Plant Physiology

doi: 10.1104/pp.17.01763pmid: 29317522

doi: 10.1104/pp.17.01763pmid: N/A

The editors and staff of Plant Physiology thank all of those listed below whose insight and contributions in reviewing manuscripts from December 8, 2016, to December 7, 2017, have helped make the journal a success. We would also like to thank the Monitoring Editors who have concluded their term of service, and we welcome those Monitoring Editors who are joining the Ed Board in 2018. REVIEWERS Mark G.M. Aarts Pierre Abad Steffen Abel Maria Jazmin Abraham-Juarez Isabel A. Abreu Shunsuke Adachi Hiroaki Adachi Zach Adam Bartosz Adamczyk Keith L. Adams Ji Hoon Ahn Mitsuhiro Aida Elizabeth Ainsworth Tariq A. Akhtar Eduard Akhunov Garo Akmakjian Emre Aksoy David Alabadi Josefa M. Alamillo Salim Al-Babili Nick W.S. Albert Alessandro Alboresi Rubén Alcázar Yagut Allahverdiyeva-Rinne Andrew Charles Allan Douglas K. Allen Ahmad M. Alqudah Ihsan A. Al-Shehbaz Rubén Rellán Álvarez Rafael de Prado Amian Gynheung An Charles T. Anderson Ryan G. Anderson Mats X. Andersson Vasilios Andriotis Fernando Aniento Aldwin M. Anterola Takashi Aoyama Jens Appel Ingo Appelhagen Edgar Demesa Arevalo German Martinez Arias Shin-ichi Arimura Steven A. Arisz William Armstrong Vincent Arondel Tadao Asami Hiroshi Ashihara Motoyuki Ashikari Israel Ausin Tamar Avin-Wittenberg Michael J. Axtell Brian G. Ayre Ivan Baccelli Thomas J. Bach Murray Badger Shu-Nong Bai Soren Bak Bénédicte Bakan Sureshkumar Balasubramanian Patricia Baldrich Ian T. Baldwin Janneke Balk Marilyn C. Ball Steven G. Ball Carlos L. Ballare Kelly Balmant Maya Bar François Barbier Margaret M. Barbour Muriel Bardor Bronwyn J. Barkla Fredy Barneche Celia Baroux Jaime Barros-Rios Cornelius S. Barry Andrea Barta Dorothea Bartels Khurram Bashir Georg Basler Hank Bass Gilles Basset Roberto Bassi Elias Bassil Philip David Bates Henri Batoko Marie Baucher Petra Bauer Isabel Baurle Ivan R. Baxter Frederic Beaudoin Claude Becker Jorg D. Becker Gerold Beckers Diane M. Beckles Patricia A. Bedinger Gerrit T.S. Beemster David Beerling Richard Belanger Daniela Bellincampi Michele Bellucci M.P. Benavides Abdelhafid Bendahmane Moussa Benhamed Christoph Benning Andrew F. Bent Stephane Bentolila Frederic Berger Dominique Bergmann Gerald Berkowitz Oliver Berkowitz Lien Bertier Edoardo Bertolini Sebastien Besseau Rishikesh Bhalerao Thomas S. Bibby Amir Bidhendi Gerd Patrick Bienert Wolfgang Bilger Stefan Binder James A. Birchler David A. Bird Kenneth D. Birnbaum Gerard James Bishop Ton Bisseling Christopher Black Joshua J. Blakeslee Elison B. Blancaflor Robert E. Blankenship Miguel A. Blazquez Andreas Blennow Bernhard Blob Anna Block Maryse A. Block Cecilia K. Blomstedt Tiina Blomster Arnold Jeffrey Bloom Eduardo Blumwald Scott Andrew Boden Wout A. Boerjan Jochen Bogs Joerg Bohlmann Alexandra Bohne Tomas Bohr Aurelien Boisson-Dernier Cordelia Bolle Thomas Boller Atle Bones Paola Bonfante Peta C. Bonham-Smith Cristina Bonza Michael Borg Ljudmilla Borisjuk Frederik Börnke Gerd Bossinger Miguel A. Botella Javier Francisco Botto Frédéric Bouché David Bouchez Arezki Boudaoud Cécile Bousquet-Antonelli John L. Bowman Lewis Bowman Tolga Osman Bozkurt Helen Brabham Hans-Peter Braun Andrea Bräutigam Melissa Brazier-Hicks Philip B. Brewer Craig Brodersen Peter Brodersen Mikael Brosche John W.S. Brown Kathleen Brown Patrick Hugh Brown Karen S. Browning John Browse Friederike Bruessow Tzvetina Brumbarova Amy Brunner Kerry Bubb Thomas J. Buckhout Thomas N. Buckley Vincent Bulone Daniele Del Buono Tessa Burch-Smith Vincent Burlat Florian A. Busch Daniel Bush Eugenio Butelli Melinka A. Butenko Nathaniel M. Butler Diana Mihaela Buzas Caitlin Siobhan Byrt Giampiero Cai Cristiane Calixto Judy Callis Douglas A. Campbell Josep Canavate Maura C. Cannon Francisco Javier Cano Shuqing Cao Allan B. Caplan Maura Cardarelli Elizabete Carmo-Silva Nicholas C. Carpita John P. Carr Jorge Jose Casal Paula Casati Gladys I. Cassab Carmen Castresana Stefano Cazzaniga Francisco J. Cejudo Jose Miguel Celedon Lucas A. Cernusak Martin Cerny Heriberto D. Cerutti Felice Cervone Usawadee Chaiprom Sanhita Chakraborty Krishna Reddy Challa Kai Xun Chan Kai Chan Kent D. Chapman Joe Chappell Clint Chapple Yee-Yung Charng Guillaume Charrier Caspar Chater Francois Chaumont Youssef Chebli Changbin Chen Feng Chen Meng Chen Ming-Shun Chen Shaoliang Chen Sixue Chen Xiao-Ya Chen Yi-Fang Chen Zhixiang Chen Zhonghua Chen Yi Chen Liang Chen Dr Yi Chen Youfa Cheng Zong-Ming Cheng Choong-Ill Cheon Christian Chervin Alice Y. Cheung Cécilia Cheval Christian Chevalier Andrea Chini Viswanathan Chinnusamy Tzyy-Jen Chiou Daniel H. Chitwood Brendan Choat Sunghwa Choe Sang-Bong Choi Yeonhee Choi Kang Chong Yi-hsiang Chou Fred W.S. Chow Shawn Christensen John Christie Pascal-Antoine Christin Chengcai Chu George Chuck Randy T. Clark Nicole Kho Clay Andrew Clayton Stephan Clemens Tom E. Clemente Inge De Clercq Herve Cochard Robert Coe Patricia Coello Catharina Coenen Sivio Collani Luca Comai Erin L. Connolly Michael J. Considine David Cook George Coupland Vincent Courdavault Kevin Cox William A. Cramer Martin Crespi Jose L. Crespo Mauro Cresti Peter Alexander Crisp Roberta Croce Felipe Cruz-Garcia Yuhai Cui Ian Cummins Jennifer Cunniff Catherine Curie John C. Cushman Sean R. Cutler Yasin F. Dagdas Firas Bou Daher David A. Dalton Chris D. Dardick Pradeep Das Rebecca Dauwe Christopher Davies Kevin M. Davies Seth J. Davis Brad Day David A. Day Alexis De Angeli Inge De De Clercq Stefan de Folter Gerardo De Leon Vincenzo De Luca Sylvia de Pater Bert De Rybel Ive De Smet Lieven De Veylder Roger Deal Gillian Dean Ross Milton Deans Seth DeBolt Jan P. Dekker Juan Carlos del Pozo John Paul Délano-Frier Carolin Delker Dean DellaPenna Massimo Delledonne Serge Delrot Charles F. Delwiche Sylvain Delzon Jurgen Denecke Xing Wang Deng Thierry M. Desnos Bénédicte Desvoyes Om Parkash Dhankher Kanwarpal Dhugga Antonio Diaz-Espejo Jazz Dickinson Rebecca Dickstein Carol Dieckman Daniela Dietrich Karl-Josef Dietz Feike A. Dijkstra Paul Dijkwel Savithramma P. Dinesh-Kumar Shou-Wei Ding Yezhang Ding Yong Ding Zhaojun Ding Yong Ding Jose R. Dinneny Ram Dixit David P. Dixon Laura Dixon Michael Anthony Djordjevic Monika Susanne Doblin Anna A. Dobritsa Ian C. Dodd Antony Dodd Peter N. Dodds Valerian V. Dolja Jean-Christophe Domec Juan Dong Suomeng Dong Xinnian Dong Jia Dong Teresa Donze-Reiner Fiona M. Doohan Marcela C. Dotto Jeff J. Doyle GA Drakakaki John Drake Paul L. Drake Thomas Dresselhaus Steven Michiel Driever Professor Liqun Du Liqun Du Jorge Dubcovsky Marieke Dubois Christian Dubos Stephen O. Duke Paul Dupree Lionel Dupuy Paula Duque Joerg Durner John M. Dyer Derek Eamus Peter J. Eastmond Berit Ebert Patrick P. Edger Erika Edwards Everard J. Edwards Gerald E. Edwards Juergen Ehlting Cornelia Eisenach R.J. Neil Emery Michael J. Emes Masaki Endo Ingo Ensminger Bernard L. Epel Matthias Erb Yuval Eshed Mark Estelle Jose Estevez Thomas Eulgem Matthew Evans Chris Exley Peter J. Facchini Ahmed Faik Jiří Fajkus Huihui Fang Christian Fankhauser Mario Fares Edward E. Farmer Graham D. Farquhar Eva M. Farre Sara Farrona Bruno Favery Attila Feher Zhangjun Fei Jose A. Feijo Georg Felix Antje C. Feller Matyas Fendrych Baomin Feng Emilio Fernandez Alisdair R. Fernie Cristina Ferrandiz Ivo Feussner Oliver Fiehn Sergei A. Filichkin Giovanni Finazzi Ruth R. Finkelstein Iris Finkemeier Fabio Fiorani Anne-Sophie Fiorucci Teresa B. Fitzpatrick Andrew J. Fleming Christina Fliege Padraic J. Flood Francisco J. Florencio Victor Flors Tim Flowers Robert Fluhr Elizabeth P.B. Fontes Eloise Foo Enrique Martinez Force Brian G. Forde Ana M. Fortes Vasileios Fotopoulos Fabrice Foucher John E. Fowler Christine H. Foyer Keara A. Franklin Vernonica E. Franklin-Tong Peter J. Franks Henk Franssen Analiese Franz Valerie Fraser Rupert George Fray Michael Freeling Monika Frey Wieland Fricke Lorenzo Frigerio Jiří Friml Rikard Fristedt Hillel Fromm Florian Frugier Ying Fu Xiangdong Fu Yasunari Fujita Toru Fujiwara Takeshi Fukao Akihito Fukodome Hideya Fukuzawa Dietmar Funck Dominique Gagliardi Jorge Gago Todd A. Gaines Massimo Galbiati David W. Galbraith Gad Galili Andrea Gallavotti Aurora Galvan Pascal Gamas Dongying Gao Győző Garab Carlos Garcia-Mata Gary Gardner Kimberly Garland-Campbell Debora Gasperini Christiane Gatz Sonia Gazzarrini Yufeng Ge Tsanko Gechev Danny N.V. Geelen Chris Gehring Peter Geigenberger Markus Geisler Niko Geldner Anthony R. Gendall Bernard Genty Milen I. Georgiev Jonathan Gershenzon Koen Geuten Godelieve Gheysen Daniel Gibbs Mike Gidley Ricardo Fabiano Hettwer Giehl Mark Gijzen Matthew E. Gilbert Glenda E. Gillaspy Simon Gilroy James J. Giovannoni Andrew Gipson Anthony D.M. Glass Gaetan Glauser Jane Glazebrook David Goad Ian D. Godwin Alexander Goldschmidt Michel Goldschmidt-Clermont Gregory Goldsmith Charlotte M.M. Gommers Jiming Gong Eliana Gonzales-Vigil Daniel H. Gonzalez Jonas Goossens Michael A. Gore Daphne Goring Sven B. Gould Aska Goverse Emmanuelle Graciet Michelle A. Graham Antonio Granell Christine Granier Katja Graumann Gordon R. Gray Julie E. Gray William M. Gray Ian K. Greaves Markus Grebe Beverley R. Green Rachel M. Green Brian D. Gregory Peter Michael Gresshoff Etienne Grienenberger Lawrence R. Griffing Howard Griffiths L. Grillet Jacqueline Grima-Pettenati Bernhard Grimm Erich Grotewold Georg Groth Martin Groth Anil Grover Xiaofeng Gu Xingyou Gu Ying Gu Jose Manuel Gualberto Michael T. Guarnieri Frederique C. Guinel Fang-Qing Guo Hongwei Guo Hui Shan Guo Yan Guo Zhongxin Guo Mainak Das Gupta Camilla Gustafsson Kirstin Gutekunst Michael Gutensohn Crisanto Gutierrez Emilio Gutierrez-Beltran Caroline Gutjahr Georg Haberer Uwe G. Hacke Martin Hagemann Nigel G. Halford Barbara Ann Halkier Karen Jane Halliday Larry J. Halverson Bjoern Hamberger John D. Hamill Ulrich Zeno Hammes John P. Hammond Shaojie Han Yi Han Kosuke Hanada Marc Hanikenne Guy Hanke Meredith T. Hanlon Matthew A. Hannah David J. Hannapel Andrew D. Hanson David T. Hanson Johannes Hanson Mats Hansson Yoshie Hanzawa Yu-Jin Hao Jeremy Harbinson Christian S. Hardtke Stacey Harmer Jeanne M. Harris Philip J. Harris S. Hartley John L. Harwood Mirza Hasanuzzaman Mike Hasegawa Takashi Hashimoto Richard P. Haslam Tegan M. Haslam Ronald D. Hatfield George Haughn Bettina Hause Michel Havaux Chris Hawes Tim R. Hawkes Angela Hay Fiona R. Hay Ken-ichiro Hayashi Scott Hayes Joel Haywood Jan Hazebroek Junxian He Ping He Xin-Jian He Yuehui He Guangming He Kim Henrik Hebelstrup Kim Hebelstrup Peter Hedden Antje Heese Manfred Heinlein Marcus G. Heisler Jan Hejatko Yrjo Helariutta Tieme Helderman Piers A. Hemsley Jung-ok Heo Eliot M. Herman Miguel A. Hernandez-Prieto Joshua R. Herr Christine Hervé Wolfgang R. Hess Alistair M. Hetherington Tarek Hewezi Julian M. Hibberd Lee Hickey Kouki Hikosaka David Hildebrand Henk W.M. Hilhorst Monika Hilker Adrian Hills Pierre Hilson Andreas Hiltbrunner Dirk K. Hincha Michael Hippler Ko Hirano Bertrand Hirel Cory D. Hirsch Heribert Hirt Toru Hisabori Uri Hochberg Ute Hoecker Rainer Hoefgen Georg Hoelzl Daniel Hofius Herman Hofte N. Michele Holbrook Michael J. Holdsworth Ben F. Holt Jia Honglei David Honys Paul J.J. Hooykaas Brian Hopkinson Hanna Horak Harry T. Horner Walter J. Horst Stefan Hortensteiner Christopher J. Howe Stephen H. Howell Chuan-hih Hsu Polly Hsu Hanhua Hu Jianping Hu Junjie Hu Yuxin Hu Wei Hu Anthony H.C. Huang Jirong Huang Sanwen Huang Shanjin Huang Matthew Hudson Peter Huijser Lee Hunt Charles T. Hunter Enamul Huq Vaughan Hurry Aman Husbands Søren Husted Inhwan Hwang Kazuya Ichimura Yoko Iijima Ryozo Imai Takato Imaizumi Richard G.H. Immink Takehito Inaba Roger W. Innes Yoshiaki Inukai Dirk Inzé Vivian F. Irish Sandra Irmisch Till Ischebeck Sumie Ishiguro Jean-Charles Isner Emmanuelle Issakidis-Bourguet Toshiro Ito Alexander A. Ivakov Anjali S. Iyer-Pascuzzi Joe Jacobowitz Anna L. Jacobsen Yvon Jaillais Euan K. James Georg Jander Jyan-Chyun Jang Steven Jansen Dick B. Janssen Jose A. Jarillo Michael C. Jarvis Paul Jarvis Andrzej Jerzmanowski Mareike Jezek Zhongtao Jia Liwen Jiang Ning Jiang Yuling Jiao José Jiminez Jin-Pu Jin Henrik Johansson Giles N. Johnson Kim L. Johnson Mark A. Johnson Matt Johnson Matthew P. Johnson Xenie Johnson Alan M. Jones Matthew A. Jones Matthieu H.A.J. Joosten Lucia Jordá Gregory J. Jordan Juliette Jouhet Mithila Jugulam Ivan Juric Aardra Kachroo Yasuhiro Kadota Shawn M. Kaeppler Brent N. Kaiser Kaisa Kajala Kriton Kalantidis Lothar Kalmbach Masatake Kanai Masahiro Kanaoka Angelos K. Kanellis Zhensheng Kang Joohyun Kang Saijaliisa Kangasjärvi Stanislaw Karpinski Hiroyuki Kasahara Fumiaki Katagiri Yusuke Kato Shiv Shankhar Kaundun Tsutomu Kawasaki Kenneth Keegstra Beat Keller Tim Kelliher Elizabeth A. Kellogg Pavel I. Kerchev Nir Keren Andre Kessler Felix Kessler Sharon A. Kessler Diederik H. Keuskamp Minsung Kim Woo Taek Kim Yoomee Kim Hyojung Kim Jeong Im Kim Daewon Kim Seisuke Kimura Anthony J. Kinney Toshinori Kinoshita Helmut Kirchhoff Viktor Kirik Diana Kirilovsky Jurgen Kleine-Vehn Daniel J. Kliebenstein Marc R. Knight Donghwi Ko Kappei Kobayashi Leon Vincent Kochian Daniel Koenig Claudia Kohler Hisashi Koiwa Nozomu Koizumi Hannes Kollist Josef Komenda Eva Kondorosi Abraham J.K. Koo Maarten Koornneef Peter M. Kopittke Joachim Kopka Anna Koprivova Maria von Korff Tomokazu Koshiba Dylan K. Kosma Klara Kosova Toshihisa Kotake Hiroyuki Koyama Laszlo Kozma-Bognar Friedrich Kragler David M. Kramer Björn Krenz Anja Krieger-Liszkay Beth A. Krizek Thomas Kroj Johannes Kromdijk Karin Krupinska Patrick J. Krysan Hannah Kuhn Ivan Kulich Fabian Kunzl Hendrik Kupper Dorota Kwiatkowska Tina Kyndt Junko Kyozuka Benoit Lacroix Debabrata Laha Agu Laisk Jean-Francois Laliberte Hans Lambers Zhaobo Lang Jane A. Langdale Bernd Markus Lange Caspar Langenbach Anthony William Larkum Paul B. Larsen Emily R. Larson Therese LaRue Sascha Laubinger Patrick Laufs Tracy Lawson Ana M. Laxalt Peter J. Lea Andrew Leakey Byeong-ha Lee Hojoung Lee Ilha Lee Ji-Young Lee Myeong Min Lee Youngsook Lee Dr. Hyo-Jun Lee Richard C. Leegood Benoit Lefebvre Cecile Lefoulon Mingguang Lei Dario Leister Stephane D. Lemaire Michael Lenhard Patricia Leon Verhage Leonie Jeffrey Leung Thomas Leustek Amit Levy Norman G. Lewis Mathew Graham Lewsey Chuanyou Li Gang Li Jia Li Lei Li Li Li Lin Li Ping Li Qing Li Qingliang Li Xia Li Xin Li Xu Li Xuan Li Yi Li Yunhai Li Xiaoyang Li Ying Li Jie Liang Wanqi Liang Hong Liao Conrad Paul Lichtenstein David Lightfoot Cathrine Lillo Erik Limpens Rongcheng Lin Senjie Lin Wen-Hui Lin Marika Lindahl Pia Lindberg Christian Lindermayr Volker Lipka Emmanuel Liscum Amy Litt Bao Liu Bo Liu Chang-Jun Liu Chun Yan Liu Dong Liu Haipei Liu Hongtao Liu Ming-Jung Liu Qiao Quan Liu Yao-Guang Liu Yongxiu Liu Yule Liu Zhongchi Liu Dianyi Liu James R. Lloyd Gary J. Loake Guillume Lobet Martin Lohr Terri A. Long Yanping Long Patricia Lopez Enrique Lopez-Juez Ann E. Loraine Rafael Lozano Rosa Lozano-Duran Chaofu Lu Dihong Lu Hua Lu Yan Lu Ying-Tang Lu Sheng Luan Martha Ludwig John E. Lunn Chongyuan Luo Hong Luo Ming Luo Zhibin Luo Joseph Lynch Hong Ma Jian Feng Ma Jianxin Ma Lijun Ma You-Zhi Ma Xiaofang Ma David MacAlpine John J. MacKay Sally A. Mackenzie Dan Maclean Sofia Madeira Andreas Madlung Francisco Madueno Takaki Maekawa Jurandir Magalhaes Kashif Mahmood Alexis Maizel Kristiina Maria Mäkinen Eugene Maksimov Jocelyn Malamy Pal Maliga Julin N. Maloof Pablo Andrés Manavella Jun'ichi Mano Shoji Mano Patricia Manosalva Concepcion Manzano-Fernandez Long Mao Eric Marechal Daniel Marino Lyza Gontow Maron Cathie Martin Ruth Martin Eleazar Martinez-Barajas Enrico Martinoia Samuel C.V. Martins Simona Masiero Richard Esten Mason Patrick H. Masson Sarah Mathews Jaideep Mathur Shizue Matsubara Yusuke Matsuda Sachihiro Matsunaga Takuya Matsuo Michaela Matthes Paul D. Matthews Felix Mauch Christophe Maurel Stefan Mayr Scott A.M. McAdam Bruce McClure Karen McGinnis Leah McHale Hazel McLellan Ryan McQuinn Katherine Georgina Meacham Belinda Medlyn Hipolito Medrano Andrew A. Meharg Iris Meier David Meinke Rick Meinzer Anastasios Melis Joanna Melonek Maeli Melotto Johan Memelink Benoit Menand David Mendoza-Cozatl Raphael Mercier Remy Merret Joerg Meurer Christian Meyer Knut Meyer Blake C. Meyers Alexander Meyers Amna Mhamdi Scott D. Michaels Jose Luis Micol A. Harvey Millar Andrew J. Millar Allison J Miller Tony J. Miller Peter V. Minorsky Michael Mishkind Patrick John Mitchell Rowan A.C. Mitchell Shin-ya Miyagishima Nobuyoshi Mochizuki Thomas Mock Peter Moffett Gaurav Moghe Debra Mohnen Antonio Molina Isabel Molina Ian Max Møller Attila Molnar Gabriele B. Monshausen Elena Monte Bethany Moore Caitlin E. Moore Ian Moore Takaya Moriguchi Tomas Morosinotto Richard J. Morris Adam Mott Yaseen Mottiar Zhonglin Mou Grégory Mouille Daniel S. Moura Wellington Muchero Gloria K. Muday Gary J. Muehlbauer Martin J. Mueller Douglas G. Muench Christopher David Muir Karolina Mukhtar Shahid Mukhtar Conrad W. Mullineaux Sergi Munné-Bosch Yoshiyuki Murata Erik H. Murchie Denis J. Murphy Jeremy D. Murray Jorge P. Muschietti Angelika Mustroph Marek Mutwil Philippe Nacry Satya Swathi Nadakuduti Raimund Nagel Thomas Nägele Peter Nagy Takahiro Nakamura Kazuo Nakashima Mikio Nakazono Hong Gil Nam Eiji Nambara Richard M. Napier Reena Narsai Utpal Nath Saket Navlakha Andreas Nebenführ Hilde Nelissen David C. Nelson Jennifer Nemhauser Ekkehard Neuhaus Reinat Nevo Jörg Nickelsen Andreas Niebel Chad E. Niederhuth Ulo Niinemets Basil J. Nikolau Ove Nilsson Shuh-ichi Nishikawa Kenji Nishimura Naoko K. Nishizawa Peter J. Nixon Krishna K. Niyogi Graham Noctor Joseph P. Noel Ko Noguchi Markus Nolf Kenichi Nonomura Lorena Norambuena Moritz K. Nowack Kazunari Nozue Vardis Ntoukakis Dmitri A. Nusinow Christian Obermeier Gerhard Obermeyer Devin Lee O'Connor Remko Offringa Yoshiyuki Ogata Riichi Oguchi Kyoko Ohashi-Ito Masaru Ohme-Takagi Hiroyuki Ohta Misato Ohtani Masanori Okamoto Takashi Okamoto Thomas W. Okita Giles E. Oldroyd Neil E. Olszewski Ronan O'Malley Malcolm O'Neill Coast Onoriode Jamie A O'Rourke Colin P. Osborne Oren Ostersetzer Marisa S. Otegui Sofia Otero Thomas Ott Kirk Overmyer Victor Giminez Oya Esma Özhüner Ravishankar Palanivelu Wiliam Palmer Irvin L. Pan Jianwei Pan Shen Quan Pan Olivier Panaud Shree P. Pandey Sona Pandey Yongzhen Pang Emmanuel Panteris Florent Pantin Maria Papanatsiou Jose M. Pardo Micahel Paries Christian Parisod Chung-Mo Park Jiyoung Park Martin Afan John Parry Neha Patel Andra Paterlini Andrea Paterlini Gopal Pattanayak Matthew J. Paul Germain Pauluzzi Markus Pauly Wojtek P. Pawlowski Paxton Payton Javier Paz-Ares Ole Pedersen Bjorn Panyella Pedersen Ullas Pedmale Wendy Ann Peer Soraya Pelaz Steven Penfield Pierdomenico Perata Andy Pereira Francisco Perez-Alfocea Claire Périlleux Giorgio Perrella Carole C. Perry Staffan Persson Edouard Pesquet Reuben J. Peters Morten Petersen Katia Petroni Dimitris Petroutsos Thomas Pfannschmidt Andrew L. Phillips Eran Pichersky Uri Pick John A Pickett Birgit Piechulla Brett Pike Lynn Pillitteri Priya Pimprikar Miguel A. Pineros J. Chris Pires Jon K. Pittman Claude Plassard Christop Plieth Alex Plong Barry J. Pogson Yves Poirier Jacob Pollier Mikhail Pooggin Brigitte Poppenberger Malcolm Possell Martin Potocký Thomas Potuschak Kalika Prasad Christopher Preston Tony P. Pridmore Holger Puchta Aarthi Putarjunan Ji Qi Yiping Qi Qian Qian Hong Qiao Feng Qin Genji Qin Song Qin Yuan Qin Le Qing Qu Rongda Qu Leandro Quadrana Francesca Quattrocchio Kashchandra G. Raghothama Laura Ragni Christine A. Raines Michael T. Raissig Marie-Christine Ralet John Ralph Diana Ramirez-Garces Stefanie Ranf Amanda Rasmussen Allan Rasmusson Joachim Rassow R. George Ratcliffe Pascal Ratet Cyrille B.K. Rathgeber John Rathjen John A. Raven Vivek Raxwal Cécile Raynaud Catherine Rayon Fabrice Rébeillé Anireddy Reddy Ziv Reich Jean-Philippe Reichheld James B. Reid Jorge Rencoret Zdenko Rengel Simon Renny-Byfield Stefan Reuscher Pascal Rey Matthew P. Reynolds Camilo Rey-Sanchez Dr Guilhem Reyt Richard Richards Jose Luis Riechmann Christoph Ringli Eevi Rintamäki Dae-Kyun Ro Christophe Robaglia Semidan Robaina-Estevez Neil Robbins Alison W. Roberts Daniel M. Roberts Jean-David Rochaix Fulton E. Rockwell Pedro L. Rodriguez Jorge Rodriguez-Celma Manuel Rodriguez-Concepcion Rob Roelfsema Keith Roesler Ute Roessner Hilary J. Rogers Sanja Roje Enrique Rojo Jeffrey Rollins Sabine Rosahl Jocelyn K.C. Rose Laura Rossini Pawel Roszak Christophe Rothan Anne-Lise Routier-Kierzkowska Stanley J. Roux Beth Rowan Jordan Rowley Yong-Ling Ruan Alexander V. Ruban Francisco Rubio Vicente Rubio Jason J. Rudd Carmen Marti Ruiz Mark P. Running Scott D. Russell Peter R. Ryan Wojciech Rymaszewski Ari Sadanandom Martin Sagasser Rowan F. Sage Yusuke Saijo Tatsuya Sakai Hitoshi Sakakibara Wataru Sakamoto Soulaiman Sakr Yasuhito Sakuraba Yumiko Sakuragi Patrice A. Salomé Alon Samach Arun Sampathkumar Marcus Samuel A. Lacey Samuels Ana Lopez Sanchez Nattapong Sanguankiattichai Stefano Santabarbara Diana Santelia Tomasz J. Sarnowski Shai I. Saroussi Nobuhiro Sasaki Takayuki Sasaki Takayo Sasaki Rashmi Sasidharan Christopher Saski Masa H. Sato Yutaka Sato Michael B. Sauer Norbert Sauer Margret Sauter Arnould Savoure Enrico Scarpella Gabriel Schaaf Daniel P. Schachtman Patrick Schaefer Anton R. Schaeffner Arthur A. Schaffer G. Eric Schaller Dierk Scheel Renate Scheibe H. Jochen Schenk Adam Schikora Anthony L. Schilmiller Jos H.M. Schippers Henriette Schluepmann Eric A. Schmelz Karl Schmid Wolfgang Schmidt Robert Schmitz Christian Schmitz-Linneweber James Schnable Korbinian Schneeberger Dirk Schneider Danny J. Schnell Joerg-Peter Schnitzler Thorsten Schnurbusch Herman B. Scholthof Mark Aurel Schöttler Karl Schreiber Michael Schroda Julian I. Schroeder Sven Schubert Veit Schubert Mathias Schuetz Alexander Schulz Georg E. Schulz Karin Schumacher Gadi Schuster Wilfried Schwab Alois Schweighofer Fabian Schweizer Patrick Schweizer Jorg Schwender Kathy Schwinn Steven R. Scofield Samuel Seaver John C. Sedbrook Ronald R. Sederoff Armand Seguin Thorsten Seidel Jennifer Selinski Herve Sentenac Hak Soo Seo Graham B. Seymour Jon Shaff Jyoti Shah Libo Shan Eilon Shani Thomas D. Sharkey Robert E. Sharwood Sidney L. Shaw Wen-Biao Shen Wen-Hui Shen Craig Sherman Anya Sherwood Huazhong Shi Yulan Shi Naoto Shibuya Toshiharu Shikanai Hiroyuki Shimono Jay Shockey Elena D. Shpak Andrew John Simkin Sara Simonini Shanteri Singh Jasleen Singh Folke Sitbon Michael Skelly Rebecca A. Slattery R. Keith Slotkin Jan Smalle Sjef C.M. Smeekens Andrei Smertenko Nicholas Smirnoff Alison G. Smith Alison M. Smith Rebecca A. Smith Stacey Smith Steven M. Smith William J. Snell Roman Sobotka Moon-Soo Soh Doug Soltis David E. Somers Imre E. Somssich Chun-Peng Song Fengming Song Liang Song Qingfeng Song Sophia Sonnewald Kathleen L. Soole Mariana Sottomayor Aretuza Sousa Takashi Soyano Edgar P. Spalding Imogen Sparkes Raul Antonio Sperotto John Sperry Steven Spoel Robert J. Spreitzer Nathan M. Springer Jens Staal Gary Stacey Ruth Stadler Simon Stael Christopher J. Staiger Claudia Renate Stange Leonie Steinhorst Anna N. Stepanova Martin Steup Chris Still Sophia L. Stone Jens Stougaard Jake M. Stout Benoit St-Pierre Ralf Stracke Ralf Stracke Lucia C. Strader Asa Strand Richard Strasser Johannes Stratmann Peter Streb Richard Strimbeck Paul Struik Anthony Studer Robert M. Stupar Senthil Subramanian Akifumi Sugiyama Michael L. Sullivan Ronan Sulpice Meng-Xiang Sun Tai-Ping Sun Weining Sun Venkatesan Sundaresan Bjorn Sundberg Eva Sundberg Sibum Sung Ramanjulu Sunkar Michael R. Sussman Frances C. Sussmilch Eiji Suzuki Toshiya Suzuki Ales Svatos Steve M. Swain Ranjan Swarup Lee J. Sweetlove Szymon Swiezewski László Szabados Dóra Szakonyi Daniel B. Szymanski Francisco Ramón Tadeo Hideki Takahashi Shunichi Takahashi Yuichiro Takahashi Koji Takahashi Junpei Takano Hiroshi Takatsuji Mizuki Takenaka Hidenori Takeuchi Manuel Talón Kentaro Tamura Li Tan Ayumi Tanaka Weihua Tang Wenqiang Tang Li-zhen Tao Francois Tardieu Nicolas L. Taylor Samuel Taylor Guillaume G.B. Tcherkez Markus Teige Pedro F. Teixeira Frank W. Telewski Ichiro Terashima Matthew J. Terry Mark Tester Christa Testerink Weston Testo Ian Joseph Tetlow Ted Thannhauser Steven M. Theg Frederica L. Theodoulou Guillaume Théroux-Rancourt Gerhard Thiel Dorothea Tholl Stephen G. Thomas Sebastien Thomine Hans Thordal-Christensen Michael Thorpe Feng Tian Jiang Tian Li Tian Shiping Tian Zhixi Tian Mary L. Tierney Axel Tiessen Marcel Tijsterman Mikko Tikkanen Alain F. Tissier Aude Tixier Alyson Tobin Daisuke Todaka Christopher D. Todd Takayuki Tohge Seiichi Toki James G. Tokuhisa Motoki Tominaga Rumi Tominaga-Wada Hongning Tong Christopher N. Topp Jose M. Torres-Ruiz Szilvia Z. Toth Katalin Toth Lam-Son Tran Timothy John Tranbarger Ben Trevaskis Maurizio Trovato Marco Trujillo Joshua Trujillo Chung-Jui Tsai Miltos Tsiantis Tokuji Tsuchiya Katsutoshi Tsuda Hiroyuki Tsuji Masaru Tsujii Hironaka Tsukagoshi Hirokazu Tsukaya Shih-Long Tu Roberto Tuberosa Matthew R. Tucker Paul Tudzynski Robert Turgeon Joseph A. Turner Simon R. Turner David Twell Esa Tyystjarvi Nerea Ubierna Naoyuki Uchida Michael K. Udvardi Minako Ueda Takashi Ueda Yusaku Uga Roman Ulm Masaaki Umeda Taishi Umezawa Nobuyuki Uozumi Breeanna Urbanowicz Bjoern Usadel Olivier Van Aken Frank Van Breusegem Daniel Van Damme Bram Van de Poel Harrold A. van den Burg Alexander R. van der krol Aalt D.J. van Dijk Joost T. van Dongen Kasper van Gelderen Jaimie M. Van Norman Klaas Jan van Wijk Vicki S. Vance Filip J. Vandenbussche Michiel Vandenbussche Radomira Vankova Greg C. Vanlerberghe Serena Varotto Imre Vass Olena K. Vatamaniuk Kees Venema Nathalie Verbruggen Jeanmarie Verchot Jerome Verdier Lionel Verdoucq Wim Vermaas Paul E. Verslues Silvere R.M. Vialet-Chabrand Claudia E. Vickers Andras Viczian Luis Vidali Elizabeth Vierling Usha Vijayraghavan Kris Vissenberg Corina Vlot Laurentius A.C.J. Voesenek Thomas Vogt Christian A. Voigt Albrecht G. von Arnim Shinya Wada Harkamal Walia Berkley J. Walker Elsbeth Lewis Walker Ian S. Wallace Ross Waller Justin W. Walley Chao-Wen Wang Dong Wang Enli Wang Guodong Wang Hsiao-Lin Wang Huanzhong Wang Jia-Wei Wang Kan Wang Ming-Bo Wang MingLi Wang Peng Wang Pengcheng Wang Shucai Wang Wei Wang Xiaohong Wang Xiaowu Wang Xiu-Jie Wang Xuemin Wang Yingxiang Wang Yonghong Wang Yucheng Wang Zhiyong Wang Xin Wang C.J. Rachel Wang Yan Wang John M. Ward Anton Wasson Geoffrey O. Wasteneys Cezary Waszczak Naoharu Watanabe Yuichiro Watanabe Brian M. Waters Danielle Way Alex A.R. Webb Michael Webb Andreas P.M. Weber Wolfram Weckwerth Arne Weiberg Dolf Weijers Daniel Weisz Elina Welchen Ralf Welsch Stephan Wenkel Elizabeth Weretilnyk Randall J. Weselake James Whelan Clinton Whipple Justin G.A. Whitehill Steven A. Whitham Spencer M. Whitney Philip A. Wigge Raymond Wightman Christian Wilhelm Mary E. Williams Ben Williams Valerie Moroz Williamson Robert D. Willows Zoe A. Wilson Scott Wing Lawrence J. Winship Nicolaus von Wiren Sebastian Wolf Daniel Woods Michael Wrzaczek Hen-Ming Wu Keqiang Wu Yajun Wu Yan Wu Yongrui Wu Bernhard Wurzinger Guangmin Xia Rui Xia Xinli Xia Cheng-Bin Xiang Shunyuan Xiao Wenyan Xiao Jun Xiao Daoxin Xie Kabin Xie Deyu Xie Daoxin Xie Shuping Xing Lizhong Xiong Yan Xiong Changcheng Xu Kenong Xu Lin Xu Zheng-Yi Xu Dongqing Xu Arun Yadav Masashi Yamada Kazuko Yamaguchi-Shinozaki Yasusi Yamamoto Yoshiharu Y. Yamamoto Takafumi Yamashino Wataru Yamori Feng Yan Jianbin Yan Dawei Yan Chang-Hsien Yang Chengwei Yang Hongxing Yang Shuhua Yang Wei-Cai Yang Yinong Yang Zhenbiao Yang Zhongnan Yang Marcelo Yanovsky Steven Andrew Yates De Ye Zheng-Hua Ye Trevor Yeats Trevor H Yeats Koichi Yoneyama Gyeong Mee Yoon Takuya Yoshida Kaoru Okamoto Yoshiyama Bin Yu Diqiu Yu Fei Yu Yao-Wu Yuan Dae-Jin Yun Olga A Zabotina Sabine Zachgo Patricia C. Zambryski María Eugenia Zanetti Martijn Van Zanten Viktor Zarsky Jürgen Zeier Assaf Zemach Wei Zeng Ulrike Zentgraf Philipp Zerbe William Zerges Jixian Zhai Bailong Zhang Ben Zhang Cankui Zhang Chunhua Zhang Hairong Zhang Hongxia zhang Jin-Song Zhang Peng Zhang Sheng Zhang Shuqun Zhang Wenhua Zhang Xian Sheng Zhang Xiaoyu Zhang Xiuren Zhang Yan Zhang Yijing Zhang Yong Zhang Yongjiang Zhang Zemin Zhang Meixiang Zhang Ran Zhang Yu Zhang Dazhong Zhao Dongyan Zhao Jian Zhao Yu Zhao Yunde Zhao Zhong Zhao Huanquan Zheng Shao Jian Zheng Xuehua Zhong Chuanen Zhou Dao-Xiu Zhou Jian-Min Zhou Yihua Zhou Youping Zhou Yun Zhou Chuanmei Zhu Danmeng Zhu Jianhua Zhu Xinguang Zhu Yang Zhu Keyan Zhu-Salzman Johan Zicola Laurent Zimmerli Michel Zivy Bethany K. Zolman Jitao Zou Rita Zrenner OUTGOING EDITORS FOR 2017 Associate Editors Anna Amtmann Hong Ma Monitoring Editors Francois Chaumont John Christie Vitaly Citovsky Jeff Doyle Thomas Girke Robert Edwards Anthony Kinney Leon Kochian Ferenc Nagy Shin-Han Shiu Doris Wagner Hong-Wei Xue INCOMING MES FOR 2018 Christian Fankhauser Matt Gilliham Eirini Kaiserli Markus Schwartzlaender Nick Smirnoff Christa Testerink Viktor Zarsky Jianhua Zhang © 2018 American Society of Plant Biologists. All Rights Reserved. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Szymanski, Dan; Bassham, Diane; Munnik, Teun; Sakamoto, Wataru

