can be detected in the neighboring hypophysis, indicating a non-cell-autonomy of this regulator[34,35]. In post-embryonic growth, intercellular movement of transcriptional factors regulates a variety of developmental aspects ranging from root radial patterning to root hair and trichome initiation. These mobile regulators including SHORT-ROOT (SHR), CAPRICE (CPC), TRANSPARENT TESTA GLABRA 1 (TTG1), GLABRA 3 (GL3), ENHANCER OF TRY AND CPC 3 (ETC3)/ TRIPTYCHON (TRY), UBIQUITIN-SPECIFIC PROTEASE (UBP1) have been well reviewed previously[15,21]. A previous screen estimated that around 15% of transcriptional factors in roots can move between cells[36]. In contrast, we only have limited understanding of the functionality of these mobile proteins.

      Recently, more mobile transcriptional factors have been identified (summarized in Table 1). Two closely related AT-hook family members, AT-HOOK MOTIF NUCLEAR LOCALIZED PROTEIN 3 (AHL3) and AHL4, were shown to interact in vivo and regulate the boundaries between the procambium and xylem[37]. Interestingly, their interaction seemed to be required for their intercellular trafficking. A SHR target, SCL23 displays a bidirectional radial spread and long-range movement into meristem in Arabidopsis roots. Through direct interaction, SCL23 controls movement of SHR and participate in endodermal specification in the root meristem[38].

      Table 1.  Summary of the mobile transcription factors identified in plants.

      Mobile TFsFunctionMoves from:toReference
      HY5Root growth and N uptakeShoot-to-rootChen et al. (2016)[41]
      DWARF14Regulate the development of AMsThrough phloem into axillary meristems (AMs)Kameoka et al. (2016)[139]
      BdMUTEBdMUTE is required for subsidiary cell formationGMCs to neighboring cell filesRaissig et al. (2017)[97]
      SPCHStomatal cell fateCell-to-cell diffusion in the leaf epidermis of chorusGuseman et al. (2010)[96]
      AN3Leaf developmentFrom the mesophyll to the epidermis in leavesKawade et al. (2013)[140]
      WUSMeristem maintenanceFrom the organizing centre to L1, L2 layersYadav et al. (2011)[28]
      KN1/STMMeristem maintenanceBroadly in the SAMKim et al. (2003)[31], 2005[32]
      PLT2Longitudinal root zonationLongitudinally from the root meristem forming a gradientMahonen et al. (2014)[141]; Galinha et al. (2007)[142]
      SHRRoot radial patterning and RAM maintenanceWithin Stele; Stele into endodermis, QC, CEI and CEDKoizumi et al. (2011)[44], Nakajima et al. (2001)[78]
      AHL3/AHL4Xylem specificationFrom procambium cells to the xylemZhou et al. (2013)[37]
      WOX5Stem cell maintenanceQC to CSCPi et al. (2015)[30]
      TMO7Recruitment of the hypophysisEmbryo into the upper cell of suspensorSchlereth (2010)[34]; Lu et al. (2018)[35]
      Cyp1Root growthFrom leaves to root in tomatoSpiegelman et al. (2015)[143]
      UBP1Transition from cell division to elongationStele and LRC to cells into transition/elongation zoneTsukagoshi et al. (2010)[144]
      SCL23Endodermal cell fateBidirectional radial spread and movement into meristemLong et al. (2015)[38]
      TTG1Trichome patterningAtrichoblasts into trichome initials
      CPCTrichome patterning, root hair initiationTrichome initials into Atrichoblasts; non-root hair cell into root hair cellWester et al. (2009)[90]
      GL3/EGL3Root hair initiationRoot hair cell into non-root hair cellKang et al. (2013)[91]

      Besides the local regulation, transcriptional factors were also found to traffic long-distance between organs to direct global developmental transition in plants in Fig 1. An early example is the detection of FLOWERING LOCUS T (FT) trafficking from leaves where it is synthesized in response to day length, to the SAM to trigger flowering[39,40]. Recently, a light-activated transcriptional factor, ELONGATED HYPOCOTYL 5 (HY5) was shown to move via phloem from shoot-to-root. This translocation of HY5 was proposed to mediate light-activated root growth and N uptake from the soil to balance photosynthetic carbon fixation in the leaf[41].

      Figure 1. 

