Structure and growth of plant cell walls.

Journal

Nature reviews. Molecular cell biology
ISSN: 1471-0080
Titre abrégé: Nat Rev Mol Cell Biol
Pays: England
ID NLM: 100962782

Informations de publication

Date de publication:
15 Dec 2023
Historique:
accepted: 15 11 2023
medline: 16 12 2023
pubmed: 16 12 2023
entrez: 15 12 2023
Statut: aheadofprint

Résumé

Plant cells build nanofibrillar walls that are central to plant growth, morphogenesis and mechanics. Starting from simple sugars, three groups of polysaccharides, namely, cellulose, hemicelluloses and pectins, with very different physical properties are assembled by the cell to make a strong yet extensible wall. This Review describes the physics of wall growth and its regulation by cellular processes such as cellulose production by cellulose synthase, modulation of wall pH by plasma membrane H

Identifiants

DOI: 10.1038/s41580-023-00691-y PMID: 38102449
pubmed: 38102449
doi: 10.1038/s41580-023-00691-y
pii: 10.1038/s41580-023-00691-y
doi:

Types de publication

Journal Article Review

Langues

eng

Sous-ensembles de citation

IM

Informations de copyright

© 2023. Springer Nature Limited.

Références

Bidhendi, A. J. & Geitmann, A. Finite element modeling of shape changes in plant cells. Plant. Physiol. 176, 41–56 (2018).
pubmed: 29229695 doi: 10.1104/pp.17.01684
Coen, E. & Cosgrove, D. J. The mechanics of plant morphogenesis. Science 379, eade8055 (2023). A concise overview of mechanics, from fibres to fibre networks to tissues and organs, touching on some of the open questions and divergent views in plant mechanobiology.
pubmed: 36730409 doi: 10.1126/science.ade8055
Echevin, E. et al. Growth and biomechanics of shoot organs. J. Exp. Bot. 70, 3573–3585 (2019).
pubmed: 31037307 doi: 10.1093/jxb/erz205
Zhang, Y. et al. Molecular insights into the complex mechanics of plant epidermal cell walls. Science 372, 706–711 (2021). This paper uses coarse-grained molecular dynamics to simulate the physical properties of cellulose, xyloglucan and hemicellulose and to investigate wall assembly and wall mechanics.
pubmed: 33986175 doi: 10.1126/science.abf2824
Jarvis, M. C. Forces on and in the cell walls of living plants. Plant Physiol., kiad387 https://doi.org/10.1093/plphys/kiad387 (2023).
Bar-On, Y. M., Phillips, R. & Milo, R. The biomass distribution on Earth. Proc. Natl Acad. Sci. 115, 6506–6511 (2018).
pubmed: 29784790 pmcid: 6016768 doi: 10.1073/pnas.1711842115
Bastin, J. F. et al. The global tree restoration potential. Science 365, 76–79 (2019).
pubmed: 31273120 doi: 10.1126/science.aax0848
Liao, Y. et al. A sustainable wood biorefinery for low-carbon footprint chemicals production. Science 367, 1385–1390 (2020).
pubmed: 32054697 doi: 10.1126/science.aau1567
Parre, E. & Geitmann, A. Pectin and the role of the physical properties of the cell wall in pollen tube growth of Solanum chacoense. Planta 220, 582–592 (2005). This paper shows that pectin viscoelasticity plays a big role in the growth of pollen tubes, whose walls are pectin rich and cellulose poor and that regions of de-esterified pectin are mechanically stiffer.
pubmed: 15449057 doi: 10.1007/s00425-004-1368-5
Winship, L. J., Rosen, G. A. & Hepler, P. K. Apical pollen tube wall curvature correlates with growth and indicates localized changes in the yielding of the cell wall. Protoplasma 258, 1347–1358 (2021).
pubmed: 34414478 doi: 10.1007/s00709-021-01694-2
Miller, K., Strychalski, W., Nickaeen, M., Carlsson, A. & Haswell, E. S. In vitro experiments and kinetic models of Arabidopsis pollen hydration mechanics show that MSL8 is not a simple tension-gated osmoregulator. Curr. Biol. 32, 2921–2934.e3 (2022).
pubmed: 35660140 doi: 10.1016/j.cub.2022.05.033
Cosgrove, D. J. Relaxation in a high-stress environment: the molecular bases of extensible cell walls and cell enlargement. Plant. Cell 9, 1031–1041 (1997).
pubmed: 9254929 pmcid: 156977 doi: 10.1105/tpc.9.7.1031
Cosgrove, D. J. Characterization of long-term extension of isolated cell walls from growing cucumber hypocotyls. Planta 177, 121–130 (1989).
pubmed: 24212280 doi: 10.1007/BF00392162
Yamamoto, R., Shinozak, K. & Masuda, Y. Stress-relaxation properties of plant cell walls with special reference to auxin action. Plant. Cell Physiol. 11, 947–956 (1970).
doi: 10.1093/oxfordjournals.pcp.a074586
Cosgrove, D. J. Wall relaxation in growing stems: comparison of four species and assessment of measurement techniques. Planta 171, 266–278 (1987).
pubmed: 24227336 doi: 10.1007/BF00391104
Verbancic, J., Lunn, J. E., Stitt, M. & Persson, S. Carbon supply and the regulation of cell wall synthesis. Mol. Plant. 11, 75–94 (2018).
pubmed: 29054565 doi: 10.1016/j.molp.2017.10.004
Vaahtera, L., Schulz, J. & Hamann, T. Cell wall integrity maintenance during plant development and interaction with the environment. Nat. Plants 5, 924–932 (2019).
pubmed: 31506641 doi: 10.1038/s41477-019-0502-0
Frankova, L. & Fry, S. C. Biochemistry and physiological roles of enzymes that ‘cut and paste’ plant cell-wall polysaccharides. J. Exp. Bot. 64, 3519–3550 (2013).
pubmed: 23956409 doi: 10.1093/jxb/ert201
Spartz, A. K. et al. SAUR inhibition of PP2C-D phosphatases activates plasma membrane H
pubmed: 24858935 pmcid: 4079373 doi: 10.1105/tpc.114.126037
Li, Y., Zeng, H. Q., Xu, F. Y., Yan, F. & Xu, W. F. H. H
pubmed: 35231180 doi: 10.1146/annurev-arplant-102820-114551
Haruta, M., Sabat, G., Stecker, K., Minkoff, B. B. & Sussman, M. R. A peptide hormone and its receptor protein kinase regulate plant cell expansion. Science 343, 408–411 (2014).
pubmed: 24458638 pmcid: 4672726 doi: 10.1126/science.1244454
Du, M., Spalding, E. P. & Gray, W. M. Rapid auxin-mediated cell expansion. Annu. Rev. Plant. Biol. 71, 379–402 (2020).
pubmed: 32131604 pmcid: 7733314 doi: 10.1146/annurev-arplant-073019-025907
Cosgrove, D. J. Catalysts of plant cell wall loosening. F1000Res. 5, F1000 (2016).
pubmed: 26918182 pmcid: 4755413 doi: 10.12688/f1000research.7180.1
Silk, W. K. & Bogeat-Triboulot, M.-B. Deposition rates in growing tissue: implications for physiology, molecular biology, and response to environmental variation. Plant. Soil. 374, 1–17 (2014).
doi: 10.1007/s11104-013-1726-9
Haruta, M. & Sussman, M. R. Ligand receptor-mediated regulation of growth in plants. Curr. Top. Dev. Biol. 123, 331–363 (2017).
pubmed: 28236971 doi: 10.1016/bs.ctdb.2016.11.007
Pérez-Henríquez, P. & Yang, Z. Extranuclear auxin signaling: a new insight into auxin’s versatility. N. Phytol. 237, 1115–1121 (2023).
doi: 10.1111/nph.18602
True, J. H. & Shaw, S. L. Exogenous auxin induces transverse microtubule arrays through transport inhibitor response1/auxin signaling f-box receptors 1. Plant. Physiol. 182, 892–907 (2020).
pubmed: 31767691 doi: 10.1104/pp.19.00928
Adamowski, M., Li, L. & Friml, J. Reorientation of cortical microtubule arrays in the hypocotyl of Arabidopsis thaliana is induced by the cell growth process and independent of auxin signaling. Int. J. Mol. Sci. 20, 3337 (2019).
pubmed: 31284661 pmcid: 6651120 doi: 10.3390/ijms20133337
Baskin, T. I. Auxin inhibits expansion rate independently of cortical microtubules. Trends Plant. Sci. 20, 471–472 (2015).
pubmed: 26044741 doi: 10.1016/j.tplants.2015.05.008
Xie, Y. et al. FERONIA receptor kinase integrates with hormone signaling to regulate plant growth, development, and responses to environmental stimuli. Int. J. Mol. Sci. 23, 3730 (2022).
pubmed: 35409090 pmcid: 8998941 doi: 10.3390/ijms23073730
Hofte, H. The yin and yang of cell wall integrity control: brassinosteroid and FERONIA signaling. Plant. Cell Physiol. 56, 224–231 (2015).
pubmed: 25481004 doi: 10.1093/pcp/pcu182
Malivert, A. & Hamant, O. Why is FERONIA pleiotropic? Nat. Plants 9, 1018–1025 (2023).