doi: 10.1104/pp.17.01777pmid: 29317523

Szymanski, Dan; Bassham, Diane; Munnik, Teun; Sakamoto, Wataru

doi: 10.1104/pp.17.01777pmid: 29317523

Plant cells are the fundamental building blocks of growth and development. For each cell type, the size, shape, and mechanical properties of the cell wall are customized for particular physiological functions (Szymanski and Cosgrove, 2009; Winship et al., 2011). The morphogenesis of highly polarized cell types such as trichoblasts and pollen tubes is internally programmed and occurs largely in the absence of a neighbor. Most cell types differentiate in the context of a tissue. Therefore, their growth and shape change can operate at larger spatial scales to influence tissue- and organ-level processes. Because plant cells grow symplastically and are mechanically coupled to their neighboring cells, growth properties and information flow within and between tissues can feed back on and influence cell behaviors. Plant cells are also metabolically specialized. Within a single tissue or organ, cell types can differ greatly in terms of how central metabolism is fueled, the types of metabolites that accumulate, and where in the cell they are stored. Despite the structural and biochemical diversity of different cell types, their cell biology and development can be considered as a similar set of integrated systems-level processes. For example, the metabolic activity and energy status of a cell varies as a function of light levels or developmental stage. The biosynthesis and transport activities of the cytosol and endomembrane systems are integrated with metabolism over time. Cellular systems are also integrated across wide spatial scales. Proteins and protein complexes at the approximately 10- to 100-nm scale can use the cytoskeleton to position organelles and organize the cytoplasm at the approximately 1- to 10-μm spatial scale, to influence cell behaviors. Discovering and unraveling the complexity of these multiscale systems level interactions is a grand challenge in plant research. In recent years, progress has been rapid and is being driven in large part by the widespread use of multichannel quantitative time-lapse imaging. Using this approach, it is possible to create a spatial and temporal coordinate system in which multiple parameters can be measured and cross-correlated, and the effects of mutations or other experimental manipulations can be more deeply analyzed. GROWTH STRATEGIES AND BIOMECHANICS This Focus Issue includes Updates that broadly cover recent discoveries in the field of cell biology and can serve as an important resource for research and classroom instruction. The Updates section of the issue begins with two articles that describe recent discoveries on the biomechanics and control mechanisms of cells that employ either a diffuse growth (Cosgrove, 2018) or tip growth mechanism (Bascom et al., 2018). Quantitative multivariate live cell imaging enables the creation of computational models that predict the mechanics of morphogenesis. In the Update by Bidhendi and Geitmann (2018), the impact of recent papers that use finite element computational modeling to analyze cell morphogenesis is reviewed. CYTOSKELETON: CELLULAR ORGANIZATION AND FUNCTION Growing plant cells use the cytoskeleton to organize the cytoplasm and pattern the cell wall to define the roadways for intracellular transport and influence the mechanical properties of the cell wall. The cortical microtubule cytoskeleton is tightly associated with the plasma membrane and can dictate the patterns of cellulose synthesis in the cell wall (Wasteneys and Ambrose, 2009). Determining how microtubules are patterned at cellular spatial scales is an active area of research and is covered in a research Update in this issue (Elliott and Shaw, 2018b). The Focus Issue also contains a research article describing new types of cortical microtubule arrays that were discovered using spatial and temporal analysis of microtubule polymerization patterns in light-grown hypocotyl cells (Elliott and Shaw, 2018a). Long-term time-lapse imaging and cross-correlation analyses of leaf epidermal pavement cell shape and microtubule organization were used to reveal unexpected temporal and spatial variability of cortical microtubules during the process of lobe formation in pavement cells (Belteton et al., 2018). This issue also contains important new discoveries on the genetic control of microtubule arrays that position the cell division plane (Mir et al., 2018) and a plant-specific Kinesin (KinG) that uses microtubule-based transport to affect intercellular transport of a developmental regulatory protein (Spiegelman et al., 2018). The actin cytoskeleton also is required for the growth of cells that employ either tip or diffuse growth mechanisms. However, it has been difficult to assign particular functions to specific actin arrays in the cell because they are often short-lived, widely distributed, and highly variable with respect to their spatial organization in the cell. The Update by Szymanski and Staiger (2018) focuses on recent discoveries that advance our understanding of how particular actin filament arrays influence growth and organelle clustering in polarized cell types. In this issue, quantitative live cell imaging and biophysical modeling of organelle diffusion was used to demonstrate an important role for actin filaments in locally increasing vesicle concentration at the apex of tip-growing moss protonema (Bibeau et al., 2018). A major research question in the transport and morphogenesis field is centered on understanding how myosin motors are dynamically coupled to actin filament roadways and cargo. The structure and regulation of plant myosin motors is the subject of a timely Update from the Nebenführ lab (Ryan and Nebenführ, 2018). ORGANELLES Endoplasmic Reticulum: Morphology and Function The endomembrane system, plastids, and mitochondria compose an interdependent population of organelles that compartmentalize the biosynthetic and metabolic activities of the cell. The Cell Dynamics Focus Issue contains numerous Updates and research articles that analyze the biogenesis and function of many of the major plant organelles. For example, the endoplasmic reticulum (ER) consists of a highly dynamic network of membrane sheets and tubules that runs throughout the cytoplasm. The synthesis, quality control, and trafficking of various macromolecules originates in the ER, and is therefore critical for plant growth, development, and stress responses (Stefano and Brandizzi, 2018). This Update discusses the dynamics of ER morphology and how this changes under different conditions and throughout development. Recent findings on the function of the cytoskeleton and ER membrane-shaping proteins are presented. In this issue, Sun and Zheng (2018) present a detailed structure-function analysis of the RHD3, an ER membrane fusion factor. The article describes the importance and mechanism of RHD3 dimerization and how it influences ER network structure. Oil or lipid bodies are present in many cell types and are derived from the ER. Control of the size and morphology of oil bodies and the role of oil body membrane proteins in both oil body formation and oil body breakdown during seed germination are described in an Update (Shimada et al., 2018). The interaction between oil bodies and peroxisomes is discussed as an example of the dynamics of organelle interactions. Endocytosis, Early Endosomes, and the trans-Golgi Network Endocytosis is the process by which plasma membrane lipids and proteins, and soluble apoplastic proteins, are packaged and internalized within clathrin-coated vesicles. The cellular control of endocytosis has been controversial in plants, and in this issue Reynolds et al. provide an authoritative Update on the recent advances in this field (Reynolds et al., 2018). Phosphatidylinositol (PI) 3-phosphate (PI3P) is a minor phospholipid that has been implicated in the trafficking of endocytic cargo in plants (Schink et al., 2013). The trans-Golgi network (TGN) not only is a sorting station for cargo en route to the plasma membrane, but also functions as an early endosome, receiving endocytic vesicles from the plasma membrane. Therefore, the TGN is the convergence site for multiple trafficking pathways. Cargo from each of these pathways has to be correctly sorted at the TGN for transport on to the appropriate destination. An Update discusses recent work on the biogenesis of the TGN from the Golgi, the specialization of distinct TGN domains for sorting via different pathways, and role of the TGN in cell wall synthesis and cytokinesis (Rosquete et al., 2018). Chloroplasts and Mitochondria Chloroplasts and mitochondria are critical organelles for photosynthesis and respiration, respectively. This Focus Issue highlights the cell biology of their division and their membrane connectivities. The Update from Chen et al. describes our recent understanding of chloroplast division machineries (Chen et al., 2018). In bacteria, cell division involves only a single FtsZ protein that forms polymers that mark and generate the division site. In this issue, the Osteryoung group demonstrates that photosynthetic organisms encode a second FtsZ isoform and that plastid division involves the polymerization of FtsZ heteropolymers during chloroplast division (TerBush et al., 2018). Another interesting aspect of chloroplasts is their unique protruding structures, termed “stromules,” that extend from the outer chloroplast membrane. The imaging techniques to visualize and analyze stromules in the context of interorganelle communication and environmental response is reviewed (Hanson and Hines, 2018). Although chloroplasts respond to external abiotic signals, such as blue light-regulated chloroplast relocation in mesophylls, how chloroplast morphologies are affected by biotic signals is poorly understood. A research article by Jin et al. (2018) moves the field forward by using electron tomography and 3D construction of chloroplasts in virus-infected Nicotiana cells, and reveals cytoplasmic invaginations at outer envelopes as the site of virus replication. Similar to chloroplasts, mitochondria also proliferate by division of preexisting organelles. However, they are much more dynamic because of a highly active “fusion” system that chloroplasts do not possess. This fusion system plays a fundamental role in exchanging mitochondrial genetic information, as mounting evidence now indicates that a majority of mitochondria exist without DNA. Arimura (2018) reviews mitochondrial fusion and fission systems in plant cells, with emphasis on the interorganelle mitochondrial DNA exchange. Cellular Responses to Stress Fundamentally important cellular processes such as gene expression (see Updates Daszkowska-Golec [2018] and Chantarachot and Bailey-Serres [2018]) and vesicle trafficking respond to adverse conditions. Autophagy is a vesicle trafficking process in which organelles and macromolecules are transported in autophagosomes to the vacuole for degradation, often as a response to stress. An Update by Soto-Burgos et al. discusses recent advances in our understanding of how the autophagy pathway is activated by a variety of stress conditions, and also the mechanisms by which membrane remodeling and dynamics contributes to the formation of autophagosomes (Soto-Burgos et al., 2018). Peroxisomes are specialized for oxidation reactions during fatty acid catabolism, reactive oxygen species scavenging, and photorespiration. Peroxisomes not only emerge from budding ER membranes but also proliferate by division. Kao et al. provide a comprehensive Update in which peroxisome biogenesis as well as metabolic reactions are reviewed (Kao et al., 2018). Environmental stress is known to cause proliferation of peroxisomes, but the mechanisms by which this occurs are unknown. An article by Frick and Strader shows that MAP KINASE17 (MPK17) controls the number of peroxisomes via the peroxisome division factor PMD1, with an increase in peroxisome number in mpk17 mutants (Frick and Strader, 2018). Both MPK17 and PMD1 were shown to be required for proliferation of peroxisomes upon salt stress, in a process requiring actin polymerization. MPK17 therefore defines a new pathway for the regulation of peroxisome number, particularly as a response to stress conditions. The nucleus usually receives attention because of its importance in controlling gene expression in response to developmental or positional cues. The nuclear envelopes and nuclear pore complexes are important components of the nuclear periphery and help to define nuclear structure. Knowledge about how the structure of the nucleus is generated and how it responds to developmental and biotic/abiotic signals is described in the Update by Groves et al. (2018). The issue also contains an article that analyzes how nuclear position is controlled during root hair differentiation (Nakamura et al., 2018). CLOSING REMARKS Sophisticated live cell imaging pipelines are being used to quantitatively analyze organelle, cytoskeletal, and cell wall systems (often in parallel). These imaging-centric approaches, empowered further by solid genetics and biochemistry, are providing mechanistic insight into how cells control their division and morphogenesis across wide spatial and temporal scales. The field is learning more about how cellular systems allow the plant to respond adaptively to abiotic and biotic stress. Collectively, these discoveries are providing a knowledge base that can enable the engineering of improved crops with specified architectural traits or stress tolerances. There are likely to be increasing opportunities to use different types of computational modeling techniques to define specific targets and efficient strategies for cellular engineering (Zuñiga et al., 2018). As more of the proteins and cellular activities that control cellular phenotypes become known, there are new opportunities to use rapidly evolving superresolution and light sheet microscopy technologies (see Update by Komis et al. [2018]). Protein complexes and cellular dynamics can be analyzed at spatial and temporal resolutions far beyond what has been previously achieved. An important remaining challenge is centered on learning how small GTPases and other signaling molecules coordinate diverse cellular activities during plant growth and development (see Update by Feiguelman et al. [2018]). In the coming years, it will be interesting to see the extent to which new imaging technologies and computational methods accelerate the rate at which the control modules of the cell are revealed. LITERATURE CITED Arimura S ( 2018 ) Fission and fusion of plant mitochondria, and genome maintenance . Plant Physiol 176 : 152 – 161 Google Scholar Crossref Search ADS PubMed WorldCat Bascom CS Jr, Hepler PK, Bezanilla M ( 2018 ) Interplay between ions, the cytoskeleton, and cell wall properties during tip growth . Plant Physiol 176 : 28 – 40 Google Scholar Crossref Search ADS PubMed WorldCat Belteton SA , Sawchuk MG, Donohoe BS, Scarpella E, Szymanski DB ( 2018 ) Reassessing the roles of PIN proteins and anticlinal microtubules during pavement cell morphogenesis . Plant Physiol 176 : 432 – 449 Google Scholar Crossref Search ADS PubMed WorldCat Bibeau JP , Kingsley JL, Furt F, Tüzel E, Vidali L ( 2018 ) F-actin mediated focusing of vesicles at the cell tip is essential for polarized growth . Plant Physiol 176 : 352 – 363 Google Scholar Crossref Search ADS PubMed WorldCat Bidhendi AJ , Geitmann A ( 2018 ) Finite element modeling of shape changes in plant cells . Plant Physiol 176 : 41 – 56 Google Scholar Crossref Search ADS PubMed WorldCat Chantarachot T , Bailey-Serres J ( 2018 ) Polysomes, stress granules, and processing bodies: a dynamic triumvirate controlling cytoplasmic mRNA fate and function . Plant Physiol 176 : 254 – 269 Google Scholar Crossref Search ADS PubMed WorldCat Chen C , MacCready JS, Ducat DC, Osteryoung KW ( 2018 ) The molecular machinery of chloroplast division . Plant Physiol 176 : 138 – 151 Google Scholar Crossref Search ADS PubMed WorldCat Cosgrove DJ ( 2018 ) Diffuse growth of plant cell walls . Plant Physiol 176 : 16 – 27 Google Scholar Crossref Search ADS PubMed WorldCat Daszkowska-Golec A ( 2018 ) Emerging roles of the nuclear cap-binding complex in abiotic stress responses . Plant Physiol 176 : 242 – 253 Google Scholar Crossref Search ADS PubMed WorldCat Elliott A , Shaw SL ( 2018a ) Microtubule array patterns have a common underlying architecture in hypocotyl cells . Plant Physiol 176 : 307 – 325 Google Scholar Crossref Search ADS WorldCat Elliott A , Shaw SL ( 2018b ) Update: plant cortical microtubule arrays . Plant Physiol 176 : 94 – 105 Google Scholar Crossref Search ADS WorldCat Feiguelman G , Fu Y, Yalovsky S ( 2018 ) ROP GTPases structure-function and signaling pathways . Plant Physiol 176 : 57 – 79 Google Scholar Crossref Search ADS PubMed WorldCat Frick EM , Strader LC ( 2018 ) Kinase MPK17 and the peroxisome division factor PMD1 influence salt-induced peroxisome proliferation . Plant Physiol 176 : 340 – 351 Google Scholar Crossref Search ADS PubMed WorldCat Groves NR , Biel AM, Newman-Griffis AH, Meier I ( 2018 ) Dynamic changes of plant nuclear organization in response to environmental and developmental signals . Plant Physiol 176 : 230 – 241 Google Scholar Crossref Search ADS PubMed WorldCat Hanson MR , Hines KM ( 2018 ) Stromules: probing formation and function . Plant Physiol 176 : 128 – 137 Google Scholar Crossref Search ADS PubMed WorldCat Jin X , Jiang Z, Zhang K, Wang P, Cao X, Yue N, Wang X, Zhang X, Li Y, Li D, et al. ( 2018 ) Three-dimensional analysis of chloroplast structures associated with virus infection . Plant Physiol 176 : 282 – 294 Google Scholar Crossref Search ADS PubMed WorldCat Kao YT , Gonzalez KL, Bartel B ( 2018 ) Peroxisome function, biogenesis, and dynamics in plants . Plant Physiol 176 : 162 – 171 Google Scholar Crossref Search ADS PubMed WorldCat Komis G , Novák D, Ovečka M, Šamajova O, Šamaj J ( 2018 ) Advances in imaging plant cell dynamics . Plant Physiol 176 : 80 – 93 Google Scholar Crossref Search ADS PubMed WorldCat Mir R , Morris VH, Buschmann H, Rasmussen CG ( 2018 ) Division plane orientation defects revealed by a synthetic double mutant phenotype . Plant Physiol 176 : 418 – 431 Google Scholar Crossref Search ADS PubMed WorldCat Nakamura M , Claes AR, Grebe T, Hermkes R, Viotti C, Ikeda Y, Grebe M ( 2018 ) Auxin and ROP GTPase signaling of polar nuclear migration in root epidermal hair cells . Plant Physiol 176 : 378 – 391 Google Scholar Crossref Search ADS PubMed WorldCat Reynolds GD , Wang C, Pan J, Bednarek SY ( 2018 ) Inroads into internalization: five years of endocytic exploration . Plant Physiol 176 : 208 – 218 Google Scholar Crossref Search ADS PubMed WorldCat Rosquete MR , Davis DJ, Drakakaki G ( 2018 ) The plant trans-Golgi network: not just a matter of distinction . Plant Physiol 176 : 187 – 198 Google Scholar Crossref Search ADS PubMed WorldCat Ryan JM , Nebenführ A ( 2018 ) Update on myosin motors: molecular mechanisms and physiological functions . Plant Physiol 176 : 119 – 127 Google Scholar Crossref Search ADS PubMed WorldCat Schink KO , Raiborg C, Stenmark H ( 2013 ) Phosphatidylinositol 3-phosphate, a lipid that regulates membrane dynamics, protein sorting and cell signalling . BioEssays 35 : 900 – 912 Google Scholar PubMed OpenURL Placeholder Text WorldCat Shimada TL , Hayashi M, Hara-Nishimura I ( 2018 ) Membrane dynamics and multiple functions of oil bodies in seeds and leaves . Plant Physiol 176 : 199 – 207 Google Scholar Crossref Search ADS PubMed WorldCat Soto-Burgos J , Zhuang X, Jiang L, Bassham DC ( 2018 ) Dynamics of autophagosome formation . Plant Physiol 176 : 219 – 229 Google Scholar Crossref Search ADS PubMed WorldCat Spiegelman Z , Lee CM, Gallagher KL ( 2018 ) KinG is a plant-specific kinesin that regulates both intra- and intercellular movement of SHORT-ROOT . Plant Physiol 176 : 392 – 405 Google Scholar Crossref Search ADS PubMed WorldCat Stefano G , Brandizzi F ( 2018 ) Advances in plant ER architecture and dynamics . Plant Physiol 176 : 178 – 186 Google Scholar Crossref Search ADS PubMed WorldCat Sun J , Zheng H ( 2018 ) Efficient ER fusion requires a dimerization and a C-terminal tail mediated membrane anchoring of RHD3 . Plant Physiol 176 : 406 – 417 Google Scholar Crossref Search ADS PubMed WorldCat Szymanski DB , Cosgrove DJ ( 2009 ) Dynamic coordination of cytoskeletal and cell wall systems during plant cell morphogenesis . Curr Biol 19 : R800 – R811 Google Scholar Crossref Search ADS PubMed WorldCat Szymanski D , Staiger CJ ( 2018 ) The actin cytoskeleton: functional arrays for cytoplasmic organization and cell shape control . Plant Physiol 176 : 106 – 118 Google Scholar Crossref Search ADS PubMed WorldCat TerBush AD , MacCready JS, Chen C, Ducat DC, Osteryoung KW ( 2018 ) Conserved dynamics of chloroplast cytoskeletal FtsZ proteins across photosynthetic lineages . Plant Physiol 176 : 295 – 306 Google Scholar Crossref Search ADS PubMed WorldCat Wasteneys GO , Ambrose JC ( 2009 ) Spatial organization of plant cortical microtubules: close encounters of the 2D kind . Trends Cell Biol 19 : 62 – 71 Google Scholar Crossref Search ADS PubMed WorldCat Winship LJ , Obermeyer G, Geitmann A, Hepler PK ( 2011 ) Pollen tubes and the physical world . Trends Plant Sci 16 : 353 – 355 Google Scholar Crossref Search ADS PubMed WorldCat Zuñiga C , Levering J, Antoniewicz MR, Guarnieri MT, Betenbaugh MJ, Zengler K ( 2018 ) Predicting dynamic metabolic demands in the photosynthetic eukaryote Chlorella vulgaris . Plant Physiol 176 : 450 – 462 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 Address correspondence to [email protected]. www.plantphysiol.org/cgi/doi/10.1104/pp.17.01777 © 2018 American Society of Plant Biologists. All Rights Reserved. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Cosgrove, Daniel J.