      Mobile proteins and RNAs in plant development and stress response. The mobile regulators participate widely in the development of different organs (as illustrated). They can travel short-range to regulate local tissue patterning or long-distance to transduce systemic signaling. Gray arrow: phloem-based long-distance movement. WUS and STM regulate SAM maintenance; SPCH, BdMUTE, AN3, TTG1, GL3 and CPC are involved in epidermal patterning. In roots, PLT2, SHR, AtDof4.1, AHL3/AHL4, WOX5, TMO7, UBP1 and SCL23 govern a variety of processes including cell division, radial patterning, stem cell maintenance and developmental transition. Long-distance signaling regulators such as FT and HY5 can traffic from leaves to SAM to promote flowering, and from shoot to root to regulate root growth and nitrate uptake respectively. Environmental stresses can induce PD closure. Small RNAs including miR399d, 827 and 2111 move from aerial parts to roots in response to phosphate starvation.

      Considering the size of transcriptional factors, PD seems to be the most possible way for the intercellular translocation. With an iclas3m system (described in detail in a later part of this review) that blocked the PD between stele and endodermis, SHR intercellular transport was terminated[3]. Another piece of evidence supporting PD transport of transcriptional factors is the blocked movement of TMO7 from meristematic cells into the root cap in the cals3-2d, a mutant in which PD is restricted by over-accumulated callose[35]. To get access to PD, transcriptional factors could exploit intracellular apparatus including microtubules and endomembrane delivery system[42,43]. Besides, an unknown function protein named SHR INTERACTING EMBRYONIC LETHAL (SIEL) was shown to interact with a number of mobile transcriptional factors and the mutation of this gene seemed to reduce SHR intercellular movement[44]. As SIEL partially localized to endosomes, it was proposed that this protein could function as a 'shuttle' to facilitate delivery of mobile transcriptional factors. In addition, some facilitating proteins have also been identified. After passing through PD, a few mobile proteins including APS KINASE 1 (KN1), SHOOT MERISTEMLESS (STM) and TRANSPARENT TESTA GLABRA 1 (TTG1) were discovered to associate with a group of type II chaperonin complexes consisting of CHAPERONIN CONTAINING T-COMPLEX POLYPEPTIDE-1 SUBUNIT 7 and 8 (CCT7 & CCT8), which facilitate the movement possibly by promoting the protein refolding after the PD cross-over[27].

      Although no specific domain has been identified that accounts for intercellular mobility, the cell-to-cell transport of transcriptional factors seemed to be protein sequence-dependent. Homeodomain (HD) and the helical domains have been shown to be necessary and sufficient for PD-mediated transport of KN1. Unlike this, three conserved domains (HD, WUS-box, and EAR-like domain) in WUS are not required for its movement. Instead, WUS mobility seems to be controlled by a non-conserved sequence between the HD domain and WUS-box[29]. Despite triple GFP Tag impaired TMO7 movement, protein size did not seem to be the primary determinant of intercellular transport. Instead, TMO7 was found to move in a sequence-dependent manner, and both nuclear residence and protein modification are important for TMO7 mobility[35]. In two other mobile transcriptional factors, CPC and SHR, the mobility relied on multiple regions within the proteins. In addition, the mobility of these two proteins seemed to be associated with the subcellular distribution in both the cytoplasm and the nucleus.

      In addition to transcriptional factors, small RNAs also participate in transcriptional regulation of diverse developmental and physiological events in plants. Small RNAs are 21−24 nt long and can be generally divided into siRNAs and miRNAs[45]. Small RNAs function either through degrading target genes by near-perfect complementarity, or via transcriptional silencing by histone modification and DNA methylation[4650]. Small RNAs were often regarded as the long-distance signals as the initial efforts dissecting their mobility exploited the grafting system in which mutants defective in small RNAs biogenesis were included. Facilitated by high-throughput sequencing techniques, researchers identified a large number of mobile siRNAs that can traffic from shoot to root presumably via phloem. Besides siRNA, a large number of miRNAs were discovered to traffic in phloem exudates over long distance. Low-phosphate induced miR399s exhibited a shoot-to-root movement to repress downstream targets including PHO2 in the root[51]. Similarly,miR399d, miR827 and miR2111 were all found in grafting experiments to relocate from aerial parts to roots in response to phosphate starvation[52]. During rhizobial infection, miR2111 functioned as long-distance signals to post-transcriptionally regulate symbiosis suppressor TOO MUCH LOVE in roots[53]. miR395 can also translocate from wild-type scions to rootstocks of the miRNA processing mutant hen1-1 to target the APS gene[54]. In addition, both miR156 and miR172 have been confirmed as potentially phloem-mobile miRNAs that regulate tuber formation[5557].