pubmed: 37336971 doi: 10.1038/s41477-023-01434-9
Evered, C., Majevadia, B. & Thompson, D. S. Cell wall water content has a direct effect on extensibility in growing hypocotyls of sunflower (Helianthus annuus L.). J. Exp. Bot. 58, 3361–3371 (2007).
pubmed: 17898424 doi: 10.1093/jxb/erm183
Edelmann, H. G. Water potential modulates extensibility of rye coleoptile cell-walls. Botanica Acta 108, 374–380 (1995).
doi: 10.1111/j.1438-8677.1995.tb00508.x
Muhammad Aslam, M. et al. Mechanisms of abscisic acid-mediated drought stress responses in plants. Int. J. Mol. Sci. 23, 1084 (2022).
pubmed: 35163008 pmcid: 8835272 doi: 10.3390/ijms23031084
Doblin, M. S., Pettolino, F. & Bacic, A. Plant cell walls: the skeleton of the plant world. Funct. Plant. Biol. 37, 357–381 (2010).
doi: 10.1071/FP09279
Lampugnani, E. R., Khan, G. A., Somssich, M. & Persson, S. Building a plant cell wall at a glance. J. Cell Sci. 131, jcs207373 (2018).
pubmed: 29378834 doi: 10.1242/jcs.207373
Carpita, N. C. & Gibeaut, D. M. Structural models of primary cell walls in flowering plants: consistency of molecular structure with the physical properties of the walls during growth. Plant. J. 3, 1–30 (1993).
pubmed: 8401598 doi: 10.1111/j.1365-313X.1993.tb00007.x
Cosgrove, D. J. Building an extensible cell wall. Plant. Physiol. 189, 1246–1277 (2022).
pubmed: 35460252 pmcid: 9237729 doi: 10.1093/plphys/kiac184
Park, Y. B. & Cosgrove, D. J. A revised architecture of primary cell walls based on biomechanical changes induced by substrate-specific endoglucanases. Plant. Physiol. 158, 1933–1943 (2012).
pubmed: 22362871 pmcid: 3320196 doi: 10.1104/pp.111.192880
Hayashi, T. Xyloglucans in the primary cell wall. Annu. Rev. Plant. Phys. Plant. Mol. Bio 40, 139–168 (1989).
doi: 10.1146/annurev.pp.40.060189.001035
Fry, S. C. Cellulases, hemicelluloses and auxin-stimulated growth: a possible relationship. Physiologia Plant. 75, 532–536 (1989).
doi: 10.1111/j.1399-3054.1989.tb05620.x
McCann, M. C. & Roberts, K. in Cytoskeletal Basis of Plant Growth and Form (ed. Lloyd C.) 109–129 (Academic Press, 1991).
Probine, M. C. & Barber, N. F. The structure and plastic properties of the cell wall of Nitella in relation to extension growth. Aust. J. Biol. Sci. 19, 439–457 (1966).
doi: 10.1071/BI9660439
Oliveri, H., Traas, J., Godin, C. & Ali, O. Regulation of plant cell wall stiffness by mechanical stress: a mesoscale physical model. J. Math. Biol. 78, 625–653 (2019).
pubmed: 30209574 doi: 10.1007/s00285-018-1286-y
Cavalier, D. M. et al. Disrupting two Arabidopsis thaliana xylosyltransferase genes results in plants deficient in xyloglucan, a major primary cell wall component. Plant. Cell 20, 1519–1537 (2008). This study showed that genetic deletion of xyloglucan had remarkably little effect on plant development. This undermined the concept that cellulose is mechanically linked by xyloglucan, which was the dominant wall model at that time.
pubmed: 18544630 pmcid: 2483363 doi: 10.1105/tpc.108.059873
Park, Y. B. & Cosgrove, D. J. Changes in cell wall biomechanical properties in the xyloglucan-deficient xxt1/xxt2 mutant of Arabidopsis. Plant. Physiol. 158, 465–475 (2012).
pubmed: 22108526 doi: 10.1104/pp.111.189779
Aryal, B. et al. Interplay between cell wall and auxin mediates the control of differential cell elongation during apical hook development. Curr. Biol. 30, 1733–1739.e3 (2020).
pubmed: 32197084 doi: 10.1016/j.cub.2020.02.055
Kim, S.-J. et al. The synthesis of xyloglucan, an abundant plant cell wall polysaccharide, requires CSLC function. Proc. Natl Acad. Sci. USA 117, 20316–20324 (2020).
pubmed: 32737163 pmcid: 7443942 doi: 10.1073/pnas.2007245117
Zhang, T., Tang, H., Vavylonis, D. & Cosgrove, D. J. Disentangling loosening from softening: insights into primary cell wall structure. Plant. J. 100, 1101–1117 (2019).
pubmed: 31469935 doi: 10.1111/tpj.14519
Wei, W. et al. Synergism between cucumber alpha-expansin, fungal endoglucanase and pectin lyase. J. Plant. Physiol. 167, 1204–1210 (2010).
pubmed: 20478643 doi: 10.1016/j.jplph.2010.03.017
Zhang, T., Vavylonis, D., Durachko, D. M. & Cosgrove, D. J. Nanoscale movements of cellulose microfibrils in primary cell walls. Nat. Plants 3, 17056 (2017).
pubmed: 28452988 pmcid: 5478883 doi: 10.1038/nplants.2017.56
Burton, R. A., Gidley, M. J. & Fincher, G. B. Heterogeneity in the chemistry, structure and function of plant cell walls. Nat. Chem. Biol. 6, 724–732 (2010).
pubmed: 20852610 doi: 10.1038/nchembio.439
van de Meene, A. M. L., Doblin, M. S. & Bacic, A. The plant secretory pathway seen through the lens of the cell wall. Protoplasma 254, 75–94 (2017).
pubmed: 26993347 doi: 10.1007/s00709-016-0952-4
San Clemente, H., Kolkas, H., Canut, H. & Jamet, E. Plant cell wall proteomes: the core of conserved protein families and the case of non-canonical proteins. Int. J. Mol. Sci. 23, 4273 (2022).
pubmed: 35457091 pmcid: 9029284 doi: 10.3390/ijms23084273
The Arabidopsis Genome Initiative Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature 408, 796–815 (2000).
doi: 10.1038/35048692
Carpita, N. C. Update on mechanisms of plant cell wall biosynthesis: how plants make cellulose and other (1->4)-beta-D-glycans. Plant. Physiol. 155, 171–184 (2011).
pubmed: 21051553 doi: 10.1104/pp.110.163360
Wilson, T. H., Kumar, M. & Turner, S. R. The molecular basis of plant cellulose synthase complex organisation and assembly. Biochem. Soc. Trans. 49, 379–391 (2021).
pubmed: 33616627 doi: 10.1042/BST20200697
Pedersen, G. B., Blaschek, L., Frandsen, K. E. H., Noack, L. C. & Persson, S. Cellulose synthesis in land plants. Mol. Plant. 16, 206–231 (2023). After a concise summary of cell wall structure, this review dives deep into the machinery of cellulose synthesis, as far as we know it today.
pubmed: 36564945 doi: 10.1016/j.molp.2022.12.015
Purushotham, P., Ho, R. & Zimmer, J. Architecture of a catalytically active homotrimeric plant cellulose synthase complex. Science 369, 1089–1094 (2020).
pubmed: 32646917 doi: 10.1126/science.abb2978
Jarvis, M. C. Structure of native cellulose microfibrils, the starting point for nanocellulose manufacture. Philos. Trans. Ser. A Math. Phys. Eng. Sci. 376, 20170045 (2018).
Speicher, T. L., Li, P. Z. & Wallace, I. S. Phosphoregulation of the plant cellulose synthase complex and cellulose synthase-like proteins. Plants 7, 52 (2018).
pubmed: 29966291 pmcid: 6161211 doi: 10.3390/plants7030052
Paredez, A. R., Somerville, C. R. & Ehrhardt, D. W. Visualization of cellulose synthase demonstrates functional association with microtubules. Science 312, 1491–1495 (2006). A classic in the cellulose field, demonstrating tagging of cellulose synthase with fluorescent protein to monitor the movement of CSC along the plasma membrane.
pubmed: 16627697 doi: 10.1126/science.1126551
Duncombe, S. G., Chethan, S. G. & Anderson, C. T. Super-resolution imaging illuminates new dynamic behaviors of cellulose synthase. Plant. Cell 34, 273–286 (2022).
pubmed: 34524465 doi: 10.1093/plcell/koab227
Gu, Y. & Rasmussen, C. G. Cell biology of primary cell wall synthesis in plants. Plant. Cell 34, 103–128 (2021).
pmcid: 8774047 doi: 10.1093/plcell/koab249
Li, S., Lei, L., Somerville, C. R. & Gu, Y. Cellulose synthase interactive protein 1 (CSI1) links microtubules and cellulose synthase complexes. Proc. Natl Acad. Sci. USA 109, 185–190 (2012).
pubmed: 22190487 doi: 10.1073/pnas.1118560109
Chan, J. & Coen, E. Interaction between autonomous and microtubule guidance systems controls cellulose synthase trajectories. Curr. Biol. 30, 941–947 (2020).
pubmed: 32037093 doi: 10.1016/j.cub.2019.12.066
Schneider, R. et al. Two complementary mechanisms underpin cell wall patterning during xylem vessel development. Plant. Cell 29, 2433–2449 (2017).