doi: 10.1104/pp.17.01541pmid: 29138349

The primary wall of a growing cell is a versatile, subtle, and dynamic structure, with unique properties and functions in the life of the plant (Burton et al., 2010). When a cell grows, its wall stretches irreversibly as the cell enlarges in volume. Cells can start and stop this process quickly, in less than a minute in some cases, revealing that the molecular processes underlying irreversible wall expansion are dynamically controlled. Such dynamic behavior may be mediated, at least in part, by changes in wall pH (Hager, 2003; Barbez et al., 2017), which modulates the wall-loosening action of expansins (Cosgrove, 2015) and potentially other wall-modifying agents. Wall pH in turn is dynamically modulated by plasma membrane H+-ATPase activity (Haruta et al., 2014, 2015) and other processes in the wall. Because growing cell walls are thin and in close physical contact with plasma membranes, wall pH can be rapidly modulated (Bibikova et al., 1998; Monshausen et al., 2007; Barbez et al., 2017). As a result of the pH-dependent activity of expansins, the growing cell wall behaves like a “smart material”—one whose properties (extensibility in this case) reversibly and rapidly change with environment (e.g. pH). Slower changes in wall structure that influence the wall’s ability to expand also occur as part of the natural course of cell development, e.g. as cells are displaced through the elongation zone of a stem (Phyo et al., 2017), or in response to external perturbations, e.g. Sahaf and Sharon (2016). These slower changes may include changes in mechanics, such as wall stiffening, and in the density or accessibility of sites where expansins or other proteins can loosen the wall. The wall itself is synthesized in a team effort: mobile cellulose synthesis complexes (Paredez et al., 2006; Li et al., 2016b) produce long, thin, strong, stiff cellulose microfibrils at the cell surface, while matrix polysaccharides and glycoproteins are deposited to the cell surface via the secretory apparatus (Zhu et al., 2015; Kim and Brandizzi, 2016). The cytoskeleton guides the wall synthesis machinery to supply wall components to appropriate locations on the cell surface (Szymanski and Staiger, 2017), where the components assemble to form an organized, mechanically strong structure that can withstand the in-plane tensile forces generated by the outward push of cell turgor pressure yet is able to expand in a controlled manner. The structural requirements for orderly expansion of the cell wall are not well defined at this time. Moreover, except with the possible exception of tip-growing cells (Dumais et al., 2006; Rojas et al., 2011), synthesis, secretion, and wall assembly are only distantly coupled to the wall extension process itself. For instance, cellulose synthesis in carbon-limited Arabidopsis (Arabidopsis thaliana) hypocotyls was temporally distinct from cell expansion (Ivakov et al., 2017); likewise, wall deposition did not keep up with cell expansion in dark-grown hypocotyls, resulting in substantial wall thinning (Refrégier et al., 2004). On the other hand, gradients in wall thickness in the growing trichome of Arabidopsis precisely matched predictions for mechanical stability of the wall, implying good coordination between local wall deposition and expansion (Yanagisawa et al., 2015). We still have much to learn about how plant cells build a stable yet extensible wall. Open in new tabDownload slide Open in new tabDownload slide When the normal molecular assembly of the cell wall is disturbed, for instance, by mutations that affect synthesis of cellulose (Fagard et al., 2000), xyloglucan (Cavalier et al., 2008), or pectic polysaccharides (Mouille et al., 2007), cell expansion may be disrupted in unpredictable ways. Genetic results suggest that errors in wall assembly trigger surface receptor-like kinases such as FERONIA that may act as sensors of cell wall integrity (CWI; Humphrey et al., 2007; Höfte, 2015) and mechanosensation (Hamant and Haswell, 2017). The ensuing responses, which include production of reactive oxygen species and inactivation of the plasma membrane H+-ATPase, likely give rise to some of the complex phenotypes that originate from rather simple modifications of wall polysaccharides (Voxeur and Höfte, 2016). For example, growth defects stemming from mutation of a cellulose synthase gene in Arabidopsis were partially suppressed by mutation of THESEUS1 (Hématy et al., 2007), another member of the same receptor kinase family as FERONIA (Cheung and Wu, 2011; Li et al., 2016a). Evidently, CWI responses compound and confound the direct effects of cell wall defects. Defects in pectin metabolism appear particularly prone to trigger CWI responses that activate the brassinosteroid pathway, leading to diverse growth phenotypes (Wolf et al., 2012, 2014). On the other hand, FERONIA and its extracellular peptide ligand (“rapid alkalinization factor”) are also required for normal root growth and auxin responses (Haruta et al., 2014; Shih et al., 2014; Velasquez et al., 2016; Barbez et al., 2017). Cell expansion thus appears to be intimately linked to these wall sensor pathways in ways we are only beginning to fathom. This Update focuses on the growing cell wall, in particular, the structural, mechanical, and physicochemical processes underlying irreversible wall enlargement during diffuse cell growth. Diffuse growth refers to surface expansion occurring on entire facets of cell walls, for instance, the side walls of elongating cells in the body of a growing root or stem. Diffuse growth may occur with or without a directional bias, which depends partly on wall structure and partly on patterns of mechanical stress in the wall (Baskin and Jensen, 2013). Its intensity may vary along a cell wall surface and on different cell wall facets. For instance, side walls of a hypocotyl cell may elongate rapidly, whereas its end walls may not enlarge much at all (Peaucelle et al., 2015). In the jigsaw-puzzle-like pavement cells of the Arabidopsis leaf epidermis, a complex pattern of local wall surface expansion occurs in the periclinal (outer epidermal) wall as well as in the anticlinal (side) walls (Szymanski, 2014; Armour et al., 2015). These complex expansion patterns have been linked to cytoskeletal dynamics within the cell and to spatial patterns of tensile stress (Szymanski and Cosgrove, 2009; Zhang et al., 2013; Sampathkumar et al., 2014a). Diffuse growth is the dominant pattern for most cells in the plant body and is traditionally contrasted with tip growth, for instance, in pollen tubes and root hairs, where surface expansion is localized to limited regions of the hemispherical tip (Campàs et al., 2012; Sanati Nezhad and Geitmann, 2013; Velasquez et al., 2016). Despite differences in spatial patterning, tip growth may involve some of the same cell wall processes as occur in diffuse growth, but in a more intense, spatially localized manner. This point is supported by the stunted elongation of root hairs in plants with genetic lesions in expansin genes specifically expressed in root hairs (Yu et al., 2011). Although expansins have been characterized largely in the context of diffuse growth, they also are necessary for root hair growth. Leaf trichomes in Arabidopsis are fiber-like cells with a conical shape that arises from highly anisotropic diffuse growth; its tip-biased gradient in surface expansion has been related to spatial gradients in cell diameter, wall stress, and wall thickness (Yanagisawa et al., 2015). In this Update, I begin with a review of the biophysical basis of cell wall growth, followed by our changing concepts of the role and interactions of cell wall components that compose the cell wall, and end with recent insights from atomic force microscopy (AFM), showing the details of microfibril organization and motions during wall enlargement. WALL STRESS RELAXATION DRIVES CELL GROWTH What happens when a cell grows? Cell volume increases as a result of water uptake, and wall surface area enlarges irreversibly by local separation of cell wall components (Cosgrove, 2016b). The phrase “turgor-driven growth” (or its variants) is often used to describe this process, but this phrasing can be misleading if it is taken to imply that growth is simply a mechanical stretching of an inherently pliant cell wall, like stretching a piece of putty. To understand this point better, it is useful to distinguish different patterns of cell wall stretching in response to an applied tensile force (Fig. 1). Elastic deformation is by definition reversible and instantaneous upon applied force. Small wall deformations are typically elastic, whereas after large, rapid, transient deformations, many cell walls do not fully return to their initial size. The irreversible part is a plastic deformation, which occurs when the wall is stretched beyond a yield point. When force is applied to a wall in a sustained manner, the stretch (“strain” in engineering terms) is partly elastic and partly plastic or partly viscoelastic and partly viscoplastic, depending on the time scales involved (Boudaoud, 2010; Moulia, 2013). These latter two terms refer to time-dependent deformations. Without sustained cell wall loosening, a constant force applied to a wall typically results in a time-dependent strain that approaches a nearly steady value in a few minutes, depending on the sample and its strain history (Hohl and Schopfer, 1992). Such deformation is a result of the polymeric nature of plant cell walls, but the exact structural basis for these mechanical properties is largely unknown and needs further theoretical development. Figure 1. Open in new tabDownload slide Schematic comparisons of different strain patterns of cell walls (top) stretched with a uniaxial force (bottom). A, Purely elastic (reversible) strain. Cell walls often display a retarded elasticity because wall polymer motions are not instantaneous. B, A combination of elastic and plastic strain. Beyond the yield point strain is partly elastic and partly plastic. C, Viscoelastic and viscoplastic strains. Polymer motions require time to reach equilibrium. D, Wall loosening results in sustained time-dependent extension (creep). Figure 1. Open in new tabDownload slide Schematic comparisons of different strain patterns of cell walls (top) stretched with a uniaxial force (bottom). A, Purely elastic (reversible) strain. Cell walls often display a retarded elasticity because wall polymer motions are not instantaneous. B, A combination of elastic and plastic strain. Beyond the yield point strain is partly elastic and partly plastic. C, Viscoelastic and viscoplastic strains. Polymer motions require time to reach equilibrium. D, Wall loosening results in sustained time-dependent extension (creep). These purely mechanical responses of walls differ in an essential way from the sustained wall expansion that occurs during cell growth (Cosgrove, 2016b; Zhang et al., 2017), which depends on continuous loosening by expansins or other wall-loosening agents (Cosgrove, 2016a). Wall loosening results in wall stress relaxation that drives cell growth. The significance of wall stress relaxation for plant growth was first recognized by Ray et al. (1972) and later solidified by detailed theory and experimental results (Cosgrove, 1993a, 1993b; Ortega, 2017). As a result of the complex interactions of cell wall components, there are various distinctive ways in which cell walls may become mechanically softer (meaning more easily deformed by mechanical force), but they do not necessarily result in an increase in wall relaxation and growth, and they therefore do not qualify as wall loosening processes (see below and Table I). For instance, with rare exceptions (Yuan et al., 2001), lytic enzymes may soften walls, but they do not stimulate cell growth or cause long-term cell wall extension (creep; Ruesink, 1969; Cosgrove and Durachko, 1994; Fleming et al., 1997). On the other hand, α-expansins cause stress relaxation and prolonged enlargement of cell walls, but they lack wall lytic activity and they do not soften the wall, as measured by tensile tests (see figure 8 in Yuan et al. [2001]). These are remarkable facts that seem counterintuitive to expectations based on conventional models of cell walls (Carpita and Gibeaut, 1993). To illustrate the point in another way: Xyloglucan-deficient hypocotyl walls from the xxt1,xxt2 mutant of Arabidopsis are more compliant (more easily stretched) in tensile tests compared with wild-type walls, yet xxt1,xxt2 hypocotyls grow more slowly than the wild type (Xiao et al., 2016). Moreover, despite their greater mechanical compliance, the xxt1,xxt2 walls extend more slowly in creep tests, exhibit less stress relaxation, and are less responsive to α-expansins compared with wild-type walls (Park and Cosgrove, 2012a). Another example illustrates the flip side of the coin: Low temperature strongly reduced cell wall expansion in Chara cells, but wall elasticity was hardly affected (Proseus et al., 2000). Many other examples have been documented where mechanical tests do not reliably report on the growth properties of the cell wall (Cosgrove, 2016b). Brief explanation of terms related to wall mechanics and growth properties, as used here Table I. Brief explanation of terms related to wall mechanics and growth properties, as used here Term . Meaning . Extensibility General term for the ability of the cell wall to grow; in other contexts, this is the coefficient relating growth rate to turgor pressure (Cosgrove, 1993b); not elasticity and not a purely mechanical property, as it depends on wall loosening Loosening Molecular process causing wall stress relaxation, resulting in water uptake and cell growth; it confers irreversibility to wall strains; expansins are well-recognized as wall-loosening agents Softening A process that makes the wall more deformable to mechanical force Weakening A process that reduces the force or energy need to break walls Elasticity A measure of how readily the cell wall changes shape in response to a transient mechanical force Plasticity A measure of the irreversible component of wall deformation in response to a transient mechanical force Compliance The slope for strain/stress curves; it is the reciprocal of modulus or stiffness Term . Meaning . Extensibility General term for the ability of the cell wall to grow; in other contexts, this is the coefficient relating growth rate to turgor pressure (Cosgrove, 1993b); not elasticity and not a purely mechanical property, as it depends on wall loosening Loosening Molecular process causing wall stress relaxation, resulting in water uptake and cell growth; it confers irreversibility to wall strains; expansins are well-recognized as wall-loosening agents Softening A process that makes the wall more deformable to mechanical force Weakening A process that reduces the force or energy need to break walls Elasticity A measure of how readily the cell wall changes shape in response to a transient mechanical force Plasticity A measure of the irreversible component of wall deformation in response to a transient mechanical force Compliance The slope for strain/stress curves; it is the reciprocal of modulus or stiffness Open in new tab Table I. Brief explanation of terms related to wall mechanics and growth properties, as used here Term . Meaning . Extensibility General term for the ability of the cell wall to grow; in other contexts, this is the coefficient relating growth rate to turgor pressure (Cosgrove, 1993b); not elasticity and not a purely mechanical property, as it depends on wall loosening Loosening Molecular process causing wall stress relaxation, resulting in water uptake and cell growth; it confers irreversibility to wall strains; expansins are well-recognized as wall-loosening agents Softening A process that makes the wall more deformable to mechanical force Weakening A process that reduces the force or energy need to break walls Elasticity A measure of how readily the cell wall changes shape in response to a transient mechanical force Plasticity A measure of the irreversible component of wall deformation in response to a transient mechanical force Compliance The slope for strain/stress curves; it is the reciprocal of modulus or stiffness Term . Meaning . Extensibility General term for the ability of the cell wall to grow; in other contexts, this is the coefficient relating growth rate to turgor pressure (Cosgrove, 1993b); not elasticity and not a purely mechanical property, as it depends on wall loosening Loosening Molecular process causing wall stress relaxation, resulting in water uptake and cell growth; it confers irreversibility to wall strains; expansins are well-recognized as wall-loosening agents Softening A process that makes the wall more deformable to mechanical force Weakening A process that reduces the force or energy need to break walls Elasticity A measure of how readily the cell wall changes shape in response to a transient mechanical force Plasticity A measure of the irreversible component of wall deformation in response to a transient mechanical force Compliance The slope for strain/stress curves; it is the reciprocal of modulus or stiffness Open in new tab Despite this “inconvenient truth” [apologies to Al Gore (2006)], wall elasticity is frequently taken to be synonymous with cell wall growth properties. Sometimes this is a matter of convenience—elasticity is relatively straightforward to incorporate into simulations of growth (Fayant et al., 2010; Huang et al., 2015)—and numerous methods have been devised in recent years to measure elasticity of tissues and cells with mechanical devices (e.g. Routier-Kierzkowska et al., 2012; Nezhad et al., 2013; Sanati Nezhad et al., 2013; Beauzamy et al., 2015b; Vogler et al., 2015; Mosca et al., 2017). To be fair, there are indeed cases where elasticity roughly correlates with cell growth, as discussed below. Such correlation may indicate a change in wall structure that actually contributes to altered growth by amplifying wall stress relaxation (see Cosgrove [2016b]), or may be entirely coincidental, resulting from structural changes independent of wall extensibility. Elastic changes may also be a consequence of the altered growth. For instance, it has long been known that auxin treatment results in increased wall compliances in many cases (Heyn, 1932; Edelmann and Kohler, 1995; Braybrook, 2017). However, the onset of auxin-induced growth is fast and precedes the gradual change in wall mechanics (Cleland, 1984), which may reflect longer-term changes in wall structure induced by auxin or faster growth itself. Another factor in the confusion between elasticity and extensibility may be the fuzzy definition of terms such as wall loosening, softening, and weakening (Table I). We do not have an established vocabulary to distinguish the many facets of wall properties. As discussed here, wall loosening refers to a shift or cut of a load-bearing part of the wall, relaxing tensile stress in the whole wall and simultaneously reducing cell turgor, which is the Newtonian counterbalance to wall stress. The reduction in turgor enables passive water uptake by osmosis, which elastically stretches the cell wall, restoring turgor. Such loosening-dependent polymer movement might be called a chemorheological flow (Dumais, 2013; Moulia, 2013; Cosgrove, 2016b). However, this term usually implies a chemical change in covalent bonding within the cell wall, which does not appear to be necessary for cell wall creep, i.e. α-expansin facilitates cell wall creep without evidence of hydrolysis or other covalent modification of wall polymers (Cosgrove, 2015, 2016a). I use wall “softening” to denote a process that changes wall stiffness, without the implication that the wall can grow more quickly. I use wall “weakening” to refer to cases where the wall breaking strength is reduced. Thus, xxt1,xxt2 hypocotyls are weaker than the wild type because their breaking strength is reduced (Cavalier et al., 2008). Despite being softer and weaker, xxt1,xxt2 hypocotyls are less extensible in assays of growth and cell wall creep. The key conclusion is that elasticity reports on wall structure, not the dynamic relaxation processes that determine wall extensibility and that drive cell wall growth. With the definitions used here, a stretchy rubber band has significant elasticity but no extensibility. CELL WALL MODELS NEED FURTHER REFINEMENT AND TESTING This physical framework, in which wall stress relaxation initiates cell growth and couples water uptake with wall expansion, leaves numerous molecular details unresolved. What is the molecular nature of wall loosening? Which wall components of the wall are the targets of wall loosening, and which components limit the ensuing polymer motions (e.g. spreading of cellulose microfibrils)? Are there multiple ways to induce wall relaxation and loosen the wall? What are the molecular bases of wall elasticity and plasticity? The answers to these questions require an accurate model of cell wall structure and a deep understanding of the molecular basis of wall enlargement. Notwithstanding textbook models, this remains a major unmet challenge. For many years, xyloglucan was imagined to function as a load-bearing tether linking well-separated cellulose microfibrils (Carpita and Gibeaut, 1993; Albersheim et al., 2011), with pectins functioning as a compliant, hydrated, gel-like matrix between the microfibrils. This concept entailed predictions of wall mechanics that were largely untested. Recent results, however, call for reevaluation of the roles of cellulose, xyloglucan, and pectins in wall structure and growth (Cosgrove, 2016b). For instance, extensive xyloglucanase treatment of isolated cell walls from cucumber (Cucumis sativus) and Arabidopsis did not induce cell wall extension (creep) or increase mechanical compliances (stress/strain behavior in tensile tests), even though at least half of the xyloglucan was removed (Park and Cosgrove, 2012b). On the other hand, low concentrations of bifunctional endoglucanases (able to cut both xyloglucan and cellulose) induced cell wall creep and mechanical softening (increase in tensile compliances). The extent of wall hydrolysis needed to induce these biomechanical actions was very small. Enigmatically, the combination of a xyloglucan-specific endoglucanase with a cellulose-specific endoglucanase did not mimic the bifunctional enzymes in their action on cell wall creep. To explain this enigma, we proposed that the bifunctional enzymes hydrolyze relatively inaccessible load-bearing junctions between cellulose microfibrils where xyloglucan and noncrystalline cellulose are entwined (Fig. 2). Digestion of these nexus points, dubbed “biomechanical hotspots,” releases some of the load-bearing junctions between the microfibrils, enabling stress relaxation and irreversible microfibril movements. Limited accessibility and kinetics account for the failure of combined xyloglucan-specific and cellulose-specific enzymes to cause cell wall creep. Figure 2. Open in new tabDownload slide Conceptual depiction of structural features of primary cell walls. Cellulose microfibrils are represented as thick rods with hydrophobic (blue) and hydrophilic faces (orange). Xyloglucan (green) is found in solvated, coiled conformations and in extended conformations bound to the hydrophobic faces of cellulose, based on Zheng et al. (2017b). It is also depicted as entrapped between microfibrils. Pectins (yellow) are represented as coiled structures that fill the space between microfibrils and bind to the hydrophilic surfaces (based on solid-state NMR results). Microfibrils are bundled by direct contacts and at junctions where cellulose and xyloglucan intertwine. The red arrows point to cellulose-xyloglucan-cellulose junctions that are sites of wall loosening by bifunctional endoglucanases. This depiction is a synthesis based on the most recent results from AFM, FESEM, solid-state NMR, and mechanics. Figure 2. Open in new tabDownload slide Conceptual depiction of structural features of primary cell walls. Cellulose microfibrils are represented as thick rods with hydrophobic (blue) and hydrophilic faces (orange). Xyloglucan (green) is found in solvated, coiled conformations and in extended conformations bound to the hydrophobic faces of cellulose, based on Zheng et al. (2017b). It is also depicted as entrapped between microfibrils. Pectins (yellow) are represented as coiled structures that fill the space between microfibrils and bind to the hydrophilic surfaces (based on solid-state NMR results). Microfibrils are bundled by direct contacts and at junctions where cellulose and xyloglucan intertwine. The red arrows point to cellulose-xyloglucan-cellulose junctions that are sites of wall loosening by bifunctional endoglucanases. This depiction is a synthesis based on the most recent results from AFM, FESEM, solid-state NMR, and mechanics. The origin of these