      In grafting system, only small RNAs transporting from shoot-to-root via phloem could be analyzed. Other approaches that allow for the comparison between the expression areas and in situ RNA distribution patterns may help the identification of small RNAs acting locally as non-cell autonomous signals. To establish adaxial–abaxial leaf polarity, a member of Trans-acting small interfering RNA (ta-siRNA) family forms a gradient across the leaves by intercellular diffusion. This diffusion-driven pattern of ta-siRNA shapes the expression pattern of AUXIN RESPONSE FACTOR3 (ARF3), an abaxial determinant gene. Another small RNA, miR390 was proved to regulate the leaf polarity by the cell-to-cell movement from vasculature and pith region below the shoot apical meristem to the vegetative apex[54]. In addition, miRNA165/166 were discovered to move from the endodermis into the stele to regulate the xylem cell fate[58]. Moreover, miR394 was shown to regulate stem cell maintenance in SAM by the PD-mediated movement from L1 to inner cell layers to repress LEAF CURLING RESPONSIVENESS (LCR) expression[59].

      In addition to siRNA and miRNA, mRNAs have also been found to travel beyond the cells in which they are expressed in Fig 1. In addition to the early example of mobile mRNAs of KN1, potato sucrose transporter SUC1 mRNA was also confirmed to be mobile. In grafting experiments, a number of mRNAs were found to travel, such as FT, FVE and AGL24 in Arabidopsis[60], Aux/IAA in melon and Arabidopsis[61], PP16 and NACP in pumpkin[62,63], BEL5 and POTH1 in potato, SLR/IAA14 in apple[64], PFP-T6 and PS in tomato[65] (summarized in Table 2). Recently, Luo et al. developed a fluorescence-based mRNA labeling system to identify mobile mRNAs targeted to PD[66]. Their analyses revealed that only mobile rather than not non-mobile mRNAs were selectively targeted to PD, providing further evidence for PD mediated transport of mRNAs. Interestingly, using a Nicotiana benthamiana/tomato heterograft system, Xia et al. found some mRNAs have bidirectional mobility between shoots and roots. In addition, forced expression of non-mobile mRNAs in the companion cells did not confer the mobility[6771]. Thus, the movement of mRNA is likely an actively regulated process. Moreover, a large number of graft-transmissible mRNAs have been identified by high throughput sequencing in a variety of species including Arabidopsis, tobacco, grape, cucumber and tomato[6772].

      Table 2.  List of mobile RNAs with functions in organ development.

      Mobile factorFunctionMoves from: toReference
      mRNA
      KN1SAM maintenanceinjected cell to neighbouring cellsLucas et al. (1995)[26]
      SUC1Sucrose transportcompanion cells to sieve elementsKuhn et al. (1999)[145]
      FT1Flowering inductionLeaf to SAMLu et al. (2012)[60]
      Aux/IAA18Root developmentLeaf to rootNotaguchi et al. (2012)[61]
      PP16RNA transportPhloem to shoot apexXoconostle-Cazares et al. (1999)[62]
      NACPMeristem maintenancePhloem to shoot apexRuiz-Medrano et al. (1999)[146]
      StBEL5Tuber formationLeaf to rootBanerjee et al. (2009)[147]
      POTH1Leaf developmentLeaf to rootMahajan et al. (2012)[148]
      SLR/IAA14Lateral root formationShoot to rootKanehira et al. (2010)[64]
      PFP-T6Leaf developmentLeaf to leaf primordiaKim et al. (2001)[65]
      PSPathogen resistanceShoot to root and vice versaZhang et al. (2018)[149]
      GAILeaf developmenthost to parasiteRoney et al. (2007) [150]; David-Schwartz et al. (2008)[151]
      ATCFloral initiationLeaf to flower apicesHuang et al. (2012)[152]
      FVEfloral regulatorsRoot to SAMYang and Yu (2010)[153]
      AGL24floral regulatorsRoot to SAMYang and Yu (2010)[153]
      siRNA
      ta-siRNAEstablishment of leaf polaritythe adaxial to the abaxial side of the leafChitwood et al. (2009)[154]
      hc-siRNADNA methylationShoot to rootBaldrich et al. (2016)[155]
      miRNA
      miR165/166Xylem specificationendodermis into the steleCarlsbecker et al. (2010)[58]
      miR390Leaf polarityvasculature and pith region below the SAM to SAMChitwood et al. (2009)[154]
      miR394Meristem maintenanceL1 to inner layers in the shoot meristemKnauer et al. (2013)[59]
      miR395Sulfate homeostasisgraft unionsBuhtz et al. (2010)[54]
      miR399dPhosphate homeostasisshoot to root and vice versaPant et al. (2008)[156]; Lin et al. (2008)[51]
      miR172regulate tuber formationLeaf to rootMartin et al. (2009)[55]
      miR2111Phosphate homeostasis;
      Rhizobial infection;
      shoot to root and vice versaHuen et al. (2017)[52];
      Tsikou et al. (2018)[53]
      miR827Phosphate homeostasisshoot to root and vice versaHuen et al. (2017)[52]
    • A plant organ is usually composed of morphologically and functionally different cell types in different positions. Small molecules can move between cells and across plasmodesmata, which mediates crucial intercellular communication for the growth and development of plant tissues and organs. For example, a plant root is composed of concentrically arranged cell layers with epidermis, cortex, endodermis, and stele locating from outside to inside[73]. This anatomic arrangement highlights the regulation of tissue patterning instructed by positional information, often through the exchange of signaling molecules between cells. A number of developmental processes including root radial patterning, root hair initiation and trichome formation, have emerged as the model system for studying tissue patterning in plants.