pubmed: 28947492 pmcid: 5774576 doi: 10.1105/tpc.17.00309
Vain, T. et al. The cellulase KORRIGAN is part of the cellulose synthase complex. Plant. Physiol. 165, 1521–1532 (2014).
pubmed: 24948829 pmcid: 4119035 doi: 10.1104/pp.114.241216
Haigler, C. H. & Roberts, A. W. Structure/function relationships in the rosette cellulose synthesis complex illuminated by an evolutionary perspective. Cellulose 26, 227–247 (2019).
doi: 10.1007/s10570-018-2157-9
Nixon, B. T. et al. Comparative structural and computational analysis supports eighteen cellulose synthases in the plant cellulose synthesis complex. Sci. Rep. 6, 28696 (2016). This study argues for an 18-chain microfibril based on the structure of the cellulose synthase complex.
pubmed: 27345599 pmcid: 4921827 doi: 10.1038/srep28696
Song, B., Zhao, S., Shen, W., Collings, C. & Ding, S. Y. Direct measurement of plant cellulose microfibril and bundles in native cell walls. Front. Plant. Sci. 11, 479 (2020).
pubmed: 32391038 pmcid: 7193091 doi: 10.3389/fpls.2020.00479
Del Mundo, J. T. et al. Grazing-incidence diffraction reveals cellulose and pectin organization in hydrated plant primary cell wall. Sci. Rep. 13, 5421 (2023).
pubmed: 37012389 pmcid: 10070456 doi: 10.1038/s41598-023-32505-8
Paajanen, A., Zitting, A., Rautkari, L., Ketoja, J. A. & Penttilä, P. A. Nanoscale mechanism of moisture-induced swelling in wood microfibril bundles. Nano Lett. 22, 5143–5150 (2022).
pubmed: 35767745 pmcid: 9284609 doi: 10.1021/acs.nanolett.2c00822
Wang, T. & Hong, M. Solid-state NMR investigations of cellulose structure and interactions with matrix polysaccharides in plant primary cell walls. J. Exp. Bot. 67, 503–514 (2016).
pubmed: 26355148 doi: 10.1093/jxb/erv416
Tai, H.-C. et al. Wood cellulose microfibrils have a 24-chain core–shell nanostructure in seed plants. Nat. Plants 9, 1154–1168 (2023).
pubmed: 37349550 doi: 10.1038/s41477-023-01430-z
Zhang, T., Zheng, Y. & Cosgrove, D. J. Spatial organization of cellulose microfibrils and matrix polysaccharides in primary plant cell walls as imaged by multichannel atomic force microscopy. Plant. J. 85, 179–192 (2016).
pubmed: 26676644 doi: 10.1111/tpj.13102
Langan, P. et al. Common processes drive the thermochemical pretreatment of lignocellulosic biomass. Green. Chem. 16, 63–68 (2014).
doi: 10.1039/C3GC41962B
Li, T. et al. Developing fibrillated cellulose as a sustainable technological material. Nature 590, 47–56 (2021).
pubmed: 33536649 doi: 10.1038/s41586-020-03167-7
Diotallevi, F. & Mulder, B. The cellulose synthase complex: a polymerization driven supramolecular motor. Biophys. J. 92, 2666–2673 (2007).
pubmed: 17237206 pmcid: 1831695 doi: 10.1529/biophysj.106.099473
Gutierrez, R., Lindeboom, J. J., Paredez, A. R., Emons, A. M. & Ehrhardt, D. W. Arabidopsis cortical microtubules position cellulose synthase delivery to the plasma membrane and interact with cellulose synthase trafficking compartments. Nat. Cell Biol. 11, 797–806 (2009).
pubmed: 19525940 doi: 10.1038/ncb1886
Crowell, E. F. et al. Pausing of Golgi bodies on microtubules regulates secretion of cellulose synthase complexes in Arabidopsis. Plant. Cell 21, 1141–1154 (2009).
pubmed: 19376932 pmcid: 2685615 doi: 10.1105/tpc.108.065334
Zhu, Y. & McFarlane, H. E. Regulation of cellulose synthesis via exocytosis and endocytosis. Curr. Opin. Plant. Biol. 69, 102273 (2022).
pubmed: 35987011 doi: 10.1016/j.pbi.2022.102273
Sampathkumar, A. et al. Patterning and lifetime of plasma membrane-localized cellulose synthase is dependent on actin organization in Arabidopsis interphase cells. Plant. Physiol. 162, 675–688 (2013).
pubmed: 23606596 pmcid: 3668062 doi: 10.1104/pp.113.215277
Ghassemi, N. et al. Solid-state NMR investigations of extracellular matrixes and cell walls of algae, bacteria, fungi, and plants. Chem. Rev. 122, 10036–10086 (2022).
pubmed: 34878762 doi: 10.1021/acs.chemrev.1c00669
Yang, J. et al. Biochemical and genetic analysis identify CSLD3 as a beta-1,4-glucan synthase that functions during plant cell wall synthesis. Plant. Cell 32, 1749–1767 (2020).
pubmed: 32169960 pmcid: 7203914 doi: 10.1105/tpc.19.00637
Yang, J. et al. Functional relations of CSLD2, CSLD3, and CSLD5 proteins during cell wall synthesis in Arabidopsis. Preprint at bioRxiv https://doi.org/10.1101/2023.04.25.538313 (2023).
Nicolas, W. J. et al. Cryo-electron tomography of the onion cell wall shows bimodally oriented cellulose fibers and reticulated homogalacturonan networks. Curr. Biol. 32, 2375–2389.e6 (2022).
pubmed: 35508170 pmcid: 9240970 doi: 10.1016/j.cub.2022.04.024
Chan, J., Calder, G., Fox, S. & Lloyd, C. Cortical microtubule arrays undergo rotary movements in Arabidopsis hypocotyl epidermal cells. Nat. Cell Biol. 9, 171–175 (2007).
pubmed: 17220881 doi: 10.1038/ncb1533
Jarvis, M. C. Hydrogen bonding and other non-covalent interactions at the surfaces of cellulose microfibrils. Cellulose 30, 667–687 (2023).
doi: 10.1007/s10570-022-04954-3
Zhang, Q., Brumer, H., Agren, H. & Tu, Y. The adsorption of xyloglucan on cellulose: effects of explicit water and side chain variation. Carbohydr. Res. 346, 2595–2602 (2011).
pubmed: 21974911 doi: 10.1016/j.carres.2011.09.007
Zhao, Z., Crespi, V. H., Kubicki, J. D., Cosgrove, D. J. & Zhong, L. H. Molecular dynamics simulation study of xyloglucan adsorption on cellulose surfaces: effects of surface hydrophobicity and side-chain variation. Cellulose 21, 1025–1039 (2014).
doi: 10.1007/s10570-013-0041-1
Kishani, S., Benselfelt, T., Wagberg, L. & Wohlert, J. Entropy drives the adsorption of xyloglucan to cellulose surfaces — a molecular dynamics study. J. Colloid Interface Sci. 588, 485–493 (2021).
pubmed: 33429345 doi: 10.1016/j.jcis.2020.12.113
Cosgrove, D. J. Nanoscale structure, mechanics and growth of epidermal cell walls. Curr. Opin. Plant. Biol. 46, 77–86 (2018).
pubmed: 30142487 doi: 10.1016/j.pbi.2018.07.016
Scheller, H. V. & Ulvskov, P. Hemicelluloses. Annu. Rev. Plant. Biol. 61, 263–289 (2010).
pubmed: 20192742 doi: 10.1146/annurev-arplant-042809-112315
Hoffmann, N., King, S., Samuels, A. L. & McFarlane, H. E. Subcellular coordination of plant cell wall synthesis. Dev. Cell 56, 933–948 (2021). A detailed overview of the cell biology of cell wall synthesis.
pubmed: 33761322 doi: 10.1016/j.devcel.2021.03.004
Wang, P., Chen, X., Goldbeck, C., Chung, E. & Kang, B. H. A distinct class of vesicles derived from the trans-Golgi mediates secretion of xylogalacturonan in the root border cell. Plant. J. 92, 596–610 (2017).
pubmed: 28865155 doi: 10.1111/tpj.13704
Yu, L. et al. Eudicot primary cell wall glucomannan is related in synthesis, structure, and function to xyloglucan. Plant. Cell 34, 4600–4622 (2022).
pubmed: 35929080 pmcid: 9614514 doi: 10.1093/plcell/koac238
Park, Y. B. & Cosgrove, D. J. Xyloglucan and its interactions with other components of the growing cell wall. Plant. Cell Physiol. 56, 180–194 (2015).
pubmed: 25613914 doi: 10.1093/pcp/pcu204
Schultink, A., Liu, L., Zhu, L. & Pauly, M. Structural diversity and function of xyloglucan sidechain substituents. Plants 3, 526–542 (2014).
pubmed: 27135518 pmcid: 4844278 doi: 10.3390/plants3040526
Pauly, M. & Keegstra, K. Biosynthesis of the plant cell wall matrix polysaccharide xyloglucan. Annu. Rev. Plant. Biol. 67, 235–259 (2016).
pubmed: 26927904 doi: 10.1146/annurev-arplant-043015-112222
Chen, M., Cathala, B. & Lahaye, M. Adsorption of apple xyloglucan on cellulose nanofiber depends on molecular weight, concentration and building blocks. Carbohydr. Polym. 296, 119994 (2022).
pubmed: 36088015 doi: 10.1016/j.carbpol.2022.119994
Velasquez, S. M. et al. Xyloglucan remodeling defines auxin-dependent differential tissue expansion in plants. Int. J. Mol. Sci. 22, 9222 (2021).