proposed junctions is uncertain. One possibility is that they form spontaneously by physical entrapment and self-assembly as cellulose and xyloglucan are deposited to the cell surface. Coordination of vesicle secretion with cellulose synthesis might facilitate their formation. Another possibility is that hotspots are formed enzymatically. Recent reports have identified enzymes (xyloglucan endotransglucosylase/hydrolase) that can cut noncrystalline cellulose and ligate xyloglucan onto the cellulose end (Hrmova et al., 2007; Simmons et al., 2015; Shinohara et al., 2017). The specific activity of these enzymes in performing such hetero-transglucosylations is very low, so the biological significance of such reactions is uncertain. Further work is needed to uncover the origin of biomechanical hotspots. The hotspot concept gains support from a study that used a novel solid-state NMR strategy to characterize the target of expansin binding in complex walls from Arabidopsis seedlings (Wang et al., 2013). With use of a sensitivity-boosting technique called dipolar nuclear polarization to enhance 1H-13C cross-polarization, nuclear magnetic spins were selectively filtered through 15N,13C-labeled expansin protein to 13C in wall components within close proximity to expansin. The results showed that expansin binds to cellulose with a slightly different chemical shift than the bulk of the cellulose, indicating cellulose chains that are packed together differently than most of the cellulose. Moreover, there was evidence of xyloglucan in close proximity to expansin. These NMR characteristics resemble those that might be expected for the biomechanical junctions targeted by wall-loosening bifunctional endoglucanases (Park and Cosgrove, 2012b). Additional NMR characterization found that the expansin-binding sites are on the surface of cellulose microfibrils, yet are relatively distant from water, consistent with limited accessibility (Wang et al., 2016b). The concept of wall structure emerging from these results emphasizes the importance of direct connectivity between cellulose microfibrils, as opposed to previous concepts where microfibrils were depicted as well separated by matrix polymers and connected only via tethers. As described below, AFM reveals the organization of cellulose microfibrils to be a network of laterally bundled microfibrils rather than a collection of well-spaced microfibrils connected only by matrix, as traditionally depicted in textbook models of primary cell walls. Another cellulose-related topic emerging from recent work is the potential significance of the cross-sectional shape of cellulose microfibrils, which influences the proportion of hydrophilic and hydrophobic faces on the microfibril (Newman et al., 2013; Cosgrove, 2014; Wang and Hong, 2016). This issue is important for cell wall models because of the potential for these two faces to interact differently with matrix components. The hydrophilic faces of cellulose microfibrils are populated by hydroxyl groups extending from the sides of the Glc residues, whereas the hydrophobic faces are those that expose the Glc rings with their nonpolar −CH groups. X-ray crystallography studies show that expansins bind the hydrophobic face of cellulose chains (Georgelis et al., 2012). Computational studies indicate that xyloglucan likewise preferentially binds to the hydrophobic face of cellulose (Zhao et al., 2014). This computational result is supported by a recent field emission scanning electron microscopy (FESEM)-based study showing that xyloglucan indeed covers the hydrophobic faces of cellulose microfibrils in onion (Allium cepa) cell walls (Zheng et al., 2017b). Complementing these results, recent experimental and computational studies indicate that substituted xylans in secondary cell walls may bind the hydrophilic faces of cellulose (Simmons et al., 2016; Grantham et al., 2017; Pereira et al., 2017). Primary cell walls generally contain only small amounts of xylan, e.g. Zablackis et al. (1995), with the notable exception of cell walls from grass species (Carpita, 1996). Nevertheless, a recent study characterized a xylan in primary walls of Arabidopsis (Mortimer et al., 2015) that might selectively bind to the hydrophilic faces of microfibrils. Because β-expansin was recently shown to target xylans in grass cell walls (Wang et al., 2016a), it seems plausible that α- and β-expansins may loosen the connections between different faces of cellulose microfibrils in the wall. Direct cellulose-cellulose contacts may be prevalent in primary walls, but whether such interactions are mediated via their hydrophilic or hydrophobic faces is uncertain and indeed may differ for primary and secondary cell walls. Some studies propose that microfibrils aggregate via contacts of their hydrophilic faces (Ding et al., 2012; Oehme et al., 2015), whereas an AFM analysis of microfibril patterns in onion cell walls suggested that microfibril bundling occurs via the hydrophobic faces of microfibrils (Zhang et al., 2016). In Arabidopsis hypocotyls of the xyloglucan-deficient xxt1,xxt2 mutant (Xiao et al., 2016), cellulose microfibrils were more aligned and closely packed than in the wild type, an indication that xyloglucan may promote dispersion of microfibrils within a lamella. It is also possible that these changes in cellulose organization are a consequence of CWI responses to wall defects. PECTINS As the idea of a direct cellulose network within primary cell walls has gained support, pectins have also attracted new attention as potential modifiers of wall enlargement. The case has been most cogently argued for pollen tubes (Parre and Geitmann, 2005; Rojas et al., 2011; Sanati Nezhad et al., 2014), where pectins dominate wall structure, and for the giant-celled alga Chara corallina (Boyer, 2016). In Arabidopsis cell walls, results from multidimensional solid-state NMR indicate extensive noncovalent cellulose-pectin interactions (Wang et al., 2012, 2015). This is surprising because such interactions are not observed in binding studies in vitro (Zykwinska et al., 2008a, 2008b). A recent study of cell wall properties along the axial growth gradient of the Arabidopsis stem gives additional clues (Phyo et al., 2017). Pectins in the apical (faster-growing and softer) region of the stem are more mobile, more hydrated, more esterified, and more branched compared with pectins in the lower (more slowly growing and stiffer) region of the stem. How these correlated structural changes in pectin properties influence pectin-cellulose interactions, cell wall mechanics, and growth needs further testing. In contrast, a very different concept of pectin was proposed in a study that characterized a proteoglycan with covalently linked pectin and xylan domains (Tan et al., 2013). This structure is reminiscent of the early macromolecular model of primary cell walls by Keegstra et al. (1973). The potential role of such proteoglycans in wall structure, mechanics, and growth remains to be evaluated. In another vein, a series of elegant experiments with clear but perplexing results implicate localized de-esterification of homogalacturonan as a signature event in the auxin-induced patterning of the shoot apical meristem of Arabidopsis, resulting in elastically softer regions of the meristem surface, as measured by microindentation techniques, where leaf primordia emerge (Peaucelle et al., 2008, 2011; Braybrook and Peaucelle, 2013). Experiments with the auxin-transport pin1 mutant and genes encoding pectin methyl esterase inhibitor proteins suggest that auxin patterning of the shoot apical meristem requires pectin de-esterification. The correlation of de-esterified pectin with softer meristem regions is perplexing because, in the broader context of cell wall properties, pectin de-esterification is commonly thought to result in stiffer, not softer walls, as a result of increased ability for calcium-mediated cross-linking of homogalacturonan. For instance, in the hemispherical tip of growing pollen tubes, de-esterified homogalacturonan is associated with stiffer walls and cessation of wall expansion (Geitmann and Parre, 2004; Sanati Nezhad et al., 2014). Likewise, pectin de-esterification is associated with the decline in the growth rate and increased wall stiffness along the apical-to-basal gradient of growing stems (Goldberg et al., 1986; Phyo et al., 2017). Moreover, wall loosening by expansin is hindered in the basal regions of growing stems where the extent of de-esterified pectin is high (Cosgrove, 1996), and this hindrance may be partially reversed by removal of pectins and calcium (Zhao et al., 2008). Hocq et al. (2017) have questioned the concept of wall stiffening by calcium cross-linking of pectins, yet older results with calcium chelators yielded a nuanced conclusion: that calcium cross-links are indeed load-bearing but are not broken during acid-induced (expansin-mediated) wall loosening (Virk and Cleland, 1990). Likewise, imaging with AFM shows that addition and removal of calcium reversibly stiffens pectins on the surface of onion epidermal cell walls (Zhang et al., 2016), yet parallel experiments show little effect of calcium on wall extension in vitro. Thus, more research is needed to clarify the perplexing observations about pectin esterification, calcium cross-linking, wall softening, auxin responses, and cell growth in the meristem. Are CWI responses complicating this story? The perplexing results about pectin esterification reported for the shoot apical meristem raise the following question: Is cell wall enlargement in the meristem regulated in the same way as in subapical zones of rapid cell enlargement, or do walls in the meristem follow a different set of rules? Indeed, we know rather little about cell wall structure and wall extensibility in meristematic regions, other than the limited information inferred from immunohistochemistry (Yang et al., 2016) and osmomechanical probing of cell elasticity (Kierzkowski et al., 2012; Nakayama et al., 2012; Routier-Kierzkowska and Smith, 2013). In a recent multifaceted study of the swollen shoot apical meristems from the clavata3-2 mutant of Arabidopsis, wall composition was found to consist of approximately 30% cellulose, approximately 26% pectin, and 15% xyloglucan (Yang et al., 2016). This is unremarkable, as it is similar to the wall composition of whole Arabidopsis seedlings (White et al., 2014). Older studies also bear on the question of cell enlargement mechanisms in meristems: Local application of α-expansin protein to the surface of the shoot apical meristem resulted in an outgrowth resembling early stages of a leaf primordium (Fleming et al., 1999), and more pronounced outgrowth resulted from transient induction of an α-expansin on the flanks of the meristem (Pien et al., 2001), indicating cell walls in the meristem are sensitive to the loosening action of α-expansin. Moreover, α-expansin is endogenously expressed at the site of incipient leaf primordia before primordium outgrowth (Reinhardt et al., 1998), evidence that modulation of cell wall enlargement is similar to that of other plant tissues. Allowing for the high frequency of dividing cells in meristems and the unique wall synthesis machinery involved in cell plate formation during cell division (Gu et al., 2016), current results indicate meristem walls are similar in composition and growth mechanisms as documented in rapidly elongating cells that emerge from meristems. Thus, the enigmatic function of pectin de-esterification in the meristem remains an open question. INSIGHTS FROM AFM OF EPIDERMAL CELL WALLS Plant developmental biologists are probably most familiar with AFM from studies that have used the device to indent the surface of growing tissues to evaluate local stiffness (e.g. Peaucelle et al., 2011; Milani et al., 2014; Sampathkumar et al., 2014a). This is a complex topic beyond the scope of this review, but readers are referred to reviews that assess the varied approaches and interpretations of these stiffness measurements (Milani et al., 2013; Mosca et al., 2017). AFM-based stiffness maps of the shoot apical meristem have been compared with maps of cell shape, cell division, auxin flow, gene expression, and cytoskeletal patterns (Nakayama et al., 2012; Robinson et al., 2013; de Reuille et al., 2014; Sassi et al., 2014). These studies contribute to sophisticated models of meristem morphogenesis and phyllotaxis in which wall stress, mechanics, enlargement, and the cytoskeleton play interacting roles (Nakayama et al., 2012; Kierzkowski et al., 2013; Sampathkumar et al., 2014b), concepts rooted in the pioneering efforts of Paul Green to understand the biophysics of meristem dynamics (Green et al., 1996). AFM can also be used to image cell wall surfaces at sufficiently high resolution to detect individual cellulose microfibrils (approximately 3 nm diameter; Fig. 3, A and B), revealing cellulose organization in unprecedented detail (Zhang et al., 2014). Note that fluorescence microscopy, even super resolution versions, lacks the resolution needed to see individual microfibrils in the complex and dense fibrillar network of cell walls. Transmission electron microscopy generally gives limited information about microfibril organization in cell walls, with the exception of the replica/shadowing method (McCann et al., 1990), which has been superseded by FESEM. FESEM has excellent resolving power for microfibril detection (Fujita and Wasteneys, 2014; Zheng et al., 2017a), but to attain high resolution, the wall must be dehydrated, which means wall polymers may become distorted or shift position as the water is removed. In contrast, AFM can be carried out under water, allowing imaging of walls in a near-native state. The surface topology can be measured with nm resolution and simultaneously probed mechanically to measure an indentation modulus (resistance to surface deformation). Figure 3. Open in new tabDownload slide AFM images showing the arrangement of cellulose microfibrils and matrix on the inner surface of the periclinal wall of onion epidermis. A, Large-scale (2 × 2 μm) peak force error map showing microfibril bundling and cross-lamellate organization of microfibrils. Image adapted from Zhang et al. (2014). B, Close-up of boxed region in A. Single microfibrils are seen to bundle into groups of two of more. Some microfibril details are obscured by relatively stiff matrix. C, Two-color merged image based on a height map (red), which highlights microfibrils, and a modulus map (green), which indicates stiffness (resistance to indentation). The area outlined in white contains regions of stiff matrix (bright green) closely associated with microfibrils. Other matrix regions are dark, indicating they are soft. D, Two-color merged image based on a height map (red, predominantly microfibrils) and a deformation map (green, predominantly matrix). Regions of soft matrix between microfibrils are obvious in this image. Images adapted from Zhang et al. (2016). E, Modulus map of onion wall in a relaxed state (small axial force); the predominance of blue color indicates low resistance to indentation. F, Modulus map of the same wall region upon application of axial force, stretching the wall in the direction indicated by the arrow. The predominance of red shows that microfibrils have been pulled taut by the axial force, indicating they are load bearing, as are matrix components, though to a lesser extent. Images adapted from Zhang et al. (2017). Figure 3. Open in new tabDownload slide AFM images showing the arrangement of cellulose microfibrils and matrix on the inner surface of the periclinal wall of onion epidermis. A, Large-scale (2 × 2 μm) peak force error map showing microfibril bundling and cross-lamellate organization of microfibrils. Image adapted from Zhang et al. (2014). B, Close-up of boxed region in A. Single microfibrils are seen to bundle into groups of two of more. Some microfibril details are obscured by relatively stiff matrix. C, Two-color merged image based on a height map (red), which highlights microfibrils, and a modulus map (green), which indicates stiffness (resistance to indentation). The area outlined in white contains regions of stiff matrix (bright green) closely associated with microfibrils. Other matrix regions are dark, indicating they are soft. D, Two-color merged image based on a height map (red, predominantly microfibrils) and a deformation map (green, predominantly matrix). Regions of soft matrix between microfibrils are obvious in this image. Images adapted from Zhang et al. (2016). E, Modulus map of onion wall in a relaxed state (small axial force); the predominance of blue color indicates low resistance to indentation. F, Modulus map of the same wall region upon application of axial force, stretching the wall in the direction indicated by the arrow. The predominance of red shows that microfibrils have been pulled taut by the axial force, indicating they are load bearing, as are matrix components, though to a lesser extent. Images adapted from Zhang et al. (2017). We used AFM to characterize cellulose microfibril organization of the outer (periclinal) wall of onion scales (the fleshy leaf of the bulb). Epidermal peels from onion scales have been used in numerous studies to connect tissue-level mechanics, growth, and net cellulose orientation (Wilson et al., 2000; Hepworth and Bruce, 2004; Suslov et al., 2009; Beauzamy et al., 2015a). In previous studies, whole epidermal layers were prepared with intact (living) cells. We developed an alternative procedure to prepare epidermal strips in which the outer (periclinal) wall tears away from the rest of the cell. This exposes the inner (most recently synthesized) surface for imaging by AFM and provides a simpler material for mechanical studies (a sheet of outer epidermal walls rather than a layer of intact turgid cells with complex architecture). Like the outer epidermal walls in many plant organs, including Arabidopsis hypocotyls (Crowell et al., 2011), the epidermal wall in onion is thick (2 or more μm, depending on which scale is used) and has a cross-lamellate construction. In AFM images, individual microfibrils in the surface lamella are seen to form a network with single microfibrils merging into and out of bundled regions where two or more microfibrils are laterally aligned and in close contact (Fig. 3, A and B; Zhang et al., 2016). Microfibrils are arranged roughly in a common direction in each lamella, and microfibril orientation shifts abruptly by 30° to 90° between adjacent lamellae, producing a wall comprised of many, highly anisotropic lamellae in a wide range of orientations. The result of this cross-lamellate structure is that the whole wall has much weaker net structural anisotropy than the individual lamellae that make up the wall. By combining height maps with modulus maps and deformation maps, we can visualize microfibrils in the context of rigid and soft regions of the matrix (Fig. 3, C and D). Moreover, by stretching the wall, we can detect microfibril motions and detect which components become more resistant to indentation, an indication that they bear some of the tensile force (Fig. 3, E and F). This cross-lamellate construction has important implications for the patterns of microfibril separation during cell growth. It is commonly accepted that cell walls expand preferentially in the direction at right angle to the net direction of cellulose alignment (Baskin, 2005; Suslov et al., 2009). Multicellular tissues present additional structural complications beyond the scope of this review (Crowell et al., 2010; Baskin and Jensen, 2013). For a cell wall with net transverse orientation, it is easy to imagine that the distance between microfibrils increases as the cell elongates axially. This is commonly illustrated by analogy with the way a wound spring elongates (like a Slinky toy). But what happens in a cross-lamellate wall where cellulose is organized in lamellae that are aligned in many orientations, axially, transversely, and at a spectrum of angles between these two orthogonal directions? Specifically, how do microfibrils move in lamellae where cellulose microfibrils are aligned parallel to the direction of maximal growth? In considering this question, I assume that microfibrils are too stiff to stretch appreciably and too strong to break, and also assume that adjacent lamellae do not slip past each other. With these assumptions, it seems that microfibrils in these axially aligned lamellae must have a mechanism of side-by-side gliding or axial shearing (Fig. 4). If axial shearing of this type requires more force than lateral separation of microfibrils in transversely aligned lamellae, then axial shearing between microfibrils aligned in the direction of cell growth may limit the rate of cell wall enlargement. This reasoning puts the focus of attention on the molecular nature of the lateral associations between microfibrils and the patterns of microfibril motions during wall expansion. We know rather little about this aspect of primary cell wall growth. Figure 4. Open in new tabDownload slide Predicted patterns of microfibril movement in a cross-lamellate cell wall stretched uniaxially. Shown is cellulose alignment in single lamellae before (left) and after (right) stretching in the vertical direction. Axial elongation is accompanied by transverse shrinkage during elastic uniaxial extensions. A, In lamellae with cellulose oriented transverse to the direction of stretch, the axial distance between microfibrils will increase and cellulose microfibrils will bend or kink in the transverse direction. B, For lamellae with cellulose oriented at approximately 45°, microfibril angle will shift in the axial direction and distance between microfibrils will be reduced. C, For lamellae with cellulose oriented in the same direction as the axial stretch, microfibrils will become straighter, more closely packed, and will undergo side-by-side sliding. Figure is adapted from Zhang et al. (2016). Figure 4. Open in new tabDownload slide Predicted patterns of microfibril movement in a cross-lamellate cell wall stretched uniaxially. Shown is cellulose alignment in single lamellae before (left) and after (right) stretching in the vertical direction. Axial elongation is accompanied by transverse shrinkage during elastic uniaxial extensions. A, In lamellae with cellulose oriented transverse to the direction of stretch, the axial distance between microfibrils will increase and cellulose microfibrils will bend or kink in the transverse direction. B, For lamellae with cellulose oriented at approximately 45°, microfibril angle will shift in the axial direction and distance between microfibrils will be reduced. C, For lamellae with cellulose oriented in the same direction as the axial stretch, microfibrils will become straighter, more closely packed, and will undergo side-by-side sliding. Figure is adapted from Zhang et al. (2016). These inferences about microfibril movements gain support from a recent study in which the nanoscale movements of cellulose microfibrils were directly monitored by AFM (Zhang et al., 2017). The cell wall was extended in a well-defined series of extensions that included elastic and plastic strains imposed by axial force as well as time-dependent creep induced by treatment with an endoglucanase with wall-loosening activity. For lamellae in which the microfibrils were oriented at 30° to 60° off the axis of applied tensile force, mechanical stretching resulted in passive reorientation of microfibrils in the direction of stretch, as predicted by multinet growth models (Preston, 1982). In addition, examples of axial shearing and lateral separation were observed, providing direct evidence for the microfibril movements inferred above from general considerations. When cell wall creep was induced by application of a wall-loosening endoglucanase, remarkably different patterns of microfibril movements were observed compared with those during elastic and plastic deformations. This difference was attributed to changes in microfibril connectivity, i.e. selective loosening at hotspots by endoglucanase action. Thus, the stretching of a wound spring does not seem an apt analogy for how microfibrils move during cell wall growth. This AFM study documents an example where microfibril movements motivated by applied force differed from those that occurred when wall expansion was induced by wall loosening. The microfibrils in differently aligned lamellae displayed the large diversity of microfibril movements that are required for enlargement of cross-lamellate walls. It is possible that different patterns of microfibril movement involve different matrix components, e.g. xyloglucans, xylans, pectins, or even water alone, and are mediated by different wall-loosening agents. Open in new tabDownload slide Open in new tabDownload slide PERSPECTIVE As new tools and new approaches have been applied to investigate cell growth, it has become