      In root, the formation of the endodermal cell layer starts from the endodermal and cortex initial cells in root stem cell niche, where two transcriptional factors, SHR and SCARECROW (SCR) promote the expression of CYCD6;1 to allow the switch of cell division pattern from anticlinal to periclinal[7477]. This results in the formation of two distinct layers of cells within the ground tissue, and the role of SHR in specifying the endodermal layer was proposed based on the fact that the endodermal layer was completely absent in shr-2 mutant. Intriguingly, SHR expression is restricted in stele, but the SHR protein is actively transported through PD from stele toward the outside to play non-cell-autonomous roles[78,79]. In the enodermis, SHR directly activates SCR which, in turn, physically binds to SHR to trap this mobile transcription factor in the nucleus of the endodermis, preventing further movement[77]. This mechanism was discovered to be conserved in rice and thus was proposed to be an evolutionarily conserved mechanism defining a single endodermal cell layer in almost all land plants[74]. However, a study on rice SHR homologs suggested that SHR alone is insufficient to determine endodermal cell fate[80]. Consistent with this argument, mis-expression of SHR indicated that SHR ability to confer endodermal identity partially relied on cell lineage and was coordinated by uncharacterized positional information, presumably derived from stele.

      Specific expression of marker genes, as often used previously to determine endodermal cell fate, is sometimes misleading. A prominent feature of the endodermis is the formation of the Casparian Strip (CS), an apoplastic barrier between vascular tissues and outer ground tissues[81]. The presence of functional CS is therefore a better trait for precise evaluation of endodermal identity. Two recent studies revealed that SHR does serves as a master regulator activating a hierarchical downstream network for CS formation[82,83]. The combination of SHR mediated cascade and another independent peptide signal derived from stele forms the minimum set of regulators that program endodermal identity, exemplified by the formation of functional CS[83]. Since both SHR and the peptide are specifically expressed in vascular tissues, CS formation represents the elaborate developmental control by stele-to-endodermis movement of mobile regulators. Besides CS, SHR and its downstream target SCR can activate the expression of miRNA165/166 in the endodermis which in turn moves back to vasculature to repress a class III homeodomain-leucine zipper transcription factors for proper xylem formation[58]. Thus the reciprocal communication between ground tissue and vasculature in root spatially defines the radial patterning in root. In Cardamine, a recent study indicated that a differential spatial distribution of miR165/166 is responsible for forming the extra cortex layer[84]. In addition to roots, miR165/166 also function in other organs including leaf primordial and ovule. By restricting PHB expression in incipient inner integument, miR165/166 promotes the correct ovule patterning[85]. Interestingly, a callose synthase mutant in maize, named tie-dyed2 (tdy-2), affects the development of vasculature, suggesting the mechanism of vascular development directed by intercellular communication (possibly via miR165/166) is likely conserved in crops[86,87]. In addition to roots, plasmodesmata also plays a key role in regulating leaf development, particularly the formation of leaf veins[88].