pubmed: 34502129 pmcid: 8430841 doi: 10.3390/ijms22179222
Schultink, A., Cheng, K., Park, Y. B., Cosgrove, D. J. & Pauly, M. The identification of two arabinosyltransferases from tomato reveals functional equivalency of xyloglucan side chain substituents. Plant. Physiol. 163, 86–94 (2013).
pubmed: 23893172 pmcid: 3762667 doi: 10.1104/pp.113.221788
Jonsson, K., Hamant, O. & Bhalerao, R. P. Plant cell walls as mechanical signaling hubs for morphogenesis. Curr. Biol. 32, R334–R340 (2022).
pubmed: 35413265 doi: 10.1016/j.cub.2022.02.036
Sowinski, E. E. et al. Lack of xyloglucan in the cell walls of the Arabidopsis xxt1/xxt2 mutant results in specific increases in homogalacturonan and glucomannan. Plant. J. 110, 212–227 (2022).
pubmed: 35041247 doi: 10.1111/tpj.15666
Xiao, C., Zhang, T., Zheng, Y., Cosgrove, D. J. & Anderson, C. T. Xyloglucan deficiency disrupts microtubule stability and cellulose biosynthesis in Arabidopsis, altering cell growth and morphogenesis. Plant. Physiol. 170, 234–249 (2016).
pubmed: 26527657 doi: 10.1104/pp.15.01395
Xiang, M. et al. Xyloglucan galactosylation is essential for proper cell wall assembly that facilitates stabilization of the actin cytoskeleton and the endomembrane system. J. Exp. Bot. 74, 5104–5123 (2023).
pubmed: 37386914 doi: 10.1093/jxb/erad237
Kong, Y. et al. Galactose-depleted xyloglucan is dysfunctional and leads to dwarfism in Arabidopsis. Plant. Physiol. 167, 1296–1306 (2015).
pubmed: 25673778 pmcid: 4378170 doi: 10.1104/pp.114.255943
Pauly, M., Albersheim, P., Darvill, A. & York, W. S. Molecular domains of the cellulose/xyloglucan network in the cell walls of higher plants. Plant. J. 20, 629–639 (1999).
pubmed: 10652135 doi: 10.1046/j.1365-313X.1999.00630.x
Zheng, Y., Wang, X., Chen, Y., Wagner, E. & Cosgrove, D. J. Xyloglucan in the primary cell wall: assessment by FESEM, selective enzyme digestions and nanogold affinity tags. Plant. J. 93, 211–226 (2018).
pubmed: 29160933 doi: 10.1111/tpj.13778
Tryfona, T. et al. Grass xylan structural variation suggests functional specialization and distinctive interaction with cellulose and lignin. Plant. J. 113, 1004–1020 (2023).
pubmed: 36602010 doi: 10.1111/tpj.16096
Wang, T., Chen, Y., Tabuchi, A., Cosgrove, D. J. & Hong, M. The target of beta-expansin EXPB1 in maize cell walls from binding and solid-state NMR studies. Plant. Physiol. 172, 2107–2119 (2016).
pubmed: 27729469 pmcid: 5129719 doi: 10.1104/pp.16.01311
Duan, P. et al. Xylan structure and dynamics in native Brachypodium grass cell walls investigated by solid-state NMR spectroscopy. ACS Omega 6, 15460–15471 (2021).
pubmed: 34151124 pmcid: 8210444 doi: 10.1021/acsomega.1c01978
Carpita, N. C. Hemicellulosic polymers of cell walls of zea coleoptiles. Plant. Physiol. 72, 515–521 (1983).
pubmed: 16663034 pmcid: 1066265 doi: 10.1104/pp.72.2.515
Ropartz, D. & Ralet, M.-C. in Pectin: Technological and Physiological Properties (ed. Kontogiorgos V.) Ch. 2, 17-36 (Springer Nature, 2020).
Du, J., Anderson, C. T. & Xiao, C. Dynamics of pectic homogalacturonan in cellular morphogenesis and adhesion, wall integrity sensing and plant development. Nat. Plants 8, 332–340 (2022).
pubmed: 35411046 doi: 10.1038/s41477-022-01120-2
Atmodjo, M. A., Hao, Z. & Mohnen, D. Evolving views of pectin biosynthesis. Annu. Rev. Plant. Biol. 64, 747–779 (2013).
pubmed: 23451775 doi: 10.1146/annurev-arplant-042811-105534
Caffall, K. H. & Mohnen, D. The structure, function, and biosynthesis of plant cell wall pectic polysaccharides. Carbohydr. Res. 344, 1879–1900 (2009).
pubmed: 19616198 doi: 10.1016/j.carres.2009.05.021
Jarvis, M. C. Structure and properties of pectin gels in plant-cell walls. Plant. Cell Environ. 7, 153–164 (1984).
doi: 10.1111/1365-3040.ep11614586
Jarvis, M. C. Control of thickness of collenchyma cell walls by pectins. Planta 187, 218–220 (1992).
pubmed: 24178046 doi: 10.1007/BF00201941
Thimm, J. C., Burritt, D. J., Ducker, W. A. & Melton, L. D. Pectins influence microfibril aggregation in celery cell walls: an atomic force microscopy study. J. Struct. Biol. 168, 337–344 (2009).
pubmed: 19567269 doi: 10.1016/j.jsb.2009.06.017
Radja, A., Horsley, E. M., Lavrentovich, M. O. & Sweeney, A. M. Pollen cell wall patterns form from modulated phases. Cell 176, 856–868.e810 (2019).
pubmed: 30735635 doi: 10.1016/j.cell.2019.01.014
Radja, A. Pollen wall patterns as a model for biological self-assembly. J. Exp. Zool. B Mol. Dev. Evol. 336, 629–641 (2021).
pubmed: 32991047 doi: 10.1002/jez.b.23005
Palacio-Lopez, K. et al. Experimental manipulation of pectin architecture in the cell wall of the unicellular charophyte, Penium margaritaceum. Front. Plant. Sci. 11, 1032 (2020).
pubmed: 32733522 pmcid: 7360812 doi: 10.3389/fpls.2020.01032
Temple, H. et al. Golgi-localized putative S-adenosyl methionine transporters required for plant cell wall polysaccharide methylation. Nat. Plants 8, 656–669 (2022).
pubmed: 35681018 doi: 10.1038/s41477-022-01156-4
John, J., Ray, D., Aswal, V. K., Deshpande, A. P. & Varughese, S. Dissipation and strain-stiffening behavior of pectin–Ca gels under LAOS. Soft Matter 15, 6852–6866 (2019).
pubmed: 31410439 doi: 10.1039/C9SM00709A
John, J., Ray, D., Aswal, V. K., Deshpande, A. P. & Varughese, S. Pectin self-assembly and its disruption by water: insights into plant cell wall mechanics. Phys. Chem. Chem. Phys. 24, 22691–22698 (2022).
pubmed: 36106571 doi: 10.1039/D2CP01479C
Willats, W. G. et al. Modulation of the degree and pattern of methyl-esterification of pectic homogalacturonan in plant cell walls. Implications for pectin methyl esterase action, matrix properties, and cell adhesion. J. Biol. Chem. 276, 19404–19413 (2001).
pubmed: 11278866 doi: 10.1074/jbc.M011242200
Williams, M. A. K. et al. Polysaccharide structures in the outer mucilage of arabidopsis seeds visualized by AFM. Biomacromolecules 21, 1450–1459 (2020).
pubmed: 32058700 doi: 10.1021/acs.biomac.9b01756
Tan, L. et al. Most of the rhamnogalacturonan-I from cultured Arabidopsis cell walls is covalently linked to arabinogalactan-protein. Carbohydr. Polym. 301, 120340 (2023).
pubmed: 36446508 doi: 10.1016/j.carbpol.2022.120340
Yang, H. et al. Rhamnogalacturonan-I is a determinant of cell-cell adhesion in poplar wood. Plant. Biotechnol. J. 18, 1027–1040 (2020).
pubmed: 31584248 doi: 10.1111/pbi.13271
Saez-Aguayo, S. & Largo-Gosens, A. Rhamnogalacturonan-I forms mucilage: behind its simplicity, a cutting-edge organization. J. Exp. Bot. 73, 3299–3303 (2022).
pubmed: 36305092 pmcid: 9162176 doi: 10.1093/jxb/erac094
Saffer, A. M. et al. Cellulose assembles into helical bundles of uniform handedness in cell walls with abnormal pectin composition. Plant. J. 116, 855–870 (2023).
pubmed: 37548081 doi: 10.1111/tpj.16414
Begum, R. A., Messenger, D. J. & Fry, S. C. Making and breaking of boron bridges in the pectic domain rhamnogalacturonan-II at apoplastic pH in vivo and in vitro. Plant. J. 113, 1310–1329 (2023).
pubmed: 36658763 doi: 10.1111/tpj.16112
Lerouge, P., Carlier, M., Mollet, J.-C. & Lehner, A. The cell wall pectic rhamnogalacturonan II, an enigma in plant glycobiology. Carbohydr. Chem. 45, 553–571 (2021).
doi: 10.1039/9781839164538-00553
Fry, S. C. Polysaccharide-modifying enzymes in the plant cell wall. Ann. Rev. Plant Physiol. 46, 497–520 (1995).
doi: 10.1146/annurev.pp.46.060195.002433
Wolf, S. Cell wall signaling in plant development and defense. Annu. Rev. Plant. Biol. 73, 323–353 (2022). This review details the variety of cell wall sensors and their signalling pathways.
pubmed: 35167757 doi: 10.1146/annurev-arplant-102820-095312