evident that our conventional model of the growing cell wall falls short. Xyloglucan seems to have rather different functional roles than those hypothesized in conventional depictions of the growing cell wall of the past 40 years. New evidence for the role of pectins in wall structure and morphogenesis are tantalizing, yet raise new questions and the biological responses evoked by CWI sensors complicate the interpretation of cell wall mutants. Finally, the new appreciation of the complexity of cellulose organization in the growing wall presents opportunities for rethinking the molecular control of diffuse growth. Experimental systems that enable studies of both pectin and cellulose networks are needed for future integration of these emerging ideas. LITERATURE CITED Albersheim P , Darvill A, Roberts K, Sederoff R, Staehelin A ( 2011 ) Plant Cell Walls. Garland , New York Google Scholar Google Preview OpenURL Placeholder Text WorldCat COPAC Armour WJ , Barton DA, Law AM, Overall RL ( 2015 ) Differential growth in periclinal and anticlinal walls during lobe formation in Arabidopsis cotyledon pavement cells . Plant Cell 27 : 2484 – 2500 Google Scholar Crossref Search ADS PubMed WorldCat Barbez E , Dünser K, Gaidora A, Lendl T, Busch W ( 2017 ) Auxin steers root cell expansion via apoplastic pH regulation in Arabidopsis thaliana . Proc Natl Acad Sci USA 114 : E4884 – E4893 Google Scholar Crossref Search ADS PubMed WorldCat Baskin TI ( 2005 ) Anisotropic expansion of the plant cell wall . Annu Rev Cell Dev Biol 21 : 203 – 222 Google Scholar Crossref Search ADS PubMed WorldCat Baskin TI , Jensen OE ( 2013 ) On the role of stress anisotropy in the growth of stems . J Exp Bot 64 : 4697 – 4707 Google Scholar Crossref Search ADS PubMed WorldCat Beauzamy L , Derr J, Boudaoud A ( 2015 a ) Quantifying hydrostatic pressure in plant cells by using indentation with an atomic force microscope . Biophys J 108 : 2448 – 2456 Google Scholar Crossref Search ADS PubMed WorldCat Beauzamy L , Louveaux M, Hamant O, Boudaoud A ( 2015 b ) Mechanically, the shoot apical meristem of Arabidopsis behaves like a shell inflated by a pressure of about 1 MPa . Front Plant Sci 6 : 1038 Google Scholar Crossref Search ADS PubMed WorldCat Bibikova TN , Jacob T, Dahse I, Gilroy S ( 1998 ) Localized changes in apoplastic and cytoplasmic pH are associated with root hair development in Arabidopsis thaliana . Development 125 : 2925 – 2934 Google Scholar PubMed OpenURL Placeholder Text WorldCat Boudaoud A ( 2010 ) An introduction to the mechanics of morphogenesis for plant biologists . Trends Plant Sci 15 : 353 – 360 Google Scholar Crossref Search ADS PubMed WorldCat Boyer JS ( 2016 ) Enzyme-less growth in Chara and terrestrial plants . Front Plant Sci 7 : 866 Google Scholar Crossref Search ADS PubMed WorldCat Braybrook SA ( 2017 ) Analyzing cell wall elasticity after hormone treatment: an example using tobacco BY-2 cells and auxin . Methods Mol Biol 1497 : 125 – 133 Google Scholar Crossref Search ADS PubMed WorldCat Braybrook SA , Peaucelle A ( 2013 ) Mechano-chemical aspects of organ formation in Arabidopsis thaliana: the relationship between auxin and pectin . PLoS One 8 : e57813 Google Scholar Crossref Search ADS PubMed WorldCat Burton RA , Gidley MJ, Fincher GB ( 2010 ) Heterogeneity in the chemistry, structure and function of plant cell walls . Nat Chem Biol 6 : 724 – 732 Google Scholar Crossref Search ADS PubMed WorldCat Campàs O , Rojas E, Dumais J, Mahadevan L ( 2012 ) Strategies for cell shape control in tip-growing cells . Am J Bot 99 : 1577 – 1582 Google Scholar Crossref Search ADS PubMed WorldCat Carpita NC ( 1996 ) Structure and biogenesis of the cell walls of grasses . Annu Rev Plant Physiol Plant Mol Biol 47 : 445 – 476 Google Scholar Crossref Search ADS PubMed WorldCat Carpita NC , Gibeaut DM ( 1993 ) 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 Google Scholar Crossref Search ADS PubMed WorldCat Cavalier DM , Lerouxel O, Neumetzler L, Yamauchi K, Reinecke A, Freshour G, Zabotina OA, Hahn MG, Burgert I, Pauly M, et al. ( 2008 ) Disrupting two Arabidopsis thaliana xylosyltransferase genes results in plants deficient in xyloglucan, a major primary cell wall component . Plant Cell 20 : 1519 – 1537 Google Scholar Crossref Search ADS PubMed WorldCat Cheung AY , Wu HM ( 2011 ) THESEUS 1, FERONIA and relatives: a family of cell wall-sensing receptor kinases? Curr Opin Plant Biol 14 : 632 – 641 Google Scholar Crossref Search ADS PubMed WorldCat Cleland RE ( 1984 ) The Instron technique as a measure of immediate-past wall extensibility . Planta 160 : 514 – 520 Google Scholar Crossref Search ADS PubMed WorldCat Cosgrove DJ ( 1993 a ) How do plant cell walls extend? Plant Physiol 102 : 1 – 6 Google Scholar Crossref Search ADS PubMed WorldCat Cosgrove DJ ( 1993 b ) Wall extensibility: its nature, measurement and relationship to plant cell growth . New Phytol 124 : 1 – 23 Google Scholar Crossref Search ADS PubMed WorldCat Cosgrove DJ ( 1996 ) Plant cell enlargement and the action of expansins . BioEssays 18 : 533 – 540 Google Scholar Crossref Search ADS PubMed WorldCat Cosgrove DJ ( 2014 ) Re-constructing our models of cellulose and primary cell wall assembly . Curr Opin Plant Biol 22 : 122 – 131 Google Scholar Crossref Search ADS PubMed WorldCat Cosgrove DJ ( 2015 ) Plant expansins: diversity and interactions with plant cell walls . Curr Opin Plant Biol 25 : 162 – 172 Google Scholar Crossref Search ADS PubMed WorldCat Cosgrove DJ ( 2016 a ) Catalysts of plant cell wall loosening . F1000Res 5 : F1000 Faculty Rev-119 Google Scholar Crossref Search ADS PubMed WorldCat Cosgrove DJ ( 2016 b ) Plant cell wall extensibility: connecting plant cell growth with cell wall structure, mechanics, and the action of wall-modifying enzymes . J Exp Bot 67 : 463 – 476 Google Scholar Crossref Search ADS PubMed WorldCat Cosgrove DJ , Durachko DM ( 1994 ) Autolysis and extension of isolated walls from growing cucumber hypocotyls . J Exp Bot 45 : 1711 – 1719 Google Scholar Crossref Search ADS PubMed WorldCat Crowell EF , Gonneau M, Vernhettes S, Höfte H ( 2010 ) Regulation of anisotropic cell expansion in higher plants . C R Biol 333 : 320 – 324 Google Scholar Crossref Search ADS PubMed WorldCat Crowell EF , Timpano H, Desprez T, Franssen-Verheijen T, Emons AM, Höfte H, Vernhettes S ( 2011 ) Differential regulation of cellulose orientation at the inner and outer face of epidermal cells in the Arabidopsis hypocotyl . Plant Cell 23 : 2592 – 2605 Google Scholar Crossref Search ADS PubMed WorldCat de Reuille PB , Robinson S, Smith RS ( 2014 ) Quantifying cell shape and gene expression in the shoot apical meristem using MorphoGraphX . Methods Mol Biol 1080 : 121 – 134 Google Scholar Crossref Search ADS PubMed WorldCat Ding SY , Liu YS, Zeng Y, Himmel ME, Baker JO, Bayer EA ( 2012 ) How does plant cell wall nanoscale architecture correlate with enzymatic digestibility? Science 338 : 1055 – 1060 Google Scholar Crossref Search ADS PubMed WorldCat Dumais J ( 2013 ) Modes of deformation of walled cells . J Exp Bot 64 : 4681 – 4695 Google Scholar Crossref Search ADS PubMed WorldCat Dumais J , Shaw SL, Steele CR, Long SR, Ray PM ( 2006 ) An anisotropic-viscoplastic model of plant cell morphogenesis by tip growth . Int J Dev Biol 50 : 209 – 222 Google Scholar Crossref Search ADS PubMed WorldCat Edelmann HG , Kohler K ( 1995 ) Auxin increases elastic wall-properties in rye coleoptiles: implications for the mechanism of wall loosening . Physiol Plant 93 : 85 – 92 Google Scholar Crossref Search ADS WorldCat Fagard M , Desnos T, Desprez T, Goubet F, Refregier G, Mouille G, McCann M, Rayon C, Vernhettes S, Höfte H ( 2000 ) PROCUSTE1 encodes a cellulose synthase required for normal cell elongation specifically in roots and dark-grown hypocotyls of Arabidopsis . Plant Cell 12 : 2409 – 2424 Google Scholar PubMed OpenURL Placeholder Text WorldCat Fayant P , Girlanda O, Chebli Y, Aubin CE, Villemure I, Geitmann A ( 2010 ) Finite element model of polar growth in pollen tubes . Plant Cell 22 : 2579 – 2593 Google Scholar Crossref Search ADS PubMed WorldCat Fleming AJ , Caderas D, Wehrli E, McQueen-Mason S, Kuhlemeier C ( 1999 ) Analysis of expansin-induced morphogenesis on the apical meristem of tomato . Planta 208 : 166 – 174 Google Scholar Crossref Search ADS WorldCat Fleming AJ , McQueen-Mason S, Mandel T, Kuhlemeier C ( 1997 ) Induction of leaf primordia by the cell wall protein expansin . Science 276 : 1415 Google Scholar Crossref Search ADS WorldCat Fujita M , Wasteneys GO ( 2014 ) A survey of cellulose microfibril patterns in dividing, expanding, and differentiating cells of Arabidopsis thaliana . Protoplasma 251 : 687 – 698 Google Scholar Crossref Search ADS PubMed WorldCat Geitmann A , Parre E ( 2004 ) The local cytomechanical properties of growing pollen tubes correspond to the axial distribution of structural cellular elements . Sex Plant Reprod 17 : 9 – 16 Google Scholar Crossref Search ADS WorldCat Georgelis N , Yennawar NH, Cosgrove DJ ( 2012 ) 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 Google Scholar Crossref Search ADS PubMed WorldCat Goldberg R , Morvan C, Roland JC ( 1986 ) Composition, properties and localization of pectins in young and mature cells of the mung bean hypocotyl . Plant Cell Physiol 27 : 417 – 429 Google Scholar OpenURL Placeholder Text WorldCat Gore A ( 2006 ) An Inconvenient Truth: The Planetary Emergency of Global Warming and What We Can Do About It. Rodale Books , New York Google Scholar Google Preview OpenURL Placeholder Text WorldCat COPAC Grantham NJ , Wurman-Rodrich J, Terrett OM, Lyczakowski JJ, Stott K, Iuga D, Simmons TJ, Durand-Tardif M, Brown SP, Dupree R, et al. ( 2017 ) An even pattern of xylan substitution is critical for interaction with cellulose in plant cell walls . Nat Plants 3 : 859 – 865 Green PB , Steele CS, Rennich SC ( 1996 ) Phyllotactic patterns: a biophysical mechanism for their origin . Ann Bot (Lond) 77 : 515 – 527 Google Scholar Crossref Search ADS WorldCat Gu F , Bringmann M, Combs JR, Yang J, Bergmann DC, Nielsen E ( 2016 ) Arabidopsis CSLD5 functions in cell plate formation in a cell cycle-dependent manner . Plant Cell 28 : 1722 – 1737 Google Scholar PubMed OpenURL Placeholder Text WorldCat Hager A ( 2003 ) Role of the plasma membrane H+-ATPase in auxin-induced elongation growth: historical and new aspects . J Plant Res 116 : 483 – 505 Google Scholar Crossref Search ADS PubMed WorldCat Hamant O , Haswell ES ( 2017 ) Life behind the wall: sensing mechanical cues in plants . BMC Biol 15 : 59 Google Scholar Crossref Search ADS PubMed WorldCat Haruta M , Gray WM, Sussman MR ( 2015 ) Regulation of the plasma membrane proton pump (H(+)-ATPase) by phosphorylation . Curr Opin Plant Biol 28 : 68 – 75 Google Scholar Crossref Search ADS PubMed WorldCat Haruta M , Sabat G, Stecker K, Minkoff BB, Sussman MR ( 2014 ) A peptide hormone and its receptor protein kinase regulate plant cell expansion . Science 343 : 408 – 411 Google Scholar Crossref Search ADS PubMed WorldCat Hématy K , Sado PE, Van Tuinen A, Rochange S, Desnos T, Balzergue S, Pelletier S, Renou JP, Höfte H ( 2007 ) A receptor-like kinase mediates the response of Arabidopsis cells to the inhibition of cellulose synthesis . Curr Biol 17 : 922 – 931 Google Scholar Crossref Search ADS PubMed WorldCat Hepworth DG , Bruce DM ( 2004 ) Relationships between primary plant cell wall architecture and mechanical properties for onion bulb scale epidermal cells . J Texture Stud 35 : 586 – 602 Google Scholar Crossref Search ADS WorldCat Heyn AJN ( 1932 ) Further investigations of the mechanism of cell elongation and the properties of the cell wall in connection with elongation . Protoplasma 19 : 78 – 97 Google Scholar Crossref Search ADS WorldCat Hocq L , Pelloux J, Lefebvre V ( 2017 ) Connecting homogalacturonan-type pectin remodeling to acid growth . Trends Plant Sci 22 : 20 – 29 Google Scholar Crossref Search ADS PubMed WorldCat Höfte H ( 2015 ) The yin and yang of cell wall integrity control: brassinosteroid and FERONIA signaling . Plant Cell Physiol 56 : 224 – 231 Google Scholar Crossref Search ADS PubMed WorldCat Hohl M , Schopfer P ( 1992 ) Physical extensibility of maize coleoptile cell walls: apparent plastic extensibility is due to elastic hysteresis . Planta 187 : 498 – 504 Google Scholar PubMed OpenURL Placeholder Text WorldCat Hrmova M , Farkas V, Lahnstein J, Fincher GB ( 2007 ) A barley xyloglucan xyloglucosyl transferase covalently links xyloglucan, cellulosic substrates, and (1,3;1,4)-beta-D-glucans . J Biol Chem 282 : 12951 – 12962 Google Scholar Crossref Search ADS PubMed WorldCat Huang R , Becker AA, Jones IA ( 2015 ) A finite strain fibre-reinforced viscoelasto-viscoplastic model of plant cell wall growth . J Eng Math 95 : 121 – 154 Google Scholar Crossref Search ADS WorldCat Humphrey TV , Bonetta DT, Goring DR ( 2007 ) Sentinels at the wall: cell wall receptors and sensors . New Phytol 176 : 7 – 21 Google Scholar Crossref Search ADS PubMed WorldCat Ivakov A , Flis A, Apelt F, Fünfgeld M, Scherer U, Stitt M, Kragler F, Vissenberg K, Persson S, Suslov D ( 2017 ) Cellulose synthesis and cell expansion are regulated by different mechanisms in growing Arabidopsis hypocotyls . Plant Cell 29 : 1305 – 1315 Google Scholar Crossref Search ADS PubMed WorldCat Keegstra K , Talmadge KW, Bauer WD, Albersheim P ( 1973 ) The structure of plant cell walls. III. A model of the walls of suspension-cultured sycamore cells based on the interconnections of the macromolecular components . Plant Physiol 51 : 188 – 197 Google Scholar Crossref Search ADS PubMed WorldCat Kierzkowski D , Lenhard M, Smith R, Kuhlemeier C ( 2013 ) Interaction between meristem tissue layers controls phyllotaxis . Dev Cell 26 : 616 – 628 Google Scholar Crossref Search ADS PubMed WorldCat Kierzkowski D , Nakayama N, Routier-Kierzkowska AL, Weber A, Bayer E, Schorderet M, Reinhardt D, Kuhlemeier C, Smith RS ( 2012 ) Elastic domains regulate growth and organogenesis in the plant shoot apical meristem . Science 335 : 1096 – 1099 Google Scholar Crossref Search ADS PubMed WorldCat Kim SJ , Brandizzi F ( 2016 ) The plant secretory pathway for the trafficking of cell wall polysaccharides and glycoproteins . Glycobiology 26 : 940 – 949 Google Scholar Crossref Search ADS PubMed WorldCat Li C , Wu HM, Cheung AY ( 2016 a ) FERONIA and her pals: functions and mechanisms . Plant Physiol 171 : 2379 – 2392 Google Scholar Crossref Search ADS PubMed WorldCat Li S , Bashline L, Zheng Y, Xin X, Huang S, Kong Z, Kim SH, Cosgrove DJ, Gu Y ( 2016 b ) Cellulose synthase complexes act in a concerted fashion to synthesize highly aggregated cellulose in secondary cell walls of plants . Proc Natl Acad Sci USA 113 : 11348 – 11353 Google Scholar Crossref Search ADS PubMed WorldCat McCann MC , Wells B, Roberts K ( 1990 ) Direct visualization of cross-links in the primary plant cell wall . J Cell Sci 96 : 323 – 334 Google Scholar OpenURL Placeholder Text WorldCat Milani P , Braybrook SA, Boudaoud A ( 2013 ) Shrinking the hammer: micromechanical approaches to morphogenesis . J Exp Bot 64 : 4651 – 4662 Google Scholar Crossref Search ADS PubMed WorldCat Milani P , Mirabet V, Cellier C, Rozier F, Hamant O, Das P, Boudaoud A ( 2014 ) Matching patterns of gene expression to mechanical stiffness at cell resolution through quantitative tandem epifluorescence and nanoindentation . Plant Physiol 165 : 1399 – 1408 Google Scholar Crossref Search ADS PubMed WorldCat Monshausen GB , Bibikova TN, Messerli MA, Shi C, Gilroy S ( 2007 ) Oscillations in extracellular pH and reactive oxygen species modulate tip growth of Arabidopsis root hairs . Proc Natl Acad Sci USA 104 : 20996 – 21001 Google Scholar Crossref Search ADS PubMed WorldCat Mortimer JC , Faria-Blanc N, Yu X, Tryfona T, Sorieul M, Ng YZ, Zhang Z, Stott K, Anders N, Dupree P ( 2015 ) An unusual xylan in Arabidopsis primary cell walls is synthesised by GUX3, IRX9L, IRX10L and IRX14 . Plant J 83 : 413 – 426 Google Scholar Crossref Search ADS PubMed WorldCat Mosca G , Sapala A, Strauss S, Routier-Kierzkowska AL, Smith RS ( 2017 ) On the micro-indentation of plant cells in a tissue context . Phys Biol 14 : 015003 Google Scholar Crossref Search ADS PubMed WorldCat Mouille G , Ralet MC, Cavelier C, Eland C, Effroy D, Hématy K, McCartney L, Truong HN, Gaudon V, Thibault JF, et al. ( 2007 ) Homogalacturonan synthesis in Arabidopsis thaliana requires a Golgi-localized protein with a putative methyltransferase domain . Plant J 50 : 605 – 614 Google Scholar Crossref Search ADS PubMed WorldCat Moulia B ( 2013 ) Plant biomechanics and mechanobiology are convergent paths to flourishing interdisciplinary research . J Exp Bot 64 : 4617 – 4633 Google Scholar Crossref Search ADS PubMed WorldCat Nakayama N , Smith RS, Mandel T, Robinson S, Kimura S, Boudaoud A, Kuhlemeier C ( 2012 ) Mechanical regulation of auxin-mediated growth . Curr Biol 22 : 1468 – 1476 Google Scholar Crossref Search ADS PubMed WorldCat Newman RH , Hill SJ, Harris PJ ( 2013 ) Wide-angle x-ray scattering and solid-state nuclear magnetic resonance data combined to test models for cellulose microfibrils in mung bean cell walls . Plant Physiol 163 : 1558 – 1567 Google Scholar Crossref Search ADS PubMed WorldCat Nezhad AS , Naghavi M, Packirisamy M, Bhat R, Geitmann A ( 2013 ) Quantification of the Young’s modulus of the primary plant cell wall using Bending-Lab-On-Chip (BLOC) . Lab Chip 13 : 2599 – 2608 Google Scholar Crossref Search ADS PubMed WorldCat Oehme DP , Doblin MS, Wagner J, Bacic A, Downton MT, Gidley MJ ( 2015 ) Gaining insight into cell wall cellulose macrofibril organisation by simulating microfibril adsorption . Cellulose 22 : 3501 – 3520 Google Scholar Crossref Search ADS WorldCat Ortega JKE ( 2017 ) Dimensionless number is central to stress relaxation and expansive growth of the cell wall . Sci Rep 7 : 3016 Google Scholar Crossref Search ADS PubMed WorldCat Paredez AR , Somerville CR, Ehrhardt DW ( 2006 ) Visualization of cellulose synthase demonstrates functional association with microtubules . Science 312 : 1491 – 1495 Google Scholar Crossref Search ADS PubMed WorldCat Park YB , Cosgrove DJ ( 2012 a ) Changes in cell wall biomechanical properties in the xyloglucan-deficient xxt1/xxt2 mutant of Arabidopsis . Plant Physiol 158 : 465 – 475 Google Scholar Crossref Search ADS PubMed WorldCat Park YB , Cosgrove DJ ( 2012 b ) A revised architecture of primary cell walls based on biomechanical changes induced by substrate-specific endoglucanases . Plant Physiol 158 : 1933 – 1943 Google Scholar Crossref Search ADS PubMed WorldCat Parre E , Geitmann A ( 2005 ) Pectin and the role of the physical properties of the cell wall in pollen tube growth of Solanum chacoense . Planta 220 : 582 – 592 Google Scholar Crossref Search ADS PubMed WorldCat Peaucelle A , Braybrook SA, Le Guillou L, Bron E, Kuhlemeier C, Höfte H ( 2011 ) Pectin-induced changes in cell wall mechanics underlie organ initiation in Arabidopsis . Curr Biol 21 : 1720 – 1726 Google Scholar Crossref Search ADS PubMed WorldCat Peaucelle A , Louvet R, Johansen JN, Höfte H, Laufs P, Pelloux J, Mouille G ( 2008 ) Arabidopsis phyllotaxis is controlled by the methyl-esterification status of cell-wall pectins . Curr Biol 18 : 1943 – 1948 Google Scholar Crossref Search ADS PubMed WorldCat Peaucelle A , Wightman R, Höfte H ( 2015 ) The control of growth symmetry breaking in the Arabidopsis hypocotyl . Curr Biol 25 : 1746 – 1752 Google Scholar Crossref Search ADS PubMed WorldCat Pereira CS , Silveira RL, Dupree P, Skaf MS ( 2017 ) Effects of xylan side-chain substitutions on xylan-cellulose interactions and implications for thermal pretreatment of cellulosic biomass . Biomacromolecules 18 : 1311 – 1321 Google Scholar Crossref Search ADS PubMed WorldCat Phyo P , Wang T, Kiemle SN, O’Neill H, Pingali SV, Hong M, Cosgrove DJ ( 2017 ) Gradients in wall mechanics and polysaccharides along growing inflorescence stems . Plant Physiol 175 : 1593 – 1607 Google Scholar Crossref Search ADS PubMed WorldCat Pien S , Wyrzykowska J, McQueen-Mason S, Smart C, Fleming A ( 2001 ) Local expression of expansin induces the entire process of leaf development and modifies leaf shape . Proc Natl Acad Sci USA 98 : 11812 – 11817 Google Scholar Crossref Search ADS PubMed WorldCat Preston RD ( 1982 ) The case for multinet growth in growing walls of plant cells . Planta 155 : 356 – 363 Google Scholar Crossref Search ADS PubMed WorldCat Proseus TE , Zhu GL, Boyer JS ( 2000 ) Turgor, temperature and the growth of plant cells: using Chara corallina as a model system . J Exp Bot 51 : 1481 – 1494 Google Scholar Crossref Search ADS PubMed WorldCat Ray PM , Green PB, Cleland R ( 1972 ) Role of turgor in plant cell growth . Nature 239 : 163 – 164 Google Scholar Crossref Search ADS WorldCat Refrégier G , Pelletier S, Jaillard D, Höfte H ( 2004 ) Interaction between wall deposition and cell elongation in dark-grown hypocotyl cells in Arabidopsis . Plant Physiol 135 : 959 – 968 Google Scholar Crossref Search ADS PubMed WorldCat Reinhardt D , Wittwer F, Mandel T, Kuhlemeier C ( 1998 ) Localized upregulation of a new expansin gene predicts the site of leaf formation in the tomato meristem . Plant Cell 10 : 1427 – 1437 Google Scholar PubMed OpenURL Placeholder Text WorldCat Robinson S , Burian A, Couturier E, Landrein B, Louveaux M, Neumann ED, Peaucelle A, Weber A, Nakayama N ( 2013 ) Mechanical control of morphogenesis at the shoot apex . J Exp Bot 64 : 4729 – 4744 Google Scholar Crossref Search ADS PubMed WorldCat Rojas ER , Hotton S, Dumais J ( 2011 ) Chemically mediated mechanical expansion of the pollen tube cell wall . Biophys J 101 : 1844 – 1853 Google Scholar Crossref Search ADS PubMed WorldCat Routier-Kierzkowska AL , Smith RS ( 2013 ) Measuring the mechanics of morphogenesis . Curr Opin Plant Biol 16 : 25 – 32 Google Scholar Crossref Search ADS PubMed WorldCat Routier-Kierzkowska AL , Weber A, Kochova P, Felekis D, Nelson BJ, Kuhlemeier C, Smith RS ( 2012 ) Cellular force microscopy for in vivo measurements of plant tissue mechanics . Plant Physiol 158 : 1514 – 1522 Google Scholar Crossref Search ADS PubMed WorldCat Ruesink AW ( 1969 ) Polysaccharidases and the control of cell wall elongation . Planta 89 : 95 – 107 Google Scholar Crossref Search ADS PubMed WorldCat Sahaf M , Sharon E ( 2016 ) The rheology of a growing leaf: stress-induced changes in the mechanical properties of leaves . J Exp Bot 67 : 5509 – 5515 Google Scholar Crossref Search ADS PubMed WorldCat Sampathkumar A , Krupinski P, Wightman R, Milani P, Berquand A, Boudaoud A, Hamant O, Jönsson H, Meyerowitz EM ( 2014 a ) Subcellular and supracellular mechanical stress prescribes cytoskeleton behavior in Arabidopsis cotyledon pavement cells . eLife 3 : e01967 Google Scholar Crossref Search ADS PubMed WorldCat Sampathkumar A , Yan A, Krupinski P, Meyerowitz EM ( 2014 b ) Physical forces regulate plant development and morphogenesis . Curr Biol 24 : R475 – R483 Google Scholar Crossref Search ADS PubMed WorldCat Sanati Nezhad A , Geitmann A ( 2013 ) The cellular mechanics of an invasive lifestyle . J Exp Bot 64 : 4709 – 4728 Google Scholar Crossref Search ADS PubMed WorldCat Sanati Nezhad A , Naghavi M, Packirisamy M, Bhat R, Geitmann A ( 2013 ) Quantification of cellular penetrative forces using lab-on-a-chip technology and finite element modeling . Proc Natl Acad Sci USA 110 : 8093 – 8098 Google Scholar Crossref Search ADS PubMed WorldCat Sanati Nezhad A , Packirisamy M, Geitmann A ( 2014 ) Dynamic, high precision targeting of growth modulating agents is able to trigger pollen tube growth reorientation . Plant J 80 : 185 – 195 Google Scholar Crossref Search ADS PubMed WorldCat Sassi M , Ali O, Boudon F, Cloarec G, Abad U, Cellier C, Chen X, Gilles B, Milani P, Friml J, et al. ( 2014 ) An auxin-mediated shift toward growth isotropy promotes organ formation at the shoot meristem in Arabidopsis . Curr Biol 24 : 2335 – 2342 Google Scholar Crossref Search ADS PubMed WorldCat Shih HW , Miller ND, Dai C, Spalding EP, Monshausen GB ( 2014 ) The receptor-like kinase FERONIA is required for mechanical signal transduction in Arabidopsis seedlings . Curr Biol 24 : 1887 – 1892 Google Scholar Crossref Search ADS PubMed WorldCat Shinohara N , Sunagawa N, Tamura S, Yokoyama R, Ueda M, Igarashi K, Nishitani K ( 2017 ) The plant cell-wall enzyme AtXTH3 catalyses covalent cross-linking between cellulose and cello-oligosaccharide . Sci Rep 7 : 46099 Google Scholar Crossref Search ADS PubMed WorldCat Simmons TJ , Mohler KE, Holland C, Goubet F, Franková L, Houston DR, Hudson AD, Meulewaeter F, Fry SC ( 2015 ) Hetero-trans-β-glucanase, an enzyme unique to Equisetum plants, functionalizes cellulose . Plant J 83 : 753 – 769 Google Scholar Crossref Search ADS PubMed WorldCat Simmons TJ , Mortimer JC, Bernardinelli OD, Pöppler A-C, Brown SP, deAzevedo ER, Dupree R, Dupree P ( 2016 ) Folding of xylan onto cellulose fibrils in plant cell walls revealed by solid-state NMR . Nat Commun 7 : 13902 Google