      Trichomes and root hairs, originating from the epidermis in leaves and roots respectively play important roles in protecting plants from bio/abiotic stresses, and promoting nutrient absorption[89,90]. In Arabidopsis, the initiation of trichomes and root hairs is precisely patterned in epidermis, indicating an essential role of cell-to-cell communication in these processes.

      In trichome initiation, both positive regulator TRANSPARENT TESTA GLABRA (TTG1) and negative regulator ENHANCER OF TRY AND CPC 3 (ETC3) and CAPRICE (CPC) move between cells. In incipient trichome cells, TTG1 protein accumulates through a trapping/depletion mechanism mediated by GLABRA3 (GL3)[91]. On the other hand, the repressor of ETC3 and CPC move into the neighboring non-trichome cells (also regulated by GL3), forming inactivated MYB/bHLH/WD40 to inhibit the development towards trichomes[92]. Recently, PdBG4 has been implicated in regulating PD permeability in Arabidopsis trichome development[93]. In root hairs, CPC serves as a positive regulator and it is trapped in the hair-position root epidermis by interacting with EGL3 and GL3 after the movement[94]. The trn1 mutant is defective in the position-dependent pattern of root hairs and cause the ectopic expression of WER, GL2 and EGL3, suggesting that TRN1 also participates in the position-dependent cell fate determination[95,96].

      Stomata on epidermis are responsible for water and gas exchange between the plants and the environments. The mature stomata structure is produced through successive cell division and differentiation process, with both processes subject to highly spatiotemporal regulation[97]. In a GLUCAN SYNTHASE-LIKE 8 (GSL8) mutant in which normal callose deposition is disrupted, SPCH-GFP diffused to neighboring cells from meristemoids, resulting in excessive proliferation of stomatal-lineage cells. This observation suggests that proper gating of critical regulators, likely through callose regulation, regulates the correct patterning of stomata complex[98]. MUTE, another key transcriptional factor required to terminate asymmetric division and promote the transition of meristemoids to GMCs, was shown in Brachypodium to move from GMCs to neighboring cells to induce the subsidiary cells (SCs) formation[99].

    • Plants respond to stresses often by accumulation of callose, which is negatively correlated with PD permeability in Fig 2. A variety of abiotic stresses have been associated to callose induction, such as cold stress[100,101], wounding[102,103], heat stress[104,105], and heavy metals[106109]. Although detailed mechanism is not entirely clear, callose synthases were found to participate in the callose regulation. In Arabidopsis, there are 12 callose synthase (CalS) family members. When exposed to excess iron, the cals5 and cals12 mutants showed an attenuated callose deposition in phloem, compared to wild type and other cals mutants. This result suggests that cals5 and cals12 may play specific roles in iron stress response in Arabidopsis[110]. In tomato, cold stress has long been known to cause catfacing fruits or malformed fruits by breeders and gardeners. A recent study proved this phenomenon was caused by the restriction of SlWUS intercellular movement via plasmodesmata in floral meristem[101]. The cold induced callose accumulation blocked the plasmodesmata, resulting in the excessive activation of CLV3 and TAG1, and disrupted WUS-CLV3/WUS-TAG1 negative feedback loops[101].

      Figure 2. 

      Regulation of PD permeability by callose. (a) Schematic illustration of regulation of the PD aperture by callose deposition in flanking regions of PD. Induced callose accumulation closes PD permeability and blocks the intercellular movement of transcription factors and small RNAs. (b) The design of ‘icals3m’ system that can inducibly (via estradiol induction cassette) promote callose deposition in specific cell types (via cell-type specific promoters)[128],[138].

      It has been reported that PD regulation serves as an innate defense strategy[111]. Pathogens trigger both pathogen-associated molecular pattern (PAMP) and PAMP-triggered immunity (PTI) systems, which have been reported to induce callose deposition[112]. Upon SMV virus invasion, callose was accumulated in soybean phloem which prevents the virus from traveling long distances[113]. Salicylic acid (SA) is a plant immune signal produced upon pathogen infection, which has also been shown to trigger PD closure and affect symplastic communication. Elevation of SA level seemed to be necessary for the PD response during bacterial infection, and the expression of bacterial derived salicylate hydroxylase (NahG) gene in plants resulted in higher susceptivity to bacteria[113]. Biotic stresses including pathogen infection are known to modulate ROS level and callose abundance in infected regions, which is presumably responsible for the altered PD permeability[114,115].