Novakovic, L., Guo, T. T., Bacic, A., Sampathkumar, A. & Johnson, K. L. Hitting the wall sensing and signaling pathways involved in plant cell wall remodeling in response to abiotic stress. Plants-Basel 7, 89 (2018).
pubmed: 30360552 pmcid: 6313904 doi: 10.3390/plants7040089
Lin, W. et al. Arabidopsis pavement cell morphogenesis requires FERONIA binding to pectin for activation of ROP GTPase signaling. Curr. Biol. 32, 497–507.e4 (2022).
pubmed: 34875229 doi: 10.1016/j.cub.2021.11.030
Tang, W. et al. Mechano-transduction via the pectin-FERONIA complex activates ROP6 GTPase signaling in Arabidopsis pavement cell morphogenesis. Curr. Biol. 32, 508–517.e3 (2022).
pubmed: 34875231 doi: 10.1016/j.cub.2021.11.031
Duan, Q. et al. FERONIA controls pectin- and nitric oxide-mediated male-female interaction. Nature 579, 561–566 (2020).
pubmed: 32214247 doi: 10.1038/s41586-020-2106-2
Dunser, K. et al. Extracellular matrix sensing by FERONIA and leucine-rich repeat extensins controls vacuolar expansion during cellular elongation in Arabidopsis thaliana. EMBO J. 38, e100353 (2019).
pubmed: 30850388 pmcid: 6443208 doi: 10.15252/embj.2018100353
Johnson, K. L. et al. Insights into the evolution of hydroxyproline-rich glycoproteins from 1000 plant transcriptomes. Plant. Physiol. 174, 904–921 (2017).
pubmed: 28446636 pmcid: 5462033 doi: 10.1104/pp.17.00295
Borassi, C. et al. An update on cell surface proteins containing extensin-motifs. J. Exp. Bot. 67, 477–487 (2016).
pubmed: 26475923 doi: 10.1093/jxb/erv455
Moussu, S. & Ingram, G. The EXTENSIN enigma. Cell Surf. 9, 100094 (2023).
pubmed: 36659959 pmcid: 9842689 doi: 10.1016/j.tcsw.2023.100094
Ma, Y. & Johnson, K. A. Arabinogalactan-proteins. WikiJournal Sci. 4, 1 (2021).
doi: 10.15347/WJS/2021.002
Shafee, T., Bacic, A. & Johnson, K. Evolution of sequence-diverse disordered regions in a protein family: order within the chaos. Mol. Biol. Evol. 37, 2155–2172 (2020).
pubmed: 32359163 doi: 10.1093/molbev/msaa096
Cannon, M. C. et al. Self-assembly of the plant cell wall requires an extensin scaffold. Proc. Natl Acad. Sci. USA 105, 2226–2231 (2008).
pubmed: 18256186 pmcid: 2538902 doi: 10.1073/pnas.0711980105
Sede, A. R. et al. Arabidopsis pollen extensins LRX are required for cell wall integrity during pollen tube growth. FEBS Lett. 592, 233–243 (2018).
pubmed: 29265366 doi: 10.1002/1873-3468.12947
Marzol, E. et al. Filling the gaps to solve the extensin puzzle. Mol. Plant. 11, 645–658 (2018).
pubmed: 29530817 doi: 10.1016/j.molp.2018.03.003
Doll, N. M., Berenguer, E., Truskina, J. & Ingram, G. AtEXT3 is not essential for early embryogenesis or plant viability in Arabidopsis. New Phytol. 236, 1629–1633 (2022).
pubmed: 36052714 pmcid: 9826179 doi: 10.1111/nph.18452
Hromadova, D., Soukup, A. & Tylova, E. Arabinogalactan proteins in plant roots — an update on possible functions. Front. Plant Sci. 12, 674010 (2021).
pubmed: 34079573 pmcid: 8165308 doi: 10.3389/fpls.2021.674010
Lopez-Hernandez, F. et al. Calcium binding by arabinogalactan polysaccharides is important for normal plant development. Plant. Cell 32, 3346–3369 (2020).
pubmed: 32769130 pmcid: 7534474 doi: 10.1105/tpc.20.00027
Silva, J., Ferraz, R., Dupree, P., Showalter, A. M. & Coimbra, S. Three decades of advances in arabinogalactan-protein biosynthesis. Front. Plant Sci. 11, 610377 (2020).
pubmed: 33384708 pmcid: 7769824 doi: 10.3389/fpls.2020.610377
Chen, P., Nishiyama, Y. & Wohlert, J. Quantifying the influence of dispersion interactions on the elastic properties of crystalline cellulose. Cellulose 28, 10777–10786 (2021).
doi: 10.1007/s10570-021-04210-0
Wohlert, M. et al. Cellulose and the role of hydrogen bonds: not in charge of everything. Cellulose 29, 1–23 (2021).
doi: 10.1007/s10570-021-04325-4
Glasser, W. G. et al. About the structure of cellulose: debating the Lindman hypothesis. Cellulose 19, 589–598 (2012).
doi: 10.1007/s10570-012-9691-7
Williams, M. A. K. in Pectin: Technological and Physiological Properties (ed. Kontogiorgos V.) 125–148 (Springer International Publishing, 2020).
Pieczywek, P. M., Cieśla, J., Płaziński, W. & Zdunek, A. Aggregation and weak gel formation by pectic polysaccharide homogalacturonan. Carbohydr. Polym. 256, 117566 (2021).
pubmed: 33483067 doi: 10.1016/j.carbpol.2020.117566
Talbott, L. D. & Ray, P. M. Molecular size and separability features of pea cell wall polysaccharides. Implications for models of primary wall structure. Plant. Physiol. 92, 357–368 (1992).
doi: 10.1104/pp.98.1.357
Broxterman, S. E. & Schols, H. A. Characterisation of pectin-xylan complexes in tomato primary plant cell walls. Carbohydr. Polym. 197, 269–276 (2018).
pubmed: 30007613 doi: 10.1016/j.carbpol.2018.06.003
Cornuault, V., Pose, S. & Knox, J. P. Disentangling pectic homogalacturonan and rhamnogalacturonan-I polysaccharides: evidence for sub-populations in fruit parenchyma systems. Food Chem. 246, 275–285 (2018).
pubmed: 29291850 pmcid: 5770856 doi: 10.1016/j.foodchem.2017.11.025
Moneo-Sánchez, M. et al. β-(1,4)-Galactan remodelling in Arabidopsis cell walls affects the xyloglucan structure during elongation. Planta 249, 351–362 (2019).
pubmed: 30206696 doi: 10.1007/s00425-018-3008-5
Herburger, K. et al. Hetero-trans-beta-glucanase produces cellulose-xyloglucan covalent bonds in the cell walls of structural plant tissues and is stimulated by expansin. Mol. Plant. 13, 1047–1062 (2020).
pubmed: 32376294 doi: 10.1016/j.molp.2020.04.011
Herburger, K. et al. Defining natural factors that stimulate and inhibit cellulose:xyloglucan hetero-transglucosylation. Plant. J. 105, 1549–1565 (2021).
pubmed: 33314395 pmcid: 8611796 doi: 10.1111/tpj.15131
Buanafina, M. M. de O. Feruloylation in grasses: current and future perspectives. Mol. Plant. 2, 861–872 (2009).
doi: 10.1093/mp/ssp067
Chateigner-Boutin, A.-L. & Saulnier, L. in Advances in Botanical Research (ed. Sibout R.) 104, 169–213 (Academic, 2022).
Buanafina, M. M. D. & Morris, P. The impact of cell wall feruloylation on plant growth, responses to environmental stress, plant pathogens and cell wall degradability. Agronomy 12, 1847 (2022).
doi: 10.3390/agronomy12081847
Guhados, G., Wan, W. K. & Hutter, J. L. Measurement of the elastic modulus of single bacterial cellulose fibers using atomic force microscopy. Langmuir 21, 6642–6646 (2005).
pubmed: 15982078 doi: 10.1021/la0504311
Jacobs, C. R., Huang, H. & Kwon, R. Y. Introduction to Cell Mechanics and Mechanobiology (Garland Science, 2013).
Usov, I. et al. Understanding nanocellulose chirality and structure-properties relationship at the single fibril level. Nat. Commun. 6, 7564 (2015).
pubmed: 26108282 doi: 10.1038/ncomms8564
McFarlane, H. E. Open questions in plant cell wall synthesis. J. Exp. Bot. 74, 3425–3448 (2023).
pubmed: 36961357 doi: 10.1093/jxb/erad110
Zhu, X., Li, S., Pan, S., Xin, X. & Gu, Y. CSI1, PATROL1, and exocyst complex cooperate in delivery of cellulose synthase complexes to the plasma membrane. Proc. Natl Acad. Sci. 115, E3578 (2018).
pubmed: 29581258 pmcid: 5899483 doi: 10.1073/pnas.1800182115
Schneider, R., Ehrhardt, D. W., Meyerowitz, E. M. & Sampathkumar, A. Tethering of cellulose synthase to microtubules dampens mechano-induced cytoskeletal organization in Arabidopsis pavement cells. Nat. Plants 8, 1064–1073 (2022).
pubmed: 35982303 pmcid: 9477734 doi: 10.1038/s41477-022-01218-7
Zykwinska, A. W., Ralet, M. C., Garnier, C. D. & Thibault, J. F. Evidence for in vitro binding of pectin side chains to cellulose. Plant. Physiol. 139, 397–407 (2005).
pubmed: 16126855 pmcid: 1203388 doi: 10.1104/pp.105.065912
Lipowczan, M., Borowska-Wykret, D., Natonik-Bialon, S. & Kwiatkowska, D. Growing cell walls show a gradient of elastic strain across their layers. J. Exp. Bot. 69, 4349–4362 (2018).
pubmed: 29945239 pmcid: 6093493 doi: 10.1093/jxb/ery237