Scholar Crossref Search ADS PubMed WorldCat Suslov D , Verbelen JP, Vissenberg K ( 2009 ) Onion epidermis as a new model to study the control of growth anisotropy in higher plants . J Exp Bot 60 : 4175 – 4187 Google Scholar Crossref Search ADS PubMed WorldCat Szymanski DB ( 2014 ) The kinematics and mechanics of leaf expansion: new pieces to the Arabidopsis puzzle . Curr Opin Plant Biol 22 : 141 – 148 Google Scholar Crossref Search ADS PubMed WorldCat Szymanski DB , Cosgrove DJ ( 2009 ) Dynamic coordination of cytoskeletal and cell wall systems during plant cell morphogenesis . Curr Biol 19 : R800 – R811 Google Scholar Crossref Search ADS PubMed WorldCat Szymanski DB , Staiger C ( 2017 ) The actin cytoskeleton: functional arrays for cytoplasmic organization and cell shape control . Plant Physiol Google Scholar OpenURL Placeholder Text WorldCat Tan L , Eberhard S, Pattathil S, Warder C, Glushka J, Yuan C, Hao Z, Zhu X, Avci U, Miller JS, et al. ( 2013 ) An Arabidopsis cell wall proteoglycan consists of pectin and arabinoxylan covalently linked to an arabinogalactan protein . Plant Cell 25 : 270 – 287 Google Scholar Crossref Search ADS PubMed WorldCat Velasquez SM , Barbez E, Kleine-Vehn J, Estevez JM ( 2016 ) Auxin and cellular elongation . Plant Physiol 170 : 1206 – 1215 Google Scholar Crossref Search ADS PubMed WorldCat Virk SS , Cleland RE ( 1990 ) The role of wall calcium in the extension of cell walls of soybean hypocotyls . Planta 182 : 559 – 564 Google Scholar Crossref Search ADS WorldCat Vogler H , Felekis D, Nelson BJ, Grossniklaus U ( 2015 ) Measuring the mechanical properties of plant cell walls . Plants (Basel) 4 : 167 – 182 Google Scholar Crossref Search ADS PubMed WorldCat Voxeur A , Höfte H ( 2016 ) Cell wall integrity signaling in plants: “To grow or not to grow that’s the question” . Glycobiology 26 : 950 – 960 Google Scholar Crossref Search ADS PubMed WorldCat Wang T , Chen Y, Tabuchi A, Cosgrove DJ, Hong M ( 2016 a ) The target of β-expansin EXPB1 in maize cell walls from binding and solid-state NMR studies . Plant Physiol 172 : 2107 – 2119 Google Scholar Crossref Search ADS PubMed WorldCat Wang T , Hong M ( 2016 ) Solid-state NMR investigations of cellulose structure and interactions with matrix polysaccharides in plant primary cell walls . J Exp Bot 67 : 503 – 514 Google Scholar Crossref Search ADS PubMed WorldCat Wang T , Park YB, Caporini MA, Rosay M, Zhong L, Cosgrove DJ, Hong M ( 2013 ) Sensitivity-enhanced solid-state NMR detection of expansin’s target in plant cell walls . Proc Natl Acad Sci USA 110 : 16444 – 16449 Google Scholar Crossref Search ADS PubMed WorldCat Wang T , Park YB, Cosgrove DJ, Hong M ( 2015 ) Cellulose-pectin spatial contacts are inherent to never-dried Arabidopsis primary cell walls: Evidence from solid-state nuclear magnetic resonance . Plant Physiol 168 : 871 – 884 Google Scholar Crossref Search ADS PubMed WorldCat Wang T , Yang H, Kubicki JD, Hong M ( 2016 b ) Cellulose structural polymorphism in plant primary cell walls investigated by high-field 2D solid-state NMR spectroscopy and density functional theory calculations . Biomacromolecules 17 : 2210 – 2222 Google Scholar Crossref Search ADS PubMed WorldCat Wang T , Zabotina O, Hong M ( 2012 ) Pectin-cellulose interactions in the Arabidopsis primary cell wall from two-dimensional magic-angle-spinning solid-state nuclear magnetic resonance . Biochemistry 51 : 9846 – 9856 Google Scholar Crossref Search ADS PubMed WorldCat White PB , Wang T, Park YB, Cosgrove DJ, Hong M ( 2014 ) Water-polysaccharide interactions in the primary cell wall of Arabidopsis thaliana from polarization transfer solid-state NMR . J Am Chem Soc 136 : 10399 – 10409 Google Scholar Crossref Search ADS PubMed WorldCat Wilson RH , Smith AC, Kacuráková M, Saunders PK, Wellner N, Waldron KW ( 2000 ) The mechanical properties and molecular dynamics of plant cell wall polysaccharides studied by Fourier-transform infrared spectroscopy . Plant Physiol 124 : 397 – 405 Google Scholar Crossref Search ADS PubMed WorldCat Wolf S , Mravec J, Greiner S, Mouille G, Höfte H ( 2012 ) Plant cell wall homeostasis is mediated by brassinosteroid feedback signaling . Curr Biol 22 : 1732 – 1737 Google Scholar Crossref Search ADS PubMed WorldCat Wolf S , van der Does D, Ladwig F, Sticht C, Kolbeck A, Schürholz AK, Augustin S, Keinath N, Rausch T, Greiner S, et al. ( 2014 ) A receptor-like protein mediates the response to pectin modification by activating brassinosteroid signaling . Proc Natl Acad Sci USA 111 : 15261 – 15266 Google Scholar Crossref Search ADS PubMed WorldCat Xiao C , Zhang T, Zheng Y, Cosgrove DJ, Anderson CT ( 2016 ) Xyloglucan deficiency disrupts microtubule stability and cellulose biosynthesis in Arabidopsis, altering cell growth and morphogenesis . Plant Physiol 170 : 234 – 249 Google Scholar Crossref Search ADS PubMed WorldCat Yanagisawa M , Desyatova AS, Belteton SA, Mallery EL, Turner JA, Szymanski DB ( 2015 ) Patterning mechanisms of cytoskeletal and cell wall systems during leaf trichome morphogenesis . Nat Plants 1 : 15014 Google Scholar Crossref Search ADS PubMed WorldCat Yang W , Schuster C, Beahan CT, Charoensawan V, Peaucelle A, Bacic A, Doblin MS, Wightman R, Meyerowitz EM ( 2016 ) Regulation of meristem morphogenesis by cell wall synthases in Arabidopsis . Curr Biol 26 : 1404 – 1415 Google Scholar Crossref Search ADS PubMed WorldCat Yu ZM , Kang B, He XW, Lv SL, Bai YH, Ding WN, Chen M, Cho HT, Wu P ( 2011 ) Root hair-specific expansins modulate root hair elongation in rice . Plant J 66 : 725 – 734 Google Scholar Crossref Search ADS PubMed WorldCat Yuan S , Wu Y, Cosgrove DJ ( 2001 ) A fungal endoglucanase with plant cell wall extension activity . Plant Physiol 127 : 324 – 333 Google Scholar Crossref Search ADS PubMed WorldCat Zablackis E , Huang J, Müller B, Darvill AG, Albersheim P ( 1995 ) Characterization of the cell-wall polysaccharides of Arabidopsis thaliana leaves . Plant Physiol 107 : 1129 – 1138 Google Scholar Crossref Search ADS PubMed WorldCat Zhang C , Mallery EL, Szymanski DB ( 2013 ) ARP2/3 localization in Arabidopsis leaf pavement cells: a diversity of intracellular pools and cytoskeletal interactions . Front Plant Sci 4 : 238 Google Scholar PubMed OpenURL Placeholder Text WorldCat Zhang T , Mahgsoudy-Louyeh S, Tittmann B, Cosgrove DJ ( 2014 ) Visualization of the nanoscale pattern of recently-deposited cellulose microfibrils and matrix materials in never-dried primary walls of the onion epidermis . Cellulose 21 : 853 – 862 Google Scholar Crossref Search ADS WorldCat Zhang T , Vavylonis D, Durachko DM, Cosgrove DJ ( 2017 ) Nanoscale movements of cellulose microfibrils in primary cell walls . Nat Plants 3 : 17056 Google Scholar Crossref Search ADS PubMed WorldCat Zhang T , Zheng Y, Cosgrove DJ ( 2016 ) 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 Google Scholar Crossref Search ADS PubMed WorldCat Zhao Q , Yuan S, Wang X, Zhang Y, Zhu H, Lu C ( 2008 ) Restoration of mature etiolated cucumber hypocotyl cell wall susceptibility to expansin by pretreatment with fungal pectinases and EGTA in vitro . Plant Physiol 147 : 1874 – 1885 Google Scholar Crossref Search ADS PubMed WorldCat Zhao Z , Crespi VH, Kubicki JD, Cosgrove DJ, Zhong L ( 2014 ) Molecular dynamics simulation study of xyloglucan adsorption on cellulose surfaces: effects of surface hydrophobicity and side-chain variation . Cellulose 21 : 1025 – 1039 Google Scholar Crossref Search ADS WorldCat Zheng Y , Cosgrove DJ, Ning G ( 2017 a ) High-resolution field emission scanning electron microscopy (FESEM) imaging of cellulose microfibril organization in plant primary cell walls . Microsc Microanal 23 : 1048 – 1054 Google Scholar Crossref Search ADS PubMed WorldCat Zheng Y , Wang X, Chen Y, Wagner E, Cosgrove DJ ( 2017 b ) Xyloglucan in the primary cell wall: assessment by FESEM, selective enzyme digestions and nanogold affinity tags . Plant J http://dx.doi.org/10.1111/tpj.13778 Google Scholar OpenURL Placeholder Text WorldCat Zhu C , Ganguly A, Baskin TI, McClosky DD, Anderson CT, Foster C, Meunier KA, Okamoto R, Berg H, Dixit R ( 2015 ) The fragile Fiber1 kinesin contributes to cortical microtubule-mediated trafficking of cell wall components . Plant Physiol 167 : 780 – 792 Google Scholar Crossref Search ADS PubMed WorldCat Zykwinska A , Thibault JF, Ralet MC ( 2008 a ) Competitive binding of pectin and xyloglucan with primary cell wall cellulose . Carbohydr Polym 74 : 957 – 961 Google Scholar Crossref Search ADS WorldCat Zykwinska A , Thibault JF, Ralet MC ( 2008 b ) Modelling of xyloglucan, pectins and pectic side chains binding onto cellulose microfibrils . Carbohydr Polym 74 : 23 – 30 Google Scholar Crossref Search ADS WorldCat Author notes 1 Work on expansins is supported by the U.S. Department of Energy (grant no. DE-FG2-84ER13179) and work on cell wall structure is supported as part of the Center for LignoCellulose Structure and Formation, an Energy Frontier Research Center funded by the U.S. Department of Energy, Office of Science, Basic Energy Sciences (award no. DE-SC0001090). 2 Address correspondence to [email protected]. D.J.C. wrote the article. [OPEN] Articles can be viewed without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.17.01541 © 2018 American Society of Plant Biologists. All Rights Reserved. © The Author(s) 2018. Published by Oxford University Press on behalf of American Society of Plant Biologists. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.

Cosgrove, Daniel J.

doi: 10.1104/pp.17.01541pmid: 29138349

Bascom, Carlisle S.; Hepler, Peter K.; Bezanilla, Magdalena

doi: 10.1104/pp.17.01466pmid: 29138353

Tip growth is a specialized type of growth utilized by only a few cell types in the plant kingdom. Nevertheless, these cell types perform vital functions for plants. Pollen tubes are necessary for seed plant reproduction, root hairs are vital for nutrient acquisition, and protonemata comprise the entire juvenile moss plant. Tip growth requires exquisite coordination of intracellular cargo delivery and simultaneous weakening of the apical cell wall. This process is precarious, with imbalances resulting in cell rupture or cessation of growth. In this Update, we provide context and highlight recent advances in our understanding of how ions, the cytoskeleton, and properties of the cell wall are interconnected during tip growth across three tip-growing systems: pollen tubes, root hairs, and protonemata. We propose that further mechanistic insights are likely to come from studying the commonalities among these systems, particularly focusing on the molecular basis of the communication between cell wall status and intracellular cargo delivery. Plant cells are surrounded by a cell wall that imposes both physical restrictions to shape as well as a balance to turgor pressure. However, most undifferentiated plant cells are small and uniform. During differentiation, plant cells expand, which occurs when loosening events allow the wall to yield to the existing turgor pressure, discussed further in this issue (Cosgrove, 2018). The momentary decrease in water potential results in an influx of water, causing expansion. As new cell wall material is deposited to account for this expansion, the increase in size becomes irreversible, and thus the cell has grown. Modifications to cell wall viscosity promoting expansion can result from a number of different processes. For example, cell wall-loosening enzymes may be activated in or delivered to specific regions. Alternatively, the secretion of flexible cell wall material could be spatially regulated. One extreme form of cell wall patterning is known as tip growth. In tip-growing cells, the zone of reduced cell wall viscosity is focused at the apex of the growing cell. To grow, the cell must balance the delivery of wall material with the retrieval of excess membrane, as the delivery of excess flexible wall material at the cell tip could result in lysis. Tip growth occurs across plant lineages in cell types such as pollen tubes, root hairs, moss protonemata, and bryophyte and algal rhizoids. Although tip growth mechanisms have been generalized as equivalent in pollen tubes, root hairs, and moss protonemata, it is likely that significant differences exist in order for these cells to carry out their divergent functions. Open in new tabDownload slide Open in new tabDownload slide Of the three cell types, pollen tubes are the most specialized and short-lived cell, in that their aim is to deliver the sperm cells to the ovule. By transmitting location information about the ovule from outside the pollen tube to the intracellular growth machinery, pollen tubes grow rapidly toward the female gametophyte and burst upon entry. Root hairs are the site of nutrient and water uptake for the plant. In contrast to pollen tubes, root hairs are longer-lived and must integrate many environmental cues (moisture, nutrient content, soil texture, etc.). Root hairs internalize this suite of information to maximize the efficacy of their growth. In contrast to pollen tubes and root hairs, moss protonemata have a colonization role. They germinate from the spore and race against other plants to acquire moisture, nutrients, and an optimal position for photosynthesis. This tip-growing cell continues to undergo cell divisions as the plant develops, reaching a point in which the entire juvenile plant, persisting on the scale of months, consists entirely of protonemata. Thus, moss protonemata are longer-lived than both pollen tubes and root hairs and integrate substantially more environmental information during their growth and development. Even with such divergent functions, mechanistically describing tip growth in each system promises to illuminate fundamental processes conserved throughout plant evolution as well as derived processes that may have evolved in each cell type. Ultimately, the key to understanding tip growth will require understanding a complex interaction network; the reader is directed to recent reviews that expand upon a variety of facets of tip growth not covered here (Mendrinna and Persson, 2015; Hepler, 2016; Damineli et al., 2017; Michard et al., 2017; Stephan, 2017). In this Update, we speculate that further mechanistic advances will come from identifying deeply conserved tip growth mechanisms that decipher signaling events between the cell wall and the cytoplasm. Thus, we will focus on advances in the understanding of cytoskeletal organization, intracellular ion gradients, and the cell wall itself, all key aspects involved in tip growth signaling. A link between wall status and cytoplasmic activity is critical because, in many ways, tip growth resembles a controlled explosion: the cell wall must weaken exactly enough to allow for expansion but not so much that the cell ruptures. Indeed, there are many conditions under which this balance is lost, resulting in either growth arrest or tip bursting. We have compiled a list of agents and mutants that uncouple various aspects of tip growth with similar results (Table I). While not an exhaustive list, Table I demonstrates the wide-ranging conditions that disrupt the precarious balancing act between intracellular and extracellular events during tip growth. By identifying the molecular targets of agents and cellular deficiencies in mutants leading to growth arrest or tip blowout, it may be possible to understand the molecular regulation of this controlled explosion (see Outstanding Questions). Agents and mutants abrogating tip growth Table I. Agents and mutants abrogating tip growth RNAi, RNA interference; N/A, not applicable. Treatment/Mutant . Cellular Consequence . Gene Name . Growth Consequence . System . Reference . Latrunculin B Actin depolymerization N/A Swelling, loss of polarized growth Pollen tubes,
root hairs,
protonemata Gibbon et al. (1999); Ketelaar et al. (2003);Harries et al. (2005) Calcium ionophore Equalizes calcium concentrations N/A Apical wall thickening, growth reorientation Root hairs, pollen tubes Malho and Trewavas (1996); Monshausen et al. (2008) Oryzalin Microtubule depolymerization N/A Loss of directional growth Root hairs,
moss protonemata Doonan et al. (1988); Bibikova et al. (1999) Lanthanum Calcium channel and pump blocker N/A Bursting Pollen tubes,
root hairs Monshausen et al. (2008) Cyanide ATP synthesis inhibition N/A Apical wall thickening Pollen tubes Winship et al. (2016) Propidium iodide Competes with calcium in cell wall N/A Bursting Pollen tubes Rounds et al. (2011) EGTA/EDTA Low extracellular calcium N/A Bursting Pollen tubes,
root hairs Monshausen et al. (2008) Acidic growth medium Low extracellular pH N/A Bursting Root hairs Monshausen et al. (2008) Yariv agent Arabinogalactan protein blocking N/A Expansion stops,
deposition continues Pollen tubes,
moss protonemata Roy et al. (1999); Lee et al. (2005) apg knockdown/knockout Aberrant extracellular signaling ARABINOGALACTAN Shorter cells Pollen tubes,
moss protonemata Lee et al. (2005); Levitin et al. (2008) cngc mutants Aberrant calcium signaling CYCLIC NUCELOTIDE-GATED CHANNELS Shorter, deformed cells Pollen tubes,
root hairs Gao et al. (2016);Zhang et al. (2017) glr1.2 mutant Aberrant calcium signaling GLU RECEPTOR-LIKE Deformed cells Pollen tubes Michard et al. (2011) exo70C mutant Faster growth rate, thin cell wall EXOCYTOSIS 70C2 Bursting Pollen tubes Synek et al. (2017) cog3 and cog8 single mutants Aberrant Golgi morphology, vesicle trafficking CONSERVED OLIGOMERIC GOLGI 3 and 8 Bursting Pollen tubes Tan et al. (2016) anx1,anx2 double mutant Aberrant cell wall formation ANXUR 1 and 2 Bursting Pollen tubes Boisson-Dernier et al. (2009) LePRK RNAi/overexpression Aberrant extracellular signaling POLLEN RECEPTOR KINASE Knockdown: shorter pollen tubes, bursting;
overexpression: ballooning of tip Pollen tubes Gui et al. (2014) lrx1,lrx2 double mutant Aberrant extracellular signaling LUCINE-RICH REPEAT EXTENSIN 1 and 2 Rupture after initiation Root hairs Baumberger et al. (2003) prx44 mutant Aberrant extracellular signaling PEROXIDASE44 Bursting Root hairs Kwon et al. (2015) kjk/csld3 mutant Aberrant cell wall formation KOJAK/cellulose synthase-like D3 Rupture after initiation Root hairs Favery et al. (2001) bup mutant Loss of germination plaque BURSTING POLLEN (Golgi-localized glycosyltransferase) Bursts upon germination Pollen tubes Hoedemaekers et al. (2015) ext18 mutant Aberrant cell wall formation EXTENSIN18 Bursting Pollen tubes Choudhary et al. (2015) reb1 mutant Altered sugar metabolism ROOT EPIDERMAL BULGER1 Ballooning Root hairs Andème-Onzighi et al. (2002) ROP RNAi Aberrant intracellular signaling Rho/Rac of Plants (small GTPase) Loss of polarized growth, loss of cell adhesion Moss protonemata Burkart et al. (2015) ROP overexpression Aberrant intracellular signaling Rho/Rac of Plants Swelling,
branching Pollen tubes,
root hairs Wu et al. (2001);
Jones et al. (2002) Treatment/Mutant . Cellular Consequence . Gene Name . Growth Consequence . System . Reference . Latrunculin B Actin depolymerization N/A Swelling, loss of polarized growth Pollen tubes,
root hairs,
protonemata Gibbon et al. (1999); Ketelaar et al. (2003);Harries et al. (2005) Calcium ionophore Equalizes calcium concentrations N/A Apical wall thickening, growth reorientation Root hairs, pollen tubes Malho and Trewavas (1996); Monshausen et al. (2008) Oryzalin Microtubule depolymerization N/A Loss of directional growth Root hairs,
moss protonemata Doonan et al. (1988); Bibikova et al. (1999) Lanthanum Calcium channel and pump blocker N/A Bursting Pollen tubes,
root hairs Monshausen et al. (2008) Cyanide ATP synthesis inhibition N/A Apical wall thickening Pollen tubes Winship et al. (2016) Propidium iodide Competes with calcium in cell wall N/A Bursting Pollen tubes Rounds et al. (2011) EGTA/EDTA Low extracellular calcium N/A Bursting Pollen tubes,
root hairs Monshausen et al. (2008) Acidic growth medium Low extracellular pH N/A Bursting Root hairs Monshausen et al. (2008) Yariv agent Arabinogalactan protein blocking N/A Expansion stops,
deposition continues Pollen tubes,
moss protonemata Roy et al. (1999); Lee et al. (2005) apg knockdown/knockout Aberrant extracellular signaling ARABINOGALACTAN Shorter cells Pollen tubes,
moss protonemata Lee et al. (2005); Levitin et al. (2008) cngc mutants Aberrant calcium signaling CYCLIC NUCELOTIDE-GATED CHANNELS Shorter, deformed cells Pollen tubes,
root hairs Gao et al. (2016);Zhang et al. (2017) glr1.2 mutant Aberrant calcium signaling GLU RECEPTOR-LIKE Deformed cells Pollen tubes Michard et al. (2011) exo70C mutant Faster growth rate, thin cell wall EXOCYTOSIS 70C2 Bursting Pollen tubes Synek et al. (2017) cog3 and cog8 single mutants Aberrant Golgi morphology, vesicle trafficking CONSERVED OLIGOMERIC GOLGI 3 and 8 Bursting Pollen tubes Tan et al. (2016) anx1,anx2 double mutant Aberrant cell wall formation ANXUR 1 and 2 Bursting Pollen tubes Boisson-Dernier et al. (2009) LePRK RNAi/overexpression Aberrant extracellular signaling POLLEN RECEPTOR KINASE Knockdown: shorter pollen tubes, bursting;
overexpression: ballooning of tip Pollen tubes Gui et al. (2014) lrx1,lrx2 double mutant Aberrant extracellular signaling LUCINE-RICH REPEAT EXTENSIN 1 and 2 Rupture after initiation Root hairs Baumberger et al. (2003) prx44 mutant Aberrant extracellular signaling PEROXIDASE44 Bursting Root hairs Kwon et al. (2015) kjk/csld3 mutant Aberrant cell wall formation KOJAK/cellulose synthase-like D3 Rupture after initiation Root hairs Favery et al. (2001) bup mutant Loss of germination plaque BURSTING POLLEN (Golgi-localized glycosyltransferase) Bursts upon germination Pollen tubes Hoedemaekers et al. (2015) ext18 mutant Aberrant cell wall formation EXTENSIN18 Bursting Pollen tubes Choudhary et al. (2015) reb1 mutant Altered sugar metabolism ROOT EPIDERMAL BULGER1 Ballooning Root hairs Andème-Onzighi et al. (2002) ROP RNAi Aberrant intracellular signaling Rho/Rac of Plants (small GTPase) Loss of polarized growth, loss of cell adhesion Moss protonemata Burkart et al. (2015) ROP overexpression Aberrant intracellular signaling Rho/Rac of Plants Swelling,
branching Pollen tubes,
root hairs Wu et al. (2001);
Jones et al. (2002) Open in new tab Table I. Agents and mutants abrogating tip growth RNAi, RNA interference; N/A, not applicable. Treatment/Mutant . Cellular Consequence . Gene Name . Growth Consequence . System . Reference . Latrunculin B Actin depolymerization N/A Swelling, loss of polarized growth Pollen tubes,
root hairs,
protonemata Gibbon et al. (1999); Ketelaar et al. (2003);Harries et al. (2005) Calcium ionophore Equalizes calcium concentrations N/A Apical wall thickening, growth reorientation Root hairs, pollen tubes Malho and Trewavas (1996); Monshausen et al. (2008) Oryzalin Microtubule depolymerization N/A Loss of directional growth Root hairs,
moss protonemata Doonan et al. (1988); Bibikova et al. (1999) Lanthanum Calcium channel and pump blocker N/A Bursting Pollen tubes,
root hairs Monshausen et al. (2008) Cyanide ATP synthesis inhibition N/A Apical wall thickening Pollen tubes Winship et al. (2016) Propidium iodide Competes with calcium in cell wall N/A Bursting Pollen tubes Rounds et al. (2011) EGTA/EDTA Low extracellular calcium N/A Bursting Pollen tubes,
root hairs Monshausen et al. (2008) Acidic growth medium Low extracellular pH N/A Bursting Root hairs Monshausen et al. (2008) Yariv agent Arabinogalactan protein blocking N/A Expansion stops,
deposition continues Pollen tubes,
moss protonemata Roy et al. (1999); Lee et al. (2005) apg knockdown/knockout Aberrant extracellular signaling ARABINOGALACTAN Shorter cells Pollen tubes,
moss protonemata Lee et al. (2005); Levitin et al. (2008) cngc mutants Aberrant calcium signaling CYCLIC NUCELOTIDE-GATED CHANNELS Shorter, deformed cells Pollen tubes,
root hairs Gao et al. (2016);Zhang et al. (2017) glr1.2 mutant Aberrant calcium signaling GLU RECEPTOR-LIKE Deformed cells Pollen tubes Michard et al. (2011) exo70C mutant Faster growth rate, thin cell wall EXOCYTOSIS 70C2 Bursting Pollen tubes Synek et al. (2017) cog3 and cog8 single mutants Aberrant Golgi morphology, vesicle trafficking CONSERVED OLIGOMERIC GOLGI 3 and 8 Bursting Pollen tubes Tan et al. (2016) anx1,anx2 double mutant Aberrant cell wall formation ANXUR 1 and 2 Bursting Pollen tubes Boisson-Dernier et al. (2009) LePRK RNAi/overexpression Aberrant extracellular signaling POLLEN RECEPTOR KINASE Knockdown: shorter pollen tubes, bursting;
overexpression: ballooning of tip Pollen tubes Gui et al. (2014) lrx1,lrx2 double mutant Aberrant extracellular signaling LUCINE-RICH REPEAT EXTENSIN 1 and 2 Rupture after initiation Root hairs Baumberger et al. (2003) prx44 mutant Aberrant extracellular signaling PEROXIDASE44 Bursting Root hairs Kwon et al. (2015) kjk/csld3 mutant Aberrant cell wall formation KOJAK/cellulose synthase-like D3 Rupture after initiation Root hairs Favery et al. (2001) bup mutant Loss of germination plaque BURSTING POLLEN (Golgi-localized glycosyltransferase) Bursts upon germination Pollen tubes Hoedemaekers et al. (2015) ext18 mutant Aberrant cell wall formation EXTENSIN18 Bursting Pollen tubes Choudhary et al. (2015) reb1 mutant Altered sugar metabolism ROOT EPIDERMAL BULGER1 Ballooning Root hairs Andème-Onzighi et al. (2002) ROP RNAi Aberrant intracellular signaling Rho/Rac of Plants (small GTPase) Loss of polarized growth, loss of cell adhesion Moss protonemata Burkart et al. (2015) ROP overexpression Aberrant intracellular signaling Rho/Rac of Plants Swelling,