      Virus can also regulate the mesenchymal plasmodesmata in tobacco[109] and it was recently reported that ROS-mediated PD closure is controlled by multiple pathways, either in SA- or PDLP5-dependent manners. Change of callose level in biotic stresses is also modulated by callose synthase members[112,113]. SA-dependent PD regulation requires the function of callose synthase1 (CalS1). However, the CalS8 seemed to be more involved in basal and ROS-dependent PD regulation[103]. Callose synthase members have also been widely reported in recent years. CsCalS4 function was identified in pollen development in cucumber, and CsCalS1/8 homologous genes were induced by cucumber fungus and functioned as the key factors in response to biological stress[114]. GhCalS5 and ZmCals were found to promote callose synthesis in cotton and maize in responsive to stresses[116,117].

      In addition, PD-localized proteins also emerged as the regulator of PD aperture during biotic stresses. It was shown that the PD closure triggered by chitin was dependent on the activity of PD-localized receptor-like protein LYM2[111]. Besides, bacterial flagellin could rapidly activate the expression of CML41, a PD-localized Ca2+-binding protein, which is necessary for the induction of callose at PD.

      Callose is the linear polysaccharide that is composed of β-1,3-glucan. It is a component of cell wall and is frequently found to deposit at PD, where it is believed to control the PD permeability during plant development and stress response. It was found the precise developmental transition often relies on the regulation of symplastic continuity. In birch, bud dormancy entry and release are associated with the shift between callose production and turnover. Callose accumulation at PD in the shoot apical and rib meristems can seal off the symplastic communication and promote the bud dormancy[116121]. A period of chilling, however, triggers gibberellin biosynthesis, resulting in increased expression of 1,3-β-glucanases and degradation of callose. Accumulating evidence suggests that callose regulation is actually implicated in a wide range of developmental processes, including seed germination, embryogenesis, cell division, flowering and reproduction[122124]. In tomato, a short period of cold stress is sufficient to induce callose accumulation in floral meristem and blocked intercellular movement of SlWUS, resulting in malformed fruits[101]. In olives, callose deposition, as part of cell wall modification, regulates fruit abscission[114].

      Through a genetic screen for defective vascular development, Vaten et al. (Helariutta group) identified three semi-dominant alleles of CALLOSE SYNTHASE 3 (cals3d) that caused an increase in callose deposition at PD and abnormal plant growth[3,19]. In the root, cals3d mutants all showed aberrant radial patterning and misspecification of the phloem and the xylem. Consistent with these phenotypes, cals3d roots exhibited decreased PD-mediated symplastic movement of free GFP, SHR and miRNA165/66[3,125]. It thus seemed that the identified dominant mutations can substantially enhance the ability of CALS3 to promote callose deposition at PD. By combining these mutations in a vector containing LexA-VP16-ER (XVE)-based estradiol inducible cassette, the Helariutta group designed an elegant tool named as the 'icals3m system'. Driven by specific promoters, this system can potentially be used to temporally manipulate callose at PD and symplastic communication in particular cell types[3,126].

      The initial attempts using this system in vascular tissues and lateral root development proved to be successful[3,125,127]. With specific induction of icals3m system in xylem pole pericycle, Benitez-Alfonso et al. detected a significantly increased number of initiated primordial [126]. Together with the observation of a transient symplastic isolation of the primordium prior to emergence, they confirmed the essential role of callose based symplastic connectivity between pericycle cells, founder cells, and the neighboring tissue during lateral root patterning[122]. More recently, icals3m system was used to dissect the roles of symplastic communication in root apical stem[122]. Driven by an endodermis-specific EN7 promoter, icals3m induced symplastic blockage led to severe root patterning defects, shown by disrupted cell division direction, misspecification of cell fate as well as impaired cell polarity. In root tip, different cell types including endodermis all derived from the root stem cell niche, where QC was believed to repress the differentiation of surrounding stem cells based on an early classic laser ablation experiment carried out in the 1990's[127]. However, icals3m system provides an alternative non-invasive approach to examine the role of QC. With the expression under WOX5 promoter, icals3m system was clearly shown to induce callose specifically in QC[128]. The visible callose signal based on aniline blue staining was detected as quickly as 6 h after the estradiol induction[129]. This icals3m system was further used to study the interaction between root cap and the root meristem[124,128130]. When the symplastic communication between root cap and root meristem was disrupted, developmental defects were observed in both parts: In meristem, stem cell maintenance was affected while in root cap the starch granules, the marker commonly used as an indicator of columella differentiation, disppeared[125]. An earlier study showed that starch granules in columella cells relied on auxin concentration[131]. In this study, short-term disruption of symplastic communication was sufficient to cause defects in stem cells, while it took longer for auxin distribution in root meristem to occur[125]. In fact, plasmodesmata itself can act as the channel for auxin flow[131,132]. Furthermore, icals3m system also was employed in the study of phloem unloading[132]. A phloem pole pericycle specific promoter CalS8 and a companion cell and metaphloem sieve element specific promoter psAPL were both used to drive icals3m to block the connection between different phloem cell types[133]. A direct developmental defects arose from the blocked plasmodesmata in phloem was the reduced growth of axillary buds[50].

      To summarize, callose regulation is a central mechanism to control symlastic communication during plant development. Spatiotemporal expression of icals3m system can be an effective tool to deepen our understanding of the developmental regulation mediated by symplastic signals. The power of this system can be even higher with the combination with other techniques including cell type specific OMICs. The application of this system in vegetable studies would greatly enhance our ability to dissect various aspects of development and physiology in vegetable species ranging from fruit development to stress resistance.

    • Intercellular signaling across plasmodesmata plays crucial roles in a wide range of processes in plants. The currently identified signaling molecules across plasmodesmata are mainly transcription factors and RNAs. However, accumulating evidence suggests that many other signaling pathways including calcium signaling, redox signaling, phosphorylation signaling, and hormone signaling can also function in non-cell-autonamous manner[134136]. As these pathways are often complex and interplay with each other, it is still difficult to unravel such non-cell-autonamous functions. With the advance in high-resolution imaging techniques, such as super-resolution microscopy, researchers will be able to visualize in vivo the action and mobility of the molecular players involved in intercellular signaling[137].

      In addition to visualizing the intercellular mobility of molecules, it is crucial to precisely evaluate the phenotype with a specific intercellular signaling disrupted. Developing cell type specific approaches is the key step and thus identification of promoters with restricted expression in certain cell types is important. Furthermore, abolishing gene function in a specific cell type is a valuable tool for studying intercellular signaling. Previously, cell-specific RNAi was employed but the intercellular mobility of small RNAs prevents the precise evaluation of gene function. Recent rapid development of CRISPR-Cas9 technique has emerged as a powerful tool for this purpose. The combination of cell-specific expression of Cas9 with reporters that allows for visualizing the gene editing in different cells could greatly enhance our ability to precisely evaluate the function of mobile regulators.

      Lastly, to gain a more comprehensive understanding of the plasmodesmata mediated intercellular signaling, it is important to integrate multiple approaches, such as high-resolution imaging, single-cell technique, multi-omics, and computational modeling. Although the cell-to-cell signaling often occurs locally, the impact could be systemic in plants. The complete assessment of plasmodesmata-mediated intercellular signalling, as well as derived tissue- or cell-type-specific techniques, will not only benefit the study of plant development, but also provide the opportunity for future biotechnological renovation of plants.

      • This work was supported by carbon-nitrogen high efficiency grants from Fujian Agriculture and Forestry University (118992201A).

      • The authors declare that they have no conflict of interest.

      • Copyright: © 2023 by the author(s). Published by Maximum Academic Press, Fayetteville, GA. This article is an open access article distributed under Creative Commons Attribution License (CC BY 4.0), visit https://creativecommons.org/licenses/by/4.0/.
    Figure (2)  Table (2) References (156)
  • About this article
    Cite this article
    Li M, Niu X, Li S, Li Q, Fu S, et al. 2023. Intercellular signaling across plasmodesmata in vegetable species. Vegetable Research 3:22 doi: 10.48130/VR-2023-0022
    Li M, Niu X, Li S, Li Q, Fu S, et al. 2023. Intercellular signaling across plasmodesmata in vegetable species. Vegetable Research 3:22 doi: 10.48130/VR-2023-0022

Catalog

    /

    DownLoad:  Full-Size Img  PowerPoint
    Return
    Return