Crowell, E. F. et al. Differential regulation of cellulose orientation at the inner and outer face of epidermal cells in the Arabidopsis hypocotyl. Plant. Cell 23, 2592–2605 (2011).
pubmed: 21742992 pmcid: 3226210 doi: 10.1105/tpc.111.087338
Chen, D., Melton, L. D., McGillivray, D. J., Ryan, T. M. & Harris, P. J. Changes in the orientations of cellulose microfibrils during the development of collenchyma cell walls of celery (Apium graveolens L.). Planta 250, 1819–1832 (2019).
pubmed: 31463558 doi: 10.1007/s00425-019-03262-8
Kelly-Bellow, R. et al. Brassinosteroid coordinates cell layer interactions in plants via cell wall and tissue mechanics. Science 380, 1275–1281 (2023).
pubmed: 37347863 doi: 10.1126/science.adf0752
Mollier, C. et al. Spatial consistency of cell growth direction during organ morphogenesis requires CELLULOSE SYNTHASE INTERACTIVE1. Cell Rep. 42, 112689 (2023).
pubmed: 37352099 pmcid: 10391631 doi: 10.1016/j.celrep.2023.112689
McQueen-Mason, S., Durachko, D. M. & Cosgrove, D. J. Two endogenous proteins that induce cell wall extension in plants. Plant. Cell 4, 1425–1433 (1992).
pubmed: 11538167 pmcid: 160229
Cosgrove, D. J. Plant expansins: diversity and interactions with plant cell walls. Curr. Opin. Plant. Biol. 25, 162–172 (2015).
pubmed: 26057089 pmcid: 4532548 doi: 10.1016/j.pbi.2015.05.014
Sampedro, J. & Cosgrove, D. J. The expansin superfamily. Genome Biol. 6, 242 (2005).
pubmed: 16356276 pmcid: 1414085 doi: 10.1186/gb-2005-6-12-242
Marowa, P., Ding, A. M. & Kong, Y. Z. Expansins: roles in plant growth and potential applications in crop improvement. Plant. Cell Rep. 35, 949–965 (2016).
pubmed: 26888755 pmcid: 4833835 doi: 10.1007/s00299-016-1948-4
Rayle, D. L. & Cleland, R. E. The acid growth theory of auxin-induced cell elongation is alive and well. Plant. Physiol. 99, 1271–1274 (1992).
pubmed: 11537886 pmcid: 1080619 doi: 10.1104/pp.99.4.1271
Hocq, L., Pelloux, J. & Lefebvre, V. Connecting homogalacturonan-type pectin remodeling to acid growth. Trends Plant. Sci. 22, 20–29 (2017).
pubmed: 27884541 doi: 10.1016/j.tplants.2016.10.009
Arsuffi, G. & Braybrook, S. A. Acid growth: an ongoing trip. J. Exp. Bot. 69, 137–146 (2018).
pubmed: 29211894 doi: 10.1093/jxb/erx390
Phyo, P., Gu, Y. & Hong, M. Impact of acidic pH on plant cell wall polysaccharide structure and dynamics: insights into the mechanism of acid growth in plants from solid-state NMR. Cellulose 26, 291–304 (2018).
doi: 10.1007/s10570-018-2094-7
Cho, H. T. & Kende, H. Expansins in deepwater rice internodes. Plant. Physiol. 113, 1137–1143 (1997).
pubmed: 9112771 pmcid: 158236 doi: 10.1104/pp.113.4.1137
Cleland, R. E., Cosgrove, D. J. & Tepfer, M. Long-term acid-induced wall extension in an in vitro system. Planta 170, 379–385 (1987).
pubmed: 24232968 doi: 10.1007/BF00395030
Li, L. et al. Cell surface and intracellular auxin signalling for H
pubmed: 34707283 pmcid: 7612300 doi: 10.1038/s41586-021-04037-6
Fendrych, M. et al. Rapid and reversible root growth inhibition by TIR1 auxin signalling. Nat. Plants 4, 453–459 (2018).
pubmed: 29942048 pmcid: 6104345 doi: 10.1038/s41477-018-0190-1
Li, L., Gallei, M. & Friml, J. Bending to auxin: fast acid growth for tropisms. Trends Plant. Sci. 27, 440–449 (2021).
pubmed: 34848141 doi: 10.1016/j.tplants.2021.11.006
Yuan, S., Wu, Y. & Cosgrove, D. J. A fungal endoglucanase with plant cell wall extension activity. Plant. Physiol. 127, 324–333 (2001).
pubmed: 11553760 pmcid: 117988 doi: 10.1104/pp.127.1.324
Wang, C. X., Wang, L., McQueen-Mason, S. J., Pritchard, J. & Thomas, C. R. pH and expansin action on single suspension-cultured tomato (Lycopersicon esculentum) cells. J. Plant. Res. 121, 527–534 (2008).
pubmed: 18615263 doi: 10.1007/s10265-008-0176-6
McQueen-Mason, S. J. & Cosgrove, D. J. Expansin mode of action on cell walls. Analysis of wall hydrolysis, stress relaxation, and binding. Plant. Physiol. 107, 87–100 (1995).
pubmed: 11536663 pmcid: 161171 doi: 10.1104/pp.107.1.87
McQueen-Mason, S. J., Fry, S. C., Durachko, D. M. & Cosgrove, D. J. The relationship between xyloglucan endotransglycosylase and in-vitro cell wall extension in cucumber hypocotyls. Planta 190, 327–331 (1993).
pubmed: 7763661 doi: 10.1007/BF00196961
Cosgrove, D. J. Non-enzymatic action of expansins. J. Biol. Chem. 295, 6782 (2020).
pubmed: 32385096 pmcid: 7212657 doi: 10.1074/jbc.L120.013410
Fleming, A. J., McQueen-Mason, S., Mandel, T. & Kuhlemeier, C. Induction of leaf primordia by the cell wall protein expansin. Science 276, 1415 (1997).
doi: 10.1126/science.276.5317.1415
Link, B. M. & Cosgrove, D. J. Acid-growth response and alpha-expansins in suspension cultures of bright yellow 2 tobacco. Plant. Physiol. 118, 907–916 (1998).
pubmed: 9808735 pmcid: 34801 doi: 10.1104/pp.118.3.907
Goh, H. H., Sloan, J., Malinowski, R. & Fleming, A. Variable expansin expression in Arabidopsis leads to different growth responses. J. Plant. Physiol. 171, 329–339 (2014).
pubmed: 24144490 doi: 10.1016/j.jplph.2013.09.009
Rochange, S. F., Wenzel, C. L. & McQueen-Mason, S. J. Impaired growth in transgenic plants over-expressing an expansin isoform. Plant. Mol. Biol. 46, 581–589 (2001).
pubmed: 11516151 doi: 10.1023/A:1010650217100
Samalova, M. et al. Hormone-regulated expansins: expression, localization, and cell wall biomechanics in Arabidopsis root growth. Plant. Physiol. https://doi.org/10.1093/plphys/kiad228 (2023).
McQueen-Mason, S. & Cosgrove, D. J. Disruption of hydrogen bonding between plant cell wall polymers by proteins that induce wall extension. Proc. Natl Acad. Sci. USA 91, 6574–6578 (1994).
pubmed: 11607483 pmcid: 44245 doi: 10.1073/pnas.91.14.6574
Whitney, S. E. C., Gidley, M. J. & McQueen-Mason, S. J. Probing expansin action using cellulose/hemicellulose composites. Plant. J. 22, 327–334 (2000).
pubmed: 10849349 doi: 10.1046/j.1365-313x.2000.00742.x
Georgelis, N., Tabuchi, A., Nikolaidis, N. & Cosgrove, D. J. Structure-function analysis of the bacterial expansin EXLX1. J. Biol. Chem. 286, 16814–16823 (2011).
pubmed: 21454649 pmcid: 3089525 doi: 10.1074/jbc.M111.225037
Georgelis, N., Yennawar, N. H. & Cosgrove, D. J. Structural basis for entropy-driven cellulose binding by a type-A cellulose-binding module (CBM) and bacterial expansin. Proc. Natl Acad. Sci. USA 109, 14830–14835 (2012).
pubmed: 22927418 pmcid: 3443152 doi: 10.1073/pnas.1213200109
Imai, T. et al. Disturbance of the hydrogen bonding in cellulose by bacterial expansin. Cellulose 30, https://doi.org/10.1007/s10570-023-05402-6 (2023).
Yennawar, N. H., Li, L. C., Dudzinski, D. M., Tabuchi, A. & Cosgrove, D. J. Crystal structure and activities of EXPB1 (Zea m 1), a beta-expansin and group-1 pollen allergen from maize. Proc. Natl Acad. Sci. USA 103, 14664–14671 (2006).
pubmed: 16984999 pmcid: 1595409 doi: 10.1073/pnas.0605979103
Silveira, R. L. & Skaf, M. S. Molecular dynamics of the Bacillus subtilis expansin EXLX1: interaction with substrates and structural basis of the lack of activity of mutants. Phys. Chem. Chem. Phys. 18, 3510–3521 (2016).
pubmed: 26751268 doi: 10.1039/C5CP06674C
Hayashi, T. & Kaida, R. Functions of xyloglucan in plant cells. Mol. Plant. 4, 17–24 (2011).
pubmed: 20943810 doi: 10.1093/mp/ssq063
Nishitani, K. & Vissenberg, K. in The Expanding Cell (eds. Verbelen J.-P. & Vissenberg K. eds) 89–116 (Springer Berlin Heidelberg, 2007).
Behar, H., Graham, S. W. & Brumer, H. Comprehensive cross-genome survey and phylogeny of glycoside hydrolase family 16 members reveals the evolutionary origin of EG16 and XTH proteins in plant lineages. Plant. J. 95, 1114–1128 (2018).
pubmed: 29932263 doi: 10.1111/tpj.14004
Behar, H., Tamura, K., Wagner, E. R., Cosgrove, D. J. & Brumer, H. Conservation of endo-glucanase 16 (EG16) activity across highly divergent plant lineages. Biochem. J. 478, 3063–3078 (2021).
pubmed: 34338284 doi: 10.1042/BCJ20210341
Hrmova, M., Stratilova, B. & Stratilova, E. Broad specific xyloglucan:xyloglucosyl transferases are formidable players in the re-modelling of plant cell wall structures. Int. J. Mol. Sci. 23, 1656 (2022).
pubmed: 35163576 pmcid: 8836008 doi: 10.3390/ijms23031656
Saladie, M., Rose, J. K., Cosgrove, D. J. & Catala, C. Characterization of a new xyloglucan endotransglucosylase/hydrolase (XTH) from ripening tomato fruit and implications for the diverse modes of enzymic action. Plant. J. 47, 282–295 (2006).
pubmed: 16774648 doi: 10.1111/j.1365-313X.2006.02784.x
Maris, A., Suslov, D., Fry, S. C., Verbelen, J. P. & Vissenberg, K. Enzymic characterization of two recombinant xyloglucan endotransglucosylase/hydrolase (XTH) proteins of Arabidopsis and their effect on root growth and cell wall extension. J. Exp. Bot. 60, 3959–3972 (2009).
pubmed: 19635745 doi: 10.1093/jxb/erp229
Van Sandt, V. S. T., Suslov, D., Verbelen, J. P. & Vissenberg, K. Xyloglucan endotransglucosylase activity loosens a plant cell wall. Ann. Bot. 100, 1467–1473 (2007).
pubmed: 17916584 pmcid: 2759230 doi: 10.1093/aob/mcm248
Kaewthai, N. et al. Group III-A XTH genes of Arabidopsis encode predominant xyloglucan endohydrolases that are dispensable for normal growth. Plant. Physiol. 161, 440–454 (2013).
pubmed: 23104861 doi: 10.1104/pp.112.207308
Hara, Y., Yokoyama, R., Osakabe, K., Toki, S. & Nishitani, K. Function of xyloglucan endotransglucosylase/hydrolases in rice. Ann. Bot. 114, 1309–1318 (2014).
pubmed: 24363334 doi: 10.1093/aob/mct292
Niraula, P. M., Zhang, X., Jeremic, D., Lawrence, K. S. & Klink, V. P. Xyloglucan endotransglycosylase/hydrolase increases tightly-bound xyloglucan and chain number but decreases chain length contributing to the defense response that Glycine max has to Heterodera glycines. PLoS ONE 16, e0244305 (2021).
pubmed: 33444331 pmcid: 7808671 doi: 10.1371/journal.pone.0244305
Osato, Y., Yokoyama, R. & Nishitani, K. A principal role for AtXTH18 in Arabidopsis thaliana root growth: a functional analysis using RNAi plants. J. Plant. Res. 119, 153–162 (2006).
pubmed: 16477366 doi: 10.1007/s10265-006-0262-6
Miedes, E. et al. Xyloglucan endotransglucosylase and cell wall extensibility. J. Plant. Physiol. 168, 196–203 (2011).
pubmed: 20828871 doi: 10.1016/j.jplph.2010.06.029
Miedes, E. et al. Xyloglucan endotransglucosylase/hydrolase (XTH) overexpression affects growth and cell wall mechanics in etiolated Arabidopsis hypocotyls. J. Exp. Bot. 64, 2481–2497 (2013).
pubmed: 23585673 doi: 10.1093/jxb/ert107
Kushwah, S. et al. Arabidopsis XTH4 and XTH9 contribute to wood cell expansion and secondary wall formation. Plant Physiol. 182, 1946–1965 (2020).
pubmed: 32005783 pmcid: 7140944 doi: 10.1104/pp.19.01529
Herburger, K., Schoenaers, S., Vissenberg, K. & Mravec, J. Shank-localized cell wall growth contributes to Arabidopsis root hair elongation. Nat. Plants 8, 1222–1232 (2022).
pubmed: 36303011 doi: 10.1038/s41477-022-01259-y
Zabotina, O. A. et al. Mutations in multiple XXT genes of Arabidopsis reveal the complexity of xyloglucan biosynthesis. Plant. Physiol. 159, 1367–1384 (2012).
pubmed: 22696020 pmcid: 3425184 doi: 10.1104/pp.112.198119
Ohmiya, Y. et al. Evidence that endo-1,4-beta-glucanases act on cellulose in suspension-cultured poplar cells. Plant. J. 24, 147–158 (2000).
pubmed: 11069690 doi: 10.1046/j.1365-313x.2000.00860.x
Peaucelle, A., Braybrook, S. & Hofte, H. Cell wall mechanics and growth control in plants: the role of pectins revisited. Front. Plant. Sci. 3, 121 (2012). A minireview that attempts to integrate the concepts of growth control by pectins and by expansin-induced loosening of cellulose. The authors speculate that pectin control is an evolutionarily primitive system that emerged in algae and that expansin-mediated control emerged later.
pubmed: 22685449 pmcid: 3368173 doi: 10.3389/fpls.2012.00121
Haas, K. T., Wightman, R., Peaucelle, A. & Hofte, H. The role of pectin phase separation in plant cell wall assembly and growth. Cell Surf. 7, 100054 (2021).
pubmed: 34141960 pmcid: 8185244 doi: 10.1016/j.tcsw.2021.100054
Taiz, L. Plant-cell expansion — regulation of cell-wall mechanical-properties. Annu. Rev. Plant. Physiol. 35, 585–657 (1984).
doi: 10.1146/annurev.pp.35.060184.003101
Levesque-Tremblay, G., Pelloux, J., Braybrook, S. A. & Muller, K. Tuning of pectin methylesterification: consequences for cell wall biomechanics and development. Planta 242, 791–811 (2015).
pubmed: 26168980 doi: 10.1007/s00425-015-2358-5
Peaucelle, A. et al. Pectin-induced changes in cell wall mechanics underlie organ initiation in Arabidopsis. Curr. Biol. 21, 1720–1726 (2011).
pubmed: 21982593 doi: 10.1016/j.cub.2011.08.057
Milani, P., Braybrook, S. A. & Boudaoud, A. Shrinking the hammer: micromechanical approaches to morphogenesis. J. Exp. Bot. 64, 4651–4662 (2013).
pubmed: 23873995 doi: 10.1093/jxb/ert169
Chebli, Y. & Geitmann, A. Cellular growth in plants requires regulation of cell wall biochemistry. Curr. Opin. Cell Biol. 44, 28–35 (2017).
pubmed: 28131101 doi: 10.1016/j.ceb.2017.01.002
Phyo, P. et al. Gradients in wall mechanics and polysaccharides along growing inflorescence stems. Plant. Physiol. 175, 1593–1607 (2017).
pubmed: 29084904 pmcid: 5717741 doi: 10.1104/pp.17.01270
Liberman, M. et al. Mung bean hypocotyl homogalacturonan: localization, organization and origin. Ann. Bot. 84, 225–233 (1999).
doi: 10.1006/anbo.1999.0914
Wang, X., Wilson, L. & Cosgrove, D. J. Pectin methylesterase selectively softens the onion epidermal wall yet reduces acid-induced creep. J. Exp. Bot. 71, 2629–2640 (2020).
pubmed: 32006044 pmcid: 7210771 doi: 10.1093/jxb/eraa059
Safran, J. et al. Plant polygalacturonase structures specify enzyme dynamics and processivities to fine-tune cell wall pectins. Plant. Cell 35, 3073–3091 (2023).
pubmed: 37202370 doi: 10.1093/plcell/koad134
Palin, R. & Geitmann, A. The role of pectin in plant morphogenesis. Bio Syst. 109, 397–402 (2012).
Haas, K. T., Wightman, R., Meyerowitz, E. M. & Peaucelle, A. Pectin homogalacturonan nanofilament expansion drives morphogenesis in plant epidermal cells. Science 367, 1003–1007 (2020).
pubmed: 32108107 pmcid: 7932746 doi: 10.1126/science.aaz5103
Altartouri, B. et al. Pectin chemistry and cellulose crystallinity govern pavement cell morphogenesis in a multi-step mechanism. Plant. Physiol. 181, 127–141 (2019).
pubmed: 31363005 pmcid: 6716242 doi: 10.1104/pp.19.00303
Belteton, S. A. et al. Real-time conversion of tissue-scale mechanical forces into an interdigitated growth pattern. Nat. Plants 7, 826–841 (2021).
pubmed: 34112988 doi: 10.1038/s41477-021-00931-z
Braybrook, S. A., Hofte, H. & Peaucelle, A. Probing the mechanical contributions of the pectin matrix: insights for cell growth. Plant. Signal. Behav. 7, 1037–1041 (2012).
pubmed: 22836501 pmcid: 3474675 doi: 10.4161/psb.20768
White, P. B., Wang, T., Park, Y. B., Cosgrove, D. J. & Hong, M. Water-polysaccharide interactions in the primary cell wall of Arabidopsis thaliana from polarization transfer solid-state NMR. J. Am. Chem. Soc. 136, 10399–10409 (2014).
pubmed: 24984197 doi: 10.1021/ja504108h
Cho, H.-T. & Cosgrove, D. J. in Plant Hormones: Biosynthesis, Signal Transduction, Action! (ed. Davies P. J.) 262–281 (Springer Netherlands, 2010).
Pacifici, E., Di Mambro, R., Dello Ioio, R., Costantino, P. & Sabatini, S. Acidic cell elongation drives cell differentiation in the Arabidopsis root. EMBO J. 37, e99134 (2018).
pubmed: 30012836 pmcid: 6092616 doi: 10.15252/embj.201899134
Ramakrishna, P. et al. EXPANSIN A1-mediated radial swelling of pericycle cells positions anticlinal cell divisions during lateral root initiation. Proc. Natl Acad. Sci. USA 116, 8597–8602 (2019).
pubmed: 30944225 pmcid: 6486723 doi: 10.1073/pnas.1820882116
Vermeer, J. E. et al. A spatial accommodation by neighboring cells is required for organ initiation in Arabidopsis. Science 343, 178–183 (2014).
pubmed: 24408432 doi: 10.1126/science.1245871
Keuskamp, D. H. et al. Blue-light-mediated shade avoidance requires combined auxin and brassinosteroid action in Arabidopsis seedlings. Plant. J. 67, 208–217 (2011).
pubmed: 21457374 doi: 10.1111/j.1365-313X.2011.04597.x
Rauf, M. et al. NAC transcription factor SPEEDY HYPONASTIC GROWTH regulates flooding-induced leaf movement in Arabidopsis. Plant. Cell 25, 4941–4955 (2013).
pubmed: 24363315 pmcid: 3903997 doi: 10.1105/tpc.113.117861
Braybrook, S. A. & Peaucelle, A. Mechano-chemical aspects of organ formation in Arabidopsis thaliana: the relationship between auxin and pectin. PLoS ONE 8, e57813 (2013).
pubmed: 23554870 pmcid: 3595255 doi: 10.1371/journal.pone.0057813
Fleming, A. J., Caderas, D., Wehrli, E., McQueen-Mason, S. & Kuhlemeier, C. Analysis of expansin-induced morphogenesis on the apical meristem of tomato. Planta 208, 166–174 (1999).
doi: 10.1007/s004250050546
Reinhardt, D., Wittwer, F., Mandel, T. & Kuhlemeier, C. Localized upregulation of a new expansin gene predicts the site of leaf formation in the tomato meristem. Plant. Cell 10, 1427–1437 (1998).
pubmed: 9724690 pmcid: 144079 doi: 10.1105/tpc.10.9.1427
Nakayama, N. et al. Mechanical regulation of auxin-mediated growth. Curr. Biol. 22, 1468–1476 (2012).
pubmed: 22818916 doi: 10.1016/j.cub.2012.06.050
Sampathkumar, A., Yan, A., Krupinski, P. & Meyerowitz, E. M. Physical forces regulate plant development and morphogenesis. Curr. Biol. 24, R475–483 (2014).
pubmed: 24845680 pmcid: 4049271 doi: 10.1016/j.cub.2014.03.014
Bacete, L. et al. THESEUS1 modulates cell wall stiffness and abscisic acid production in Arabidopsis thaliana. Proc. Natl Acad. Sci. USA 119, e2119258119 (2022).
pubmed: 34949719 doi: 10.1073/pnas.2119258119
Gonneau, M. et al. Receptor kinase THESEUS1 is a rapid alkalinization factor 34 receptor in Arabidopsis. Curr. Biol. 28, 2452–2458.e2454 (2018).
pubmed: 30057301 doi: 10.1016/j.cub.2018.05.075
Smokvarska, M. et al. The receptor kinase FERONIA regulates phosphatidylserine localization at the cell surface to modulate ROP signaling. Sci. Adv. 9, eadd4791 (2023).
pubmed: 37027473 pmcid: 10081841 doi: 10.1126/sciadv.add4791
Pacheco, J. M. et al. Cell surface receptor kinase FERONIA linked to nutrient sensor TORC signaling controls root hair growth at low temperature linked to low nitrate in Arabidopsis thaliana. New Phytol. 238, 169–185 (2023).
pubmed: 36716782 doi: 10.1111/nph.18723
Mecchia, M. A. et al. The single Marchantia polymorpha FERONIA homolog reveals an ancestral role in regulating cellular expansion and integrity. Development 149, dev200580 (2022).
pubmed: 36178124 doi: 10.1242/dev.200580
Shih, H. W., Miller, N. D., Dai, C., Spalding, E. P. & Monshausen, G. B. The receptor-like kinase FERONIA is required for mechanical signal transduction in Arabidopsis seedlings. Curr. Biol. 24, 1887–1892 (2014).
pubmed: 25127214 doi: 10.1016/j.cub.2014.06.064
Li, C., Wu, H. M. & Cheung, A. Y. FERONIA and her pals: functions and mechanisms. Plant. Physiol. 171, 2379–2392 (2016).
pubmed: 27342308 pmcid: 4972288 doi: 10.1104/pp.16.00667
Wang, P. et al. Integrated omics reveal novel functions and underlying mechanisms of the receptor kinase FERONIA in Arabidopsis thaliana. Plant. Cell 34, 2594–2614 (2022).
pubmed: 35435236 pmcid: 9252503 doi: 10.1093/plcell/koac111
Steinwand, B. J. et al. Alterations in auxin homeostasis suppress defects in cell wall function. PLoS ONE 9, e98193 (2014).
pubmed: 24859261 pmcid: 4032291 doi: 10.1371/journal.pone.0098193
Anderson, C. T. & Kieber, J. J. Dynamic construction, perception, and remodeling of plant cell walls. Annu. Rev. Plant. Biol. 71, 39–69 (2020).
pubmed: 32084323 doi: 10.1146/annurev-arplant-081519-035846
Kohorn, B. D. & Kohorn, S. L. The cell wall-associated kinases, WAKs, as pectin receptors. Front. Plant. Sci. 3, 88 (2012).
pubmed: 22639672 pmcid: 3355716 doi: 10.3389/fpls.2012.00088
Wagner, T. A. & Kohorn, B. D. Wall-associated kinases are expressed throughout plant development and are required for cell expansion. Plant. Cell 13, 303–318 (2001).
pubmed: 11226187 pmcid: 102244 doi: 10.1105/tpc.13.2.303
Yue, Z. L. et al. The receptor kinase OsWAK11 monitors cell wall pectin changes to fine-tune brassinosteroid signaling and regulate cell elongation in rice. Curr. Biol. 32, 2454–2466.e7 (2022).
pubmed: 35512695 doi: 10.1016/j.cub.2022.04.028
Hamant, O., Inoue, D., Bouchez, D., Dumais, J. & Mjolsness, E. Are microtubules tension sensors? Nat. Commun. 10, 2360 (2019).
pubmed: 31142740 pmcid: 6541610 doi: 10.1038/s41467-019-10207-y
Li, J., Szymanski, D. B. & Kim, T. Probing stress-regulated ordering of the plant cortical microtubule array via a computational approach. BMC Plant Biol. 23, 308 (2023).
pubmed: 37291489 pmcid: 10251582 doi: 10.1186/s12870-023-04252-5
Fruleux, A., Verger, S. & Boudaoud, A. Feeling stressed or strained? A biophysical model for cell wall mechanosensing in plants. Front. Plant. Sci. 10, 757 (2019).
pubmed: 31244875 pmcid: 6581727 doi: 10.3389/fpls.2019.00757
Williamson, R. E. Alignment of cortical microtubules by anisotropic wall stresses. Aust. J. Plant. Physiol. 17, 601–613 (1990).
Decreux, A. & Messiaen, J. Wall-associated kinase WAK1 interacts with cell wall pectins in a calcium-induced conformation. Plant Cell Physiol. 46, 268–278 (2005).
pubmed: 15769808 doi: 10.1093/pcp/pci026
Hamilton, E. S. et al. Mechanosensitive channel MSL8 regulates osmotic forces during pollen hydration and germination. Science 350, 438–441 (2015).
pubmed: 26494758 pmcid: 4764502 doi: 10.1126/science.aac6014
Chebli, Y., Bidhendi, A. J., Kapoor, K. & Geitmann, A. Cytoskeletal regulation of primary plant cell wall assembly. Curr. Biol. 31, R681–R695 (2021).
pubmed: 34033798 doi: 10.1016/j.cub.2021.03.092
Leyser, O. Auxin signaling. Plant. Physiol. 176, 465–479 (2018).
pubmed: 28818861 doi: 10.1104/pp.17.00765
Friml, J. et al. ABP1–TMK auxin perception for global phosphorylation and auxin canalization. Nature 609, 575–581 (2022).
pubmed: 36071161 doi: 10.1038/s41586-022-05187-x
Lin, W. et al. TMK-based cell-surface auxin signalling activates cell-wall acidification. Nature 599, 278–282 (2021).
pubmed: 34707287 pmcid: 8549421 doi: 10.1038/s41586-021-03976-4
Minami, A., Takahashi, K., Inoue, S. I., Tada, Y. & Kinoshita, T. Brassinosteroid induces phosphorylation of the plasma membrane H
pubmed: 30649552 doi: 10.1093/pcp/pcz005
Ajeet, C. et al. Brassinosteroid recruits FERONIA to safeguard cell expansion in Arabidopsis. Preprint at bioRxiv 2023.2010.2001.560400, https://doi.org/10.1101/2023.10.01.560400 (2023).
Haruta, M., Gray, W. M. & Sussman, M. R. Regulation of the plasma membrane proton pump (H
pubmed: 26476298 pmcid: 4679459 doi: 10.1016/j.pbi.2015.09.005
Moussu, S. et al. Structural basis for recognition of RALF peptides by LRX proteins during pollen tube growth. Proc. Natl Acad. Sci. 117, 7494 (2020).
pubmed: 32165538 pmcid: 7132299 doi: 10.1073/pnas.2000100117
Herger, A., Dunser, K., Kleine-Vehn, J. & Ringli, C. Leucine-rich repeat extensin proteins and their role in cell wall sensing. Curr. Biol. 29, R851–R858 (2019).
pubmed: 31505187 doi: 10.1016/j.cub.2019.07.039

Auteurs

Daniel J Cosgrove (DJ)

Department of Biology, Pennsylvania State University, University Park, Pennsylvania, USA. dcosgrove@psu.edu.
ORCID: 0000-0002-4020-5786

Classifications MeSH