branching Pollen tubes,
root hairs Wu et al. (2001);
Jones et al. (2002) Treatment/Mutant . Cellular Consequence . Gene Name . Growth Consequence . System . Reference . Latrunculin B Actin depolymerization N/A Swelling, loss of polarized growth Pollen tubes,
root hairs,
protonemata Gibbon et al. (1999); Ketelaar et al. (2003);Harries et al. (2005) Calcium ionophore Equalizes calcium concentrations N/A Apical wall thickening, growth reorientation Root hairs, pollen tubes Malho and Trewavas (1996); Monshausen et al. (2008) Oryzalin Microtubule depolymerization N/A Loss of directional growth Root hairs,
moss protonemata Doonan et al. (1988); Bibikova et al. (1999) Lanthanum Calcium channel and pump blocker N/A Bursting Pollen tubes,
root hairs Monshausen et al. (2008) Cyanide ATP synthesis inhibition N/A Apical wall thickening Pollen tubes Winship et al. (2016) Propidium iodide Competes with calcium in cell wall N/A Bursting Pollen tubes Rounds et al. (2011) EGTA/EDTA Low extracellular calcium N/A Bursting Pollen tubes,
root hairs Monshausen et al. (2008) Acidic growth medium Low extracellular pH N/A Bursting Root hairs Monshausen et al. (2008) Yariv agent Arabinogalactan protein blocking N/A Expansion stops,
deposition continues Pollen tubes,
moss protonemata Roy et al. (1999); Lee et al. (2005) apg knockdown/knockout Aberrant extracellular signaling ARABINOGALACTAN Shorter cells Pollen tubes,
moss protonemata Lee et al. (2005); Levitin et al. (2008) cngc mutants Aberrant calcium signaling CYCLIC NUCELOTIDE-GATED CHANNELS Shorter, deformed cells Pollen tubes,
root hairs Gao et al. (2016);Zhang et al. (2017) glr1.2 mutant Aberrant calcium signaling GLU RECEPTOR-LIKE Deformed cells Pollen tubes Michard et al. (2011) exo70C mutant Faster growth rate, thin cell wall EXOCYTOSIS 70C2 Bursting Pollen tubes Synek et al. (2017) cog3 and cog8 single mutants Aberrant Golgi morphology, vesicle trafficking CONSERVED OLIGOMERIC GOLGI 3 and 8 Bursting Pollen tubes Tan et al. (2016) anx1,anx2 double mutant Aberrant cell wall formation ANXUR 1 and 2 Bursting Pollen tubes Boisson-Dernier et al. (2009) LePRK RNAi/overexpression Aberrant extracellular signaling POLLEN RECEPTOR KINASE Knockdown: shorter pollen tubes, bursting;
overexpression: ballooning of tip Pollen tubes Gui et al. (2014) lrx1,lrx2 double mutant Aberrant extracellular signaling LUCINE-RICH REPEAT EXTENSIN 1 and 2 Rupture after initiation Root hairs Baumberger et al. (2003) prx44 mutant Aberrant extracellular signaling PEROXIDASE44 Bursting Root hairs Kwon et al. (2015) kjk/csld3 mutant Aberrant cell wall formation KOJAK/cellulose synthase-like D3 Rupture after initiation Root hairs Favery et al. (2001) bup mutant Loss of germination plaque BURSTING POLLEN (Golgi-localized glycosyltransferase) Bursts upon germination Pollen tubes Hoedemaekers et al. (2015) ext18 mutant Aberrant cell wall formation EXTENSIN18 Bursting Pollen tubes Choudhary et al. (2015) reb1 mutant Altered sugar metabolism ROOT EPIDERMAL BULGER1 Ballooning Root hairs Andème-Onzighi et al. (2002) ROP RNAi Aberrant intracellular signaling Rho/Rac of Plants (small GTPase) Loss of polarized growth, loss of cell adhesion Moss protonemata Burkart et al. (2015) ROP overexpression Aberrant intracellular signaling Rho/Rac of Plants Swelling,
branching Pollen tubes,
root hairs Wu et al. (2001);
Jones et al. (2002) Open in new tab THE CYTOSKELETON Within the cell, both the actin and microtubule cytoskeletons contribute to tip growth. As we discuss the structure of these cytoskeletons in tip-growing cells, it is important to recall that each array is decorated with, and regulated by, microtubule- and actin-binding proteins. Thus, their localization (Table II) during tip growth provides insights into the mechanism of cytoskeletal organization and dynamics. Localization of cytoskeleton-binding proteins Table II. Localization of cytoskeleton-binding proteins MTs, Microtubules; PM, plasma membrane. Protein . Localization . Species/System . Reference . Fimbrin 1) Apical and subapical F-actin structures 1) Lily pollen tubes 1) Su et al. (2012), α-LlFIM1 antibody 2) Apical filaments 2) Arabidopsis pollen tubes 2) Zhang et al. (2016), pFIM5::FIM5-GFP Villin 1) AtVLN3, thick subapical filaments 1) Arabidopsis root hairs 1) van der Honing et al. (2012), pAtVLN3::VLN3-GFP 2) AtVLN2, thick subapical filaments, fine apical filaments 2 and 3) Arabidopsis pollen tubes 2) Qu et al. (2013), pAtVLN2::VLN2genomic-GFP 3) AtVLN5, thick subapical filaments, fine apical filaments 3) Qu et al. (2013), pAtVLN5::VLN5genomic-GFP Actin Depolymerizing Factor (ADF)/Cofilin 1) Apical and subapical actin structures 1) Tobacco pollen tubes and grains 1) Chen et al. (2002), Lat52::NtADF-GFP 2) Apical structures 2) Lily pollen tubes 2) Lovy-Wheeler et al. (2006), α-lily ADF antibody 3) Cytoplasmic 3) P. patens protonemata 3) Augustine et al. (2008), α-PpADF polyclonal antibody Profilin Cytoplasmic Lily pollen grains and tubes Vidali and Hepler (1997), α-profilin polyclonal antibody Actin Interacting Protein1 1) Apical F-actin fringe 1) Lilium spp. pollen tubes 1) Lovy-Wheeler et al. (2006), α-lily AIP antibody 2) Cytoplasmic 2) P. patens protonemata 2) Augustine et al. (2011), AIP-mCherry locus integration Class I Formin LlFH1, apical vesicles, PM Lily pollen tubes Li et al. (2017), transient overexpression of LlFH1-GFP Class II Formin PpFor2a, apex, not on PM P. patens protonemata Vidali et al. (2009b), For2a 3xGFP locus integration Class VIII Myosin PpMyo8A, cortical puncta with apical accumulation, spindle and phragmoplast midzone P. patens protonemata Wu and Bezanilla (2014), pMaizeUbiquitin::Myo8A-3xGFP, in Ɗmyo8 a,b,c,d,e Class XI Myosin 1) PpMyo11a cytosolic with apical enrichment 1) P. patens protonemata 1) Vidali et al. (2010), pMaizeUbiquitin::3xGFP-Myo11a 2) AtMyo11k, apical cloud 2) Arabidopsis root hairs 2) Park and Nebenführ (2013), pMyo11K::YFP-Myo11K Kinesin 1) KIND1a,b, plus ends of MTs focused on the tip 1) P. patens protonemata 1) Hiwatashi et al. (2014), KIND1a/1b-cerulean locus integration 2) ARK1, cytoplasmic, plus ends of MTs 2) Arabidopsis root hairs 2) Eng and Wasteneys (2014), pARK1::ARK1-GFP Actin Related Protein2/3 (ARP2/3) complex PpARPC4, apically enriched cap P. patens protonemata Perroud and Quatrano (2006), 2xYFP locus integration Brick1 (brk1) PpBRK1, apically enriched cap P. patens protonemata Perroud and Quatrano (2008), 3xYFP locus integration Rho/Rac of Plants (ROP) 1) AtROP2, apical dome 1) Tobacco pollen tubes 1) Kost et al. (1999), LAT52::GFP-AtROP2 2) ROP1, apical dome 2) Arabidopsis root hairs 2) Jones et al. (2002), 35S::GFP-ROP2 3) AtROP1-PM, apical enrichment, cytoplasm 3) Tobacco pollen tubes 3) Hwang et al. (2005), LAT52::RIC4ƊC-GFP 4) PpROP2-PM, enrichment at cross walls 4 and 5) P. patens protonemata 4) Ito et al. (2014), HSP::cerulean-PpROP2 5) PpROP4, apical PM and cross walls 5) P. patens protonemata 5) Burkart et al. (2015), pROP4::GFP-ROP4coding sequence, locus replacement ROP Activating Proteins (RopGAPs) NtRopGAP1, subapical PM Tobacco pollen tubes Klahre and Kost (2006), Lat52::YFP-NtRopGAP1 ROP Interacting CRIB-Containing Protein (RIC) 1) AtRIC 3,10, cytosolic 1 to 4) Tobacco pollen tubes 1) Wu et al. (2001), Lat52::GFP-RIC n 2) AtRIC 1, 6, 7, apical PM, cytosolic 2) Wu et al. (2001), Lat52::GFP-RIC n 3) AtRIC9, PM 3) Wu et al. (2001), Lat52::GFP-RIC 9 4) AtRIC 2, 4, 5, apical PM 4) Wu et al. (2001), Lat52::GFP-RIC n ROP Guanine Dissociation Inhibitors (RopGDIs) NtRopGDI2, cytosolic Tobacco pollen tubes Klahre et al. (2006), Lat52::YFP-NtRopGDI2 ROP Guanine Exchange Factors (RopGEFs) 1) AtRopGEF1, apical PM 1 to 3) Tobacco pollen tubes 1) Gu et al. (2006), Lat52::GFP-RopGEF1 2) AtRopGEF 8, 9, 14, apical PM, cytoplasm 2) Gu et al. (2006), Lat52::GFP-RopGEF n 3) AtRopGEF12, cytoplasm 3) Gu et al. (2006), Lat52::GFP-RopGEF12 4) PpRopGEF3, cytoplasmic apical enrichment 4) P. patens protonemata 4) Ito et al. (2014), HSP::PpRopGEF3-cerulean Microtubule-Associated Protein18 1) Shank PM, inverted cone at tip 1) Arabidopsis pollen tubes 1) Zhu et al. (2013), pAtMAP18::AtMAP18-GFP 2) Shank of PM, apical cytoplasm 2) Arabidopsis root hairs 2) Kang et al. (2017), pAtMAP18::AtMAP18-mCherry End-Binding1 1) Cortical puncta, apical accumulation 1) P. patens protonemata 1) Hiwatashi et al. (2014), PpEB1b-mCitrine locus integration 2) Cytoplasmic puncta 2) Arabidopsis root hairs 2) Eng and Wasteneys (2014), 35S::AtEB1b-GFP Protein . Localization . Species/System . Reference . Fimbrin 1) Apical and subapical F-actin structures 1) Lily pollen tubes 1) Su et al. (2012), α-LlFIM1 antibody 2) Apical filaments 2) Arabidopsis pollen tubes 2) Zhang et al. (2016), pFIM5::FIM5-GFP Villin 1) AtVLN3, thick subapical filaments 1) Arabidopsis root hairs 1) van der Honing et al. (2012), pAtVLN3::VLN3-GFP 2) AtVLN2, thick subapical filaments, fine apical filaments 2 and 3) Arabidopsis pollen tubes 2) Qu et al. (2013), pAtVLN2::VLN2genomic-GFP 3) AtVLN5, thick subapical filaments, fine apical filaments 3) Qu et al. (2013), pAtVLN5::VLN5genomic-GFP Actin Depolymerizing Factor (ADF)/Cofilin 1) Apical and subapical actin structures 1) Tobacco pollen tubes and grains 1) Chen et al. (2002), Lat52::NtADF-GFP 2) Apical structures 2) Lily pollen tubes 2) Lovy-Wheeler et al. (2006), α-lily ADF antibody 3) Cytoplasmic 3) P. patens protonemata 3) Augustine et al. (2008), α-PpADF polyclonal antibody Profilin Cytoplasmic Lily pollen grains and tubes Vidali and Hepler (1997), α-profilin polyclonal antibody Actin Interacting Protein1 1) Apical F-actin fringe 1) Lilium spp. pollen tubes 1) Lovy-Wheeler et al. (2006), α-lily AIP antibody 2) Cytoplasmic 2) P. patens protonemata 2) Augustine et al. (2011), AIP-mCherry locus integration Class I Formin LlFH1, apical vesicles, PM Lily pollen tubes Li et al. (2017), transient overexpression of LlFH1-GFP Class II Formin PpFor2a, apex, not on PM P. patens protonemata Vidali et al. (2009b), For2a 3xGFP locus integration Class VIII Myosin PpMyo8A, cortical puncta with apical accumulation, spindle and phragmoplast midzone P. patens protonemata Wu and Bezanilla (2014), pMaizeUbiquitin::Myo8A-3xGFP, in Ɗmyo8 a,b,c,d,e Class XI Myosin 1) PpMyo11a cytosolic with apical enrichment 1) P. patens protonemata 1) Vidali et al. (2010), pMaizeUbiquitin::3xGFP-Myo11a 2) AtMyo11k, apical cloud 2) Arabidopsis root hairs 2) Park and Nebenführ (2013), pMyo11K::YFP-Myo11K Kinesin 1) KIND1a,b, plus ends of MTs focused on the tip 1) P. patens protonemata 1) Hiwatashi et al. (2014), KIND1a/1b-cerulean locus integration 2) ARK1, cytoplasmic, plus ends of MTs 2) Arabidopsis root hairs 2) Eng and Wasteneys (2014), pARK1::ARK1-GFP Actin Related Protein2/3 (ARP2/3) complex PpARPC4, apically enriched cap P. patens protonemata Perroud and Quatrano (2006), 2xYFP locus integration Brick1 (brk1) PpBRK1, apically enriched cap P. patens protonemata Perroud and Quatrano (2008), 3xYFP locus integration Rho/Rac of Plants (ROP) 1) AtROP2, apical dome 1) Tobacco pollen tubes 1) Kost et al. (1999), LAT52::GFP-AtROP2 2) ROP1, apical dome 2) Arabidopsis root hairs 2) Jones et al. (2002), 35S::GFP-ROP2 3) AtROP1-PM, apical enrichment, cytoplasm 3) Tobacco pollen tubes 3) Hwang et al. (2005), LAT52::RIC4ƊC-GFP 4) PpROP2-PM, enrichment at cross walls 4 and 5) P. patens protonemata 4) Ito et al. (2014), HSP::cerulean-PpROP2 5) PpROP4, apical PM and cross walls 5) P. patens protonemata 5) Burkart et al. (2015), pROP4::GFP-ROP4coding sequence, locus replacement ROP Activating Proteins (RopGAPs) NtRopGAP1, subapical PM Tobacco pollen tubes Klahre and Kost (2006), Lat52::YFP-NtRopGAP1 ROP Interacting CRIB-Containing Protein (RIC) 1) AtRIC 3,10, cytosolic 1 to 4) Tobacco pollen tubes 1) Wu et al. (2001), Lat52::GFP-RIC n 2) AtRIC 1, 6, 7, apical PM, cytosolic 2) Wu et al. (2001), Lat52::GFP-RIC n 3) AtRIC9, PM 3) Wu et al. (2001), Lat52::GFP-RIC 9 4) AtRIC 2, 4, 5, apical PM 4) Wu et al. (2001), Lat52::GFP-RIC n ROP Guanine Dissociation Inhibitors (RopGDIs) NtRopGDI2, cytosolic Tobacco pollen tubes Klahre et al. (2006), Lat52::YFP-NtRopGDI2 ROP Guanine Exchange Factors (RopGEFs) 1) AtRopGEF1, apical PM 1 to 3) Tobacco pollen tubes 1) Gu et al. (2006), Lat52::GFP-RopGEF1 2) AtRopGEF 8, 9, 14, apical PM, cytoplasm 2) Gu et al. (2006), Lat52::GFP-RopGEF n 3) AtRopGEF12, cytoplasm 3) Gu et al. (2006), Lat52::GFP-RopGEF12 4) PpRopGEF3, cytoplasmic apical enrichment 4) P. patens protonemata 4) Ito et al. (2014), HSP::PpRopGEF3-cerulean Microtubule-Associated Protein18 1) Shank PM, inverted cone at tip 1) Arabidopsis pollen tubes 1) Zhu et al. (2013), pAtMAP18::AtMAP18-GFP 2) Shank of PM, apical cytoplasm 2) Arabidopsis root hairs 2) Kang et al. (2017), pAtMAP18::AtMAP18-mCherry End-Binding1 1) Cortical puncta, apical accumulation 1) P. patens protonemata 1) Hiwatashi et al. (2014), PpEB1b-mCitrine locus integration 2) Cytoplasmic puncta 2) Arabidopsis root hairs 2) Eng and Wasteneys (2014), 35S::AtEB1b-GFP Open in new tab Table II. Localization of cytoskeleton-binding proteins MTs, Microtubules; PM, plasma membrane. Protein . Localization . Species/System . Reference . Fimbrin 1) Apical and subapical F-actin structures 1) Lily pollen tubes 1) Su et al. (2012), α-LlFIM1 antibody 2) Apical filaments 2) Arabidopsis pollen tubes 2) Zhang et al. (2016), pFIM5::FIM5-GFP Villin 1) AtVLN3, thick subapical filaments 1) Arabidopsis root hairs 1) van der Honing et al. (2012), pAtVLN3::VLN3-GFP 2) AtVLN2, thick subapical filaments, fine apical filaments 2 and 3) Arabidopsis pollen tubes 2) Qu et al. (2013), pAtVLN2::VLN2genomic-GFP 3) AtVLN5, thick subapical filaments, fine apical filaments 3) Qu et al. (2013), pAtVLN5::VLN5genomic-GFP Actin Depolymerizing Factor (ADF)/Cofilin 1) Apical and subapical actin structures 1) Tobacco pollen tubes and grains 1) Chen et al. (2002), Lat52::NtADF-GFP 2) Apical structures 2) Lily pollen tubes 2) Lovy-Wheeler et al. (2006), α-lily ADF antibody 3) Cytoplasmic 3) P. patens protonemata 3) Augustine et al. (2008), α-PpADF polyclonal antibody Profilin Cytoplasmic Lily pollen grains and tubes Vidali and Hepler (1997), α-profilin polyclonal antibody Actin Interacting Protein1 1) Apical F-actin fringe 1) Lilium spp. pollen tubes 1) Lovy-Wheeler et al. (2006), α-lily AIP antibody 2) Cytoplasmic 2) P. patens protonemata 2) Augustine et al. (2011), AIP-mCherry locus integration Class I Formin LlFH1, apical vesicles, PM Lily pollen tubes Li et al. (2017), transient overexpression of LlFH1-GFP Class II Formin PpFor2a, apex, not on PM P. patens protonemata Vidali et al. (2009b), For2a 3xGFP locus integration Class VIII Myosin PpMyo8A, cortical puncta with apical accumulation, spindle and phragmoplast midzone P. patens protonemata Wu and Bezanilla (2014), pMaizeUbiquitin::Myo8A-3xGFP, in Ɗmyo8 a,b,c,d,e Class XI Myosin 1) PpMyo11a cytosolic with apical enrichment 1) P. patens protonemata 1) Vidali et al. (2010), pMaizeUbiquitin::3xGFP-Myo11a 2) AtMyo11k, apical cloud 2) Arabidopsis root hairs 2) Park and Nebenführ (2013), pMyo11K::YFP-Myo11K Kinesin 1) KIND1a,b, plus ends of MTs focused on the tip 1) P. patens protonemata 1) Hiwatashi et al. (2014), KIND1a/1b-cerulean locus integration 2) ARK1, cytoplasmic, plus ends of MTs 2) Arabidopsis root hairs 2) Eng and Wasteneys (2014), pARK1::ARK1-GFP Actin Related Protein2/3 (ARP2/3) complex PpARPC4, apically enriched cap P. patens protonemata Perroud and Quatrano (2006), 2xYFP locus integration Brick1 (brk1) PpBRK1, apically enriched cap P. patens protonemata Perroud and Quatrano (2008), 3xYFP locus integration Rho/Rac of Plants (ROP) 1) AtROP2, apical dome 1) Tobacco pollen tubes 1) Kost et al. (1999), LAT52::GFP-AtROP2 2) ROP1, apical dome 2) Arabidopsis root hairs 2) Jones et al. (2002), 35S::GFP-ROP2 3) AtROP1-PM, apical enrichment, cytoplasm 3) Tobacco pollen tubes 3) Hwang et al. (2005), LAT52::RIC4ƊC-GFP 4) PpROP2-PM, enrichment at cross walls 4 and 5) P. patens protonemata 4) Ito et al. (2014), HSP::cerulean-PpROP2 5) PpROP4, apical PM and cross walls 5) P. patens protonemata 5) Burkart et al. (2015), pROP4::GFP-ROP4coding sequence, locus replacement ROP Activating Proteins (RopGAPs) NtRopGAP1, subapical PM Tobacco pollen tubes Klahre and Kost (2006), Lat52::YFP-NtRopGAP1 ROP Interacting CRIB-Containing Protein (RIC) 1) AtRIC 3,10, cytosolic 1 to 4) Tobacco pollen tubes 1) Wu et al. (2001), Lat52::GFP-RIC n 2) AtRIC 1, 6, 7, apical PM, cytosolic 2) Wu et al. (2001), Lat52::GFP-RIC n 3) AtRIC9, PM 3) Wu et al. (2001), Lat52::GFP-RIC 9 4) AtRIC 2, 4, 5, apical PM 4) Wu et al. (2001), Lat52::GFP-RIC n ROP Guanine Dissociation Inhibitors (RopGDIs) NtRopGDI2, cytosolic Tobacco pollen tubes Klahre et al. (2006), Lat52::YFP-NtRopGDI2 ROP Guanine Exchange Factors (RopGEFs) 1) AtRopGEF1, apical PM 1 to 3) Tobacco pollen tubes 1) Gu et al. (2006), Lat52::GFP-RopGEF1 2) AtRopGEF 8, 9, 14, apical PM, cytoplasm 2) Gu et al. (2006), Lat52::GFP-RopGEF n 3) AtRopGEF12, cytoplasm 3) Gu et al. (2006), Lat52::GFP-RopGEF12 4) PpRopGEF3, cytoplasmic apical enrichment 4) P. patens protonemata 4) Ito et al. (2014), HSP::PpRopGEF3-cerulean Microtubule-Associated Protein18 1) Shank PM, inverted cone at tip 1) Arabidopsis pollen tubes 1) Zhu et al. (2013), pAtMAP18::AtMAP18-GFP 2) Shank of PM, apical cytoplasm 2) Arabidopsis root hairs 2) Kang et al. (2017), pAtMAP18::AtMAP18-mCherry End-Binding1 1) Cortical puncta, apical accumulation 1) P. patens protonemata 1) Hiwatashi et al. (2014), PpEB1b-mCitrine locus integration 2) Cytoplasmic puncta 2) Arabidopsis root hairs 2) Eng and Wasteneys (2014), 35S::AtEB1b-GFP Protein . Localization . Species/System . Reference . Fimbrin 1) Apical and subapical F-actin structures 1) Lily pollen tubes 1) Su et al. (2012), α-LlFIM1 antibody 2) Apical filaments 2) Arabidopsis pollen tubes 2) Zhang et al. (2016), pFIM5::FIM5-GFP Villin 1) AtVLN3, thick subapical filaments 1) Arabidopsis root hairs 1) van der Honing et al. (2012), pAtVLN3::VLN3-GFP 2) AtVLN2, thick subapical filaments, fine apical filaments 2 and 3) Arabidopsis pollen tubes 2) Qu et al. (2013), pAtVLN2::VLN2genomic-GFP 3) AtVLN5, thick subapical filaments, fine apical filaments 3) Qu et al. (2013), pAtVLN5::VLN5genomic-GFP Actin Depolymerizing Factor (ADF)/Cofilin 1) Apical and subapical actin structures 1) Tobacco pollen tubes and grains 1) Chen et al. (2002), Lat52::NtADF-GFP 2) Apical structures 2) Lily pollen tubes 2) Lovy-Wheeler et al. (2006), α-lily ADF antibody 3) Cytoplasmic 3) P. patens protonemata 3) Augustine et al. (2008), α-PpADF polyclonal antibody Profilin Cytoplasmic Lily pollen grains and tubes Vidali and Hepler (1997), α-profilin polyclonal antibody Actin Interacting Protein1 1) Apical F-actin fringe 1) Lilium spp. pollen tubes 1) Lovy-Wheeler et al. (2006), α-lily AIP antibody 2) Cytoplasmic 2) P. patens protonemata 2) Augustine et al. (2011), AIP-mCherry locus integration Class I Formin LlFH1, apical vesicles, PM Lily pollen tubes Li et al. (2017), transient overexpression of LlFH1-GFP Class II Formin PpFor2a, apex, not on PM P. patens protonemata Vidali et al. (2009b), For2a 3xGFP locus integration Class VIII Myosin PpMyo8A, cortical puncta with apical accumulation, spindle and phragmoplast midzone P. patens protonemata Wu and Bezanilla (2014), pMaizeUbiquitin::Myo8A-3xGFP, in Ɗmyo8 a,b,c,d,e Class XI Myosin 1) PpMyo11a cytosolic with apical enrichment 1) P. patens protonemata 1) Vidali et al. (2010), pMaizeUbiquitin::3xGFP-Myo11a 2) AtMyo11k, apical cloud 2) Arabidopsis root hairs 2) Park and Nebenführ (2013), pMyo11K::YFP-Myo11K Kinesin 1) KIND1a,b, plus ends of MTs focused on the tip 1) P. patens protonemata 1) Hiwatashi et al. (2014), KIND1a/1b-cerulean locus integration 2) ARK1, cytoplasmic, plus ends of MTs 2) Arabidopsis root hairs 2) Eng and Wasteneys (2014), pARK1::ARK1-GFP Actin Related Protein2/3 (ARP2/3) complex PpARPC4, apically enriched cap P. patens protonemata Perroud and Quatrano (2006), 2xYFP locus integration Brick1 (brk1) PpBRK1, apically enriched cap P. patens protonemata Perroud and Quatrano (2008), 3xYFP locus integration Rho/Rac of Plants (ROP) 1) AtROP2, apical dome 1) Tobacco pollen tubes 1) Kost et al. (1999), LAT52::GFP-AtROP2 2) ROP1, apical dome 2) Arabidopsis root hairs 2) Jones et al. (2002), 35S::GFP-ROP2 3) AtROP1-PM, apical enrichment, cytoplasm 3) Tobacco pollen tubes 3) Hwang et al. (2005), LAT52::RIC4ƊC-GFP 4) PpROP2-PM, enrichment at cross walls 4 and 5) P. patens protonemata 4) Ito et al. (2014), HSP::cerulean-PpROP2 5) PpROP4, apical PM and cross walls 5) P. patens protonemata 5) Burkart et al. (2015), pROP4::GFP-ROP4coding sequence, locus replacement ROP Activating Proteins (RopGAPs) NtRopGAP1, subapical PM Tobacco pollen tubes Klahre and Kost (2006), Lat52::YFP-NtRopGAP1 ROP Interacting CRIB-Containing Protein (RIC) 1) AtRIC 3,10, cytosolic 1 to 4) Tobacco pollen tubes 1) Wu et al. (2001), Lat52::GFP-RIC n 2) AtRIC 1, 6, 7, apical PM, cytosolic 2) Wu et al. (2001), Lat52::GFP-RIC n 3) AtRIC9, PM 3) Wu et al. (2001), Lat52::GFP-RIC 9 4) AtRIC 2, 4, 5, apical PM 4) Wu et al. (2001), Lat52::GFP-RIC n ROP Guanine Dissociation Inhibitors (RopGDIs) NtRopGDI2, cytosolic Tobacco pollen tubes Klahre et al. (2006), Lat52::YFP-NtRopGDI2 ROP Guanine Exchange Factors (RopGEFs) 1) AtRopGEF1, apical PM 1 to 3) Tobacco pollen tubes 1) Gu et al. (2006), Lat52::GFP-RopGEF1 2) AtRopGEF 8, 9, 14, apical PM, cytoplasm 2) Gu et al. (2006), Lat52::GFP-RopGEF n 3) AtRopGEF12, cytoplasm 3) Gu et al. (2006), Lat52::GFP-RopGEF12 4) PpRopGEF3, cytoplasmic apical enrichment 4) P. patens protonemata 4) Ito et al. (2014), HSP::PpRopGEF3-cerulean Microtubule-Associated Protein18 1) Shank PM, inverted cone at tip 1) Arabidopsis pollen tubes 1) Zhu et al. (2013), pAtMAP18::AtMAP18-GFP 2) Shank of PM, apical cytoplasm 2) Arabidopsis root hairs 2) Kang et al. (2017), pAtMAP18::AtMAP18-mCherry End-Binding1 1) Cortical puncta, apical accumulation 1) P. patens protonemata 1) Hiwatashi et al. (2014), PpEB1b-mCitrine locus integration 2) Cytoplasmic puncta 2) Arabidopsis root hairs 2) Eng and Wasteneys (2014), 35S::AtEB1b-GFP Open in new tab Actin Cytoskeleton Of the two cytoskeletal elements in plants, the actin cytoskeleton, which is discussed further in this issue (Szymanski and Staiger, 2018), is essential for tip growth in angiosperm pollen tubes, root hairs, and moss protonemata. To understand how actin contributes to cell expansion, it is important to determine the subcellular architecture of the F-actin arrays. However, across these systems, visualizing F-actin has been challenging. For example, in fixed cells, fluorescent phalloidin, a widely used probe for F-actin in many cell types, does not readily stain plant cells (discussed by Olyslaegers and Verbelen, 1998). Furthermore, phalloidin only interacts with a subset of F-actin arrays (Nishida et al., 1987) and stabilizes F-actin (Cooper, 1987). Thus, phalloidin often leads to an increase in bundled F-actin (Vidali et al., 2009a). Additionally, to date, GFP-actin fusions have not been successful in plants. Thus, for live-cell imaging, researchers have used fluorescent actin-binding probes, such as Lifeact (Riedl et al., 2008), talin (Kost et al., 1998), and the actin-binding domain of fimbrin (Sheahan et al., 2004). However, these probes also generate artifacts, particularly when highly expressed (Vidali et al., 2009a; van der Honing et al., 2011). Many of the issues arising from using F-actin probes stem from the fact that the binding of these probes to F-actin differs and often is affected by proteins that may be associated with the actin cytoskeleton (Table II). Thus, depending on the probe used and the relative amount of the probe, different subcellular structures can be observed. For example, in Arabidopsis (Arabidopsis thaliana) pollen tubes, fluorescent phalloidin staining revealed F-actin occupying the whole tip region, while Lifeact-GFP revealed pollen tubes with relatively little actin at the tip (Zhang et al., 2016). In root hairs, Ketelaar et al. (2003) visualized F-actin with antibodies or phalloidin. While both methods produced similar results in general, the details of the actin structure were quite different. In moss protonemata, F-actin imaging with fluorescent phalloidin (Vidali et al., 2009a) and Talin-GFP (Finka et al., 2007; Perroud and Quatrano, 2008), revealed thick apical bundles and a prominent apical accumulation. However, the expression of Lifeact-GFP at noninhibitory levels labeled finer actin filaments and provided more detail in the apical structures (Vidali et al., 2009a). Given the species-specific variation and the inherent challenges of imaging F-actin arrays, one must consider the conclusions made within the context of the visualization method and plant species used. Despite differences in F-actin structures between the probes used and the species of cell type examined (Stephan, 2017), it is nevertheless possible to recognize an emergence of commonalities. For example, most pollen tubes exhibit a fringe of short longitudinally oriented actin microfilaments located at the cell cortex near the cell tip, and long actin bundles are found along the length of the cell (Lovy-Wheeler et al., 2005; Fig. 1A). In growing pollen tubes expressing Lifeact-GFP, the fringe was observed to rapidly remodel and move forward with growth (Rounds et al., 2014). Additionally, when pollen tube growth was inhibited with cyanide, the apical fringe degraded more rapidly than the actin cables in the shank of the tube (Winship et al., 2016). In pollen grains, Vogler et al. (2015) showed an F-actin accumulation 180° from the future germination site. This accumulation translocates to the shank after growth initiates. Using a pollen-specific promoter, Actin3, to drive the expression of Lifeact-GFP, Jásik et al. (2016) reported fine F-actin at the tip and dynamic actin along the shank of germinating pollen tubes. Studies such as these demonstrate that pollen tubes have at least two distinct arrays: an apical array that is very dynamic and a subapical array that tends to have more bundled F-actin. Figure 1. Open in new tabDownload slide Apical cytoskeletal structures and ion gradients. A, Pollen tube, modeled after lily. Cortical F-actin (orange) forms a fringe overlapping with the OH− gradient and the outer reaches of the calcium gradient. Microtubules (blue) form an inverted cone that just intersects the actin fringe. B, Root hair, modeled after Arabidopsis. Short F-actin filaments fill the apex, overlapping with the highest levels of calcium and OH−. As both gradients taper off, actin bundles and microtubules are found toward the rear of the cell. C, Moss chloronemal tip, modeled after P. patens. Apical actin is enriched along the entire dome, and fine cortical actin is present along the length of the cell. Contiguous cytoplasmic microtubules originating from the nucleus (N) point toward the cell tip. D, Moss caulonemal tip, modeled after P. patens. The apical actin is a highly focused spot and also the target of polymerizing cytoplasmic microtubules. Figure 1. Open in new tabDownload slide Apical cytoskeletal structures and ion gradients. A, Pollen tube, modeled after lily. Cortical F-actin (orange) forms a fringe overlapping with the OH− gradient and the outer reaches of the calcium gradient. Microtubules (blue) form an inverted cone that just intersects the actin fringe. B, Root hair, modeled after Arabidopsis. Short F-actin filaments fill the apex, overlapping with the highest levels of calcium and OH−. As both gradients taper off, actin bundles and microtubules are found toward the rear of the cell. C, Moss chloronemal tip, modeled after P. patens. Apical actin is enriched along the entire dome, and fine cortical actin is present along the length of the cell. Contiguous cytoplasmic microtubules originating from the nucleus (N) point toward the cell tip. D, Moss caulonemal tip, modeled after P. patens. The apical actin is a highly focused spot and also the target of polymerizing cytoplasmic microtubules. F-actin structures in growing root hairs share aspects of pollen tube structures. Actin bundles are prominent along the shank of the root hair but do not invade the tip region (Fig. 1B). Rather than a fringe, fine actin filaments appear at the tip (Ketelaar, 2013). As with the apical actin structures in pollen tubes, the fine actin filaments in the apex of root hairs are dynamic, as they are more sensitive to depolymerizing drugs than actin cables in the shank (Ketelaar et al., 2003). In addition to vesicle guidance to the tip, it is thought that this fine F-actin functions to keep larger organelles out of the tip region (Emons and Ketelaar, 2009), thus maintaining the apical clear zone (Ketelaar, 2013). Using F-actin as a filter is reminiscent of what has been observed in pollen tubes (Hepler and Winship, 2015). Moss protonemata have two tip-growing cell types: chloronemata and caulonemata. Chloronemata are distinguished from caulonemata by growing more slowly and possessing perpendicular cell plates. Because caulonemata grow more quickly than chloronemata, F-actin architecture in living tip-growing cells has been analyzed predominantly in caulonemata. Moss protonemata exhibit an enrichment of F-actin at the very apex and fine cortical actin filaments along the length of the cell (Vidali et al., 2009a; Fig. 1, C and D). Bundles are apparent but are few and reside toward the rear of the cell, with subapical cells exhibiting more bundled F-actin. While an apical F-actin enrichment occurs in both cell types, their structures differ depending on the cell type. In chloronemata, apical F-actin localizes over the whole apical dome (Vidali et al., 2009a; Fig. 1C), whereas in caulonemata, F-actin exists as a focused apical spot (Vidali et al., 2010; Fig. 1D). Both the apical F-actin aggregation and the fine cortical F-actin are very dynamic, remodeling on the order of seconds (Vidali et al., 2010). Similar to the apical actin in root hairs, these structures in moss have been shown to be involved in vesicle guidance (Bibeau et al., 2017). In summary, all three tip-growing cells possess longitudinally oriented actin cables in their shanks. However, apical actin structures differ. Pollen tubes, at least in lily (Lilium longiflorum) and tobacco (Nicotiana tabacum), possess a cortical actin fringe, while moss caulonemata maintain a focused spot of F-actin. Root hairs, in contrast, have an array of fine actin at their tips. Importantly, though, all three tip structures are dynamic, which contrasts with the relatively stable F-actin in the subapical regions. Critically, it is the dynamic apical populations that are essential for tip growth (Vidali et al., 2001; Vazquez et al., 2014). Microtubule Cytoskeleton In contrast to the essential nature of F-actin, microtubule-depolymerizing drugs have little effect on angiosperm pollen tube germination and growth in vitro, suggesting that microtubules are not required. However, in both root hairs (Bibikova et al., 1999) and protonemata (Doonan et al., 1988), microtubules likely play a role in maintaining the orientation of polarity. When treated with microtubule-depolymerizing drugs, both cell types no longer grow straight. Instead, the cells tend to change direction and fork, generating two or more outgrowths (Doonan et al., 1988; Bibikova et al., 1999; Ketelaar et al., 2003; Sieberer et al., 2005; Finka et al., 2007). Regardless of their contribution to tip growth, striking microtubule arrays are evident in pollen tubes as well as root hairs and moss protonemata (Fig. 1). As opposed to F-actin visualization, which relies on actin-binding reagents, imaging the microtubule cytoskeleton is most commonly accomplished by labeling microtubules directly using antibodies to tubulin in fixed material or translational fusions of tubulin with a fluorescent protein in live material. As a result, microtubule imaging is less prone to the artifacts that plague F-actin visualization. In pollen tubes, microtubules are oriented parallel to the long axis of the cell. In both lily and tobacco pollen tubes, these long cortical microtubules are often coaligned with fine filaments, which appear to be F-actin based on immunogold labeling (Lancelle and Hepler, 1991). Additionally, lily pollen tubes possess a funnel array of microtubules at the interface between the apical clear zone and the shank (Lovy-Wheeler et al., 2005; Fig. 1A). In root hairs, microtubule organization depends on the stage of growth. Generally, root hairs have two populations of microtubules: cortical and endoplasmic (cytoplasmic). Cortical microtubules are observed in both elongating and mature root hairs (Fig. 1B). Whether cortical microtubules are visualized at the very apex is dependent on the method of fixation (Sieberer et al., 2005). The polarity of polymerizing cortical microtubules, however, is dependent on growth status. Growing root hairs have cortical microtubules that grow away from the nucleus and, thus, have baseward and tipward trajectories. Conversely, mature root hairs have exclusive tipward polymerization (Ambrose and Wasteneys, 2014). Similarly, endoplasmic microtubules are only observed in growing root hairs (Sieberer et al., 2005) and emanate from the nucleus (Van Bruaene et al., 2004; Fig. 1B). In moss protonemata, there are also two populations of microtubules: cytoplasmic and cortical. In the apical cell, the polarity of cytoplasmic microtubules between the nucleus and the cell apex is tipward, with an apical focus of microtubule plus ends (Hiwatashi et al., 2014). In contrast, the cortical microtubule array consists of shorter microtubules that lack polarity (Burkart et al., 2015; Nakaoka et al., 2015). Because microtubules appear to be required for directionality in root hairs and protonemata, one plausible hypothesis is that they spatially restrict specific F-actin populations mediating outgrowth (see Outstanding Questions). In support of this hypothesis,Hiwatashi et al. (2014) discovered that two microtubule-based motors, KINESIN FOR INTERDIGITATED MICROTUBULES1a (KIND1a) and KIND1b, regulate microtubule organization and protonemal growth. The kind1a,b knockout plants exhibit protonemata that do not grow straight but still have tipward-growing cytoplasmic microtubules. However, the apical microtubules are disorganized, suggesting that KIND1a and KIND1b focus apical microtubules in protonemata (Hiwatashi et al., 2014). Interestingly, in wild-type cells, cytoplasmic microtubules focus on the same location occupied by the apical actin spot (Fig. 1, C and D), suggesting that microtubules may interact directly with the apical F-actin structure. Myosin VIII is an actin-based motor protein that links microtubules to actin during cell division in protonemata (Wu and Bezanilla, 2014; Table II). Protonemata completely lacking myosin VIII (ƊmyoVIIIa,b,c,d,e) grow slowly and less straight than the wild type, particularly when grown under nutrient-deficient conditions (Wu and Bezanilla, 2014). Together, these studies suggest that an active link between actin and microtubules is critical for directionality during tip growth. Studies of MICROTUBULE ASSOCIATED PROTEIN18 (AtMAP18) also support a role for microtubule-actin interactions in tip growth. MAP18 binds both microtubules and F-actin (Wang et al., 2007; Zhu et al., 2013). In addition, MAP18 has calcium-dependent F-actin-severing activity (Zhu et al., 2013). Importantly, MAP18 knockdown pollen tubes cannot grow straight (Zhu et al., 2013), reminiscent of root hairs and protonemata treated with oryzalin (Doonan et al., 1988; Bibikova et al., 1999). Furthermore, F-actin arrays are altered in map18 pollen tubes: map18 pollen tubes no longer exhibit fine apical F-actin; instead, F-actin bundles invade the tip. AtMAP18-GFP localized to the plasma membrane and to an apical inverted cone pattern in map18 knockdown pollen tubes (Zhu et al., 2013; Table II). It is plausible that the location of MAP18 is determined by microtubules, and where microtubules interact with F-actin, the latter may be severed depending on the presence of calcium (Fig. 1). Interestingly, MAP18 localizes similarly in root hairs (Table II), and MAP18 knockdown plants have root hairs half the length of wild-type plants (Kang et al., 2017). To investigate MAP18’s function in root hairs, Kang et al. (2017) tested for genetic and physical interactions between MAP18 and Rho/Rac of Plants2 (ROP2), a small GTPase considered to be a master regulator of tip growth in root hairs (Jones et al., 2002). Kang et al. (2017) found that MAP18 and ROP2 are in the same genetic pathway and that they interact physically with MAP18, preferentially binding the inactive form of ROP2 and thereby promoting ROP2 activity. Kang et al. (2017) went on to show that MAP18 competes with the ROP2 guanine dissociation inhibitor for binding to ROP2. From these and earlier studies (Wang et al., 2007; Zhu et al., 2013), Kang et al. (2017) suggest that MAP18 functions through ROP2 to reinforce polarity and independently of ROP2 by modulating the cytoskeleton. IONS Modulation of the many processes and structures involved in tip growth, including the cytoskeleton, is achieved not only by regulator proteins but also by the cytoplasmic environment. Here, we highlight local ion concentrations, in particular calcium and protons, as drivers for these processes (Hepler, 2005, 2016; Michard et al., 2017). To that end, a tip-focused calcium gradient is particularly prominent in growing pollen tubes, where the high point localizes to the extreme apex of the pollen tube, directly appressed to the apical plasma membrane (Fig. 1A). Root hairs have a similarly tip-focused, albeit shallower, calcium gradient (Fig. 1B). To date, calcium in moss protonemata remains understudied. In an early report on dark-grown caulonemata injected with a calcium-sensitive fluorescent dye, Tucker et al. (2005) observed UV (340 and 380 nm light)-stimulated waves associated with both the apex and the base of apical cells. However, Tucker et al. (2005) did not observe a tip-focused calcium gradient, as seen in growing pollen tubes and root hairs. In root hairs and pollen tubes, many channels both on the plasma membrane as well as on organelles are likely to be involved in establishing and maintaining the apical calcium gradient. Patch-clamp assays have identified stretch-activated channels in pollen tubes (Dutta and Robinson, 2004), and their involvement in generating the apical calcium gradient seems plausible as a product of turgor-driven deformation of the apical plasma membrane. More recently, the Cyclic Nucleotide-Gated Channel18 (CNGC18) in Arabidopsis was implicated in tip growth, since CNGC18-null plants are male sterile (Gao et al., 2016; Table I). Investigating further, cngc18 pollen tubes were shown to have irregular calcium oscillations and have difficulty finding the ovule. Additionally, root hairs of CNGC14 knockdown and knockout plants are shorter and have altered calcium oscillation profiles (Zhang et al., 2017). Another class of transporters, the Glu receptor family (GLR), plays a role in modulating the calcium signal in pollen tubes. Atglr1.2-1 pollen tubes are deformed, and the amplitudes of calcium oscillations are diminished compared with the wild type (Michard et al., 2011; Table I). In addition to stretch-activated, CNG, and GLR channels, pharmacological assays on pollen suggest that F-actin (Wang et al., 2004) and voltage-regulated (Wu et al., 2011) calcium channels play a role in calcium homeostasis. Further study is needed to determine what complement of channels is required to establish the calcium gradients during tip growth (see Outstanding Questions). The calcium gradient may modulate growth by stimulating secretion (Winship et al., 2016) and/or regulating F-actin structures through actin-binding proteins, which we focus on here. Two well-established actin-binding proteins with calcium dependence are villin/gelsolin and profilin (Table II), which are prime candidates for playing a key role in establishing different actin arrays in root hairs and pollen tubes. While the actin-binding and bundling activities of both the gelsolin repeat domains and the C-terminal headpiece domain (Finidori et al., 1992) are calcium independent (Khurana et al., 2010), villin’s actin-severing activity is calcium dependent (Huang et al., 2004). Villin’s localization is primarily cytosolic, but it appears to bind to filaments along the shank of root hairs (van der Honing et al., 2012; Table II). It seems likely that apical villin would likely sever, not bundle, F-actin, given the tip-focused calcium gradient (Monshausen et al., 2008). In support of this, vln2vln5 pollen tubes have an enlarged apical actin structure (Qu et al., 2013). Taken together, villin is thought to be a major regulator of actin structures (Huang et al., 2015). Profilin binds to monomeric actin in a calcium-dependent manner (Kovar et al., 2000) and inhibits nucleation and polymerization. Pollen tubes of profilin mutants have aberrant (Qu et al., 2017) or absent (Liu et al., 2015) actin fringes. Transient knockdown of profilin with RNAi in moss demonstrated that protonemal growth is dependent on profilin (Vidali et al., 2007). Indeed, profilin knockdown plants are significantly less than half the size of controls, and cells grow isotropically (Vidali et al., 2007). The profilin-actin dimer is the substrate for formins: actin nucleators and elongators (Goode and Eck, 2007; Table II). Formins, which are not known to have calcium dependency, will nevertheless be affected by calcium concentrations because they require profilin-actin as a substrate. One can imagine, then, that profilin’s binding activity modulates the structure of the actin array via its interactions with various cellular nucleators of actin polymerization (Suarez et al., 2015). Pollen tubes and root hairs also possess proton or pH gradients. In growing lily pollen tubes, there is an alkaline band a few micrometers behind the tip, with a small acidic patch at the tip itself (Feijó et al., 1999; Lovy-Wheeler et al., 2006). It is noteworthy that tobacco H+-ATPase1-GFP, an enzyme responsible for creating proton gradients, localizes to the plasma membrane of the shank and an apical cytoplasmic inverted cone in tobacco pollen tubes (Certal et al., 2008), fitting well with the proton gradient (Fig. 1). Although pH gradients are difficult to image because of the rapid diffusion of protons, their importance to tip growth is apparent because modulating intracellular pH can change the direction of growth (Hu et al., 2017). Imaging both the cytosolic and extracellular environments of root hairs also has revealed localized gradients (Monshausen et al., 2007). While the pH gradient is not as steep as in pollen tubes, the gradient is present nonetheless and is coupled with root hair development (Bibikova et al., 1998). The calcium and pH gradients present at the cell apex of pollen tubes and root hairs are not static but oscillate. Indeed, analyses of these oscillations explore how ion gradients mechanistically link to tip growth. Initial studies on pollen tubes demonstrated that calcium gradient oscillations followed the oscillations in growth rate (Messerli et al., 2000; Cárdenas et al., 2008). By contrast, the apical alkaline band oscillation preceded the growth rate oscillation, virtually matching the oscillation in secretion of new wall material (Lovy-Wheeler et al., 2006). Conversely, studies on the extracellular pH in root hairs indicate that an elevation in alkalinity follows the peak in growth rate (Monshausen et al., 2007). While it is unclear how these studies fit together, the work on pollen tubes provides support for the idea that changes in pH have a close relationship with the process of growth. To explore the connection between calcium, pH gradients, and growth in pollen tubes, Winship et al. (2016) reversibly inhibited pollen tube growth with cyanide (Table I) and then followed the return of the calcium and pH gradients. Although the calcium gradient appeared before growth restarted, it was noticeably behind that of the alkaline band and the process of secretion itself. Indeed, the alkaline band reemerged even before the cyanide had been removed (Winship et al., 2016). These results provide impetus to the notion that pH changes are fundamentally involved in tip growth. Again, pH regulation of tip growth is thought to affect the actin cytoskeleton. In particular, the elevated pH (7.4), as found in the alkaline band, would stimulate the turnover of F-actin in the fringe in pollen tubes because Actin Depolymerizing Factor (ADF) optimally fragments F-actin at this pH (Bamburg, 1999; Hepler, 2016; Table II). It is additionally noteworthy that, in lily pollen tubes, ADF localizes to the region of the alkaline band and to the actin fringe (Lovy-Wheeler et al., 2006). Activation of ADF activity helps explain the constant turnover of the actin fringe as it keeps pace with elongation. Interestingly, recent work has uncovered a new calcium-ADF interaction. ADF1 from Malus domestica binds to and severs actin in a calcium-dependent manner (Yang et al., 2017), suggesting that dual regulation of ADF may be important for fine-tuning actin fringe turnover. In contrast to pollen tubes, ADF in moss protonemata is cytosolic (Augustine et al., 2008; Table II). However, ADF is absolutely essential for tip growth and F-actin dynamics in protonemata (Augustine et al., 2011), reinforcing the key role that dynamic actin plays in tip growth. While calcium oscillations in pollen tubes appear to have a single periodicity, quantification of calcium oscillations via light sheet microscopy and spectral analyses in root hairs revealed two concurrent periodicities (Candeo et al., 2017). The mechanism behind the concurrent periodicities in root hairs is not known (see Outstanding Questions). Two methods to generate different periodicities could include variable entry or variable sequestration. For entry, CNGC14 is a prime candidate for being one component of that oscillation profile, as cngc14 root hairs have altered oscillation profiles (Zhang et al., 2017). Mechanisms of sequestration are not as well understood. That being said, pollen tubes with suppressed expression of calreticulin, a resident endoplasmic reticulum protein with calcium sequestration properties, had aberrant cytosolic calcium levels (Suwińska et al., 2017), suggesting a role for calreticulin in calcium homeostasis. How calcium enters the endoplasmic reticulum of pollen tubes, however, is unclear. These works highlight the need to study the mechanism of calcium recycling in tip-growing systems (see Outstanding Questions). CELL WALL Plant cell shape, and thus growth, are both confined and defined by the cell wall. To understand how to change the cell wall, for example achieving a local wall weakening at the cell tip for tip growth, one must consider its composition. Tip-growing cells have polymers found in most plant cells, such as cellulose, hemicellulose, and pectins. However, immunochemical data show that the amounts and distribution of these polymers differ from typical plant cells, with cellulose being sparse, while pectin is considerably enriched at the tip in pollen tubes, root hairs, and protonemata (Bosch and Hepler, 2005; Berry et al., 2016). For pollen tubes, mounting evidence increasingly supports the view that pectins, either as deposited directly or following enzymatic modification, play a major role in controlling the rigidity of the wall (Caffall and Mohnen, 2009). In lily and tobacco pollen tubes, pectins are secreted in an oscillatory manner, with the increase of pectin anticipating the increase in growth rate (McKenna et al., 2009; Rounds et al., 2011). These observations have given rise to the idea that the intercalation or intussusception of wall material weakens the existing wall and allows for stress relaxation at the locus of secretion (Hepler et al., 2013). But further results in tobacco show that pectin methyl esterase (PME) is secreted with the same kinetics as the bulk pectin (McKenna et al., 2009). Thus, PME secretion anticipates growth. It is thought that PME increases the presence of free carboxyl residues, thereby enhancing calcium cross-linking and rigidifying the apical wall (Bosch and Hepler, 2005). This activity can explain the rigidity of the pollen tube shank cell wall. However, at the tube tip, a PME-induced stiffening would be contrary to the local weakening of the wall. An explanation for this paradox may come from work in nonpollen tube systems, where PME may weaken the existing cell wall (Peaucelle et al., 2011, 2012, 2015; Braidwood et al., 2014). The proposed model suggests that PME demethoxylates pectin, rendering the wall components more accessible to degrading enzymes, which, in turn, weakens the wall. Additionally, if PME removes methoxy esters in a block-wise manner, the resulting wall would be more tightly cross-linked by calcium. However, if the PME activity results in random demethoxylation, then the wall would not necessarily be strongly cross-linked. Regardless of the manner of demethoxylation, enzymatic cleavage results in the release of a proton, which locally acidifies the wall and contributes to cell wall loosening and extension. It is also possible that there could be a parallel increase in the PME inhibitor (Röckel et al., 2008), which could negate the esterase activity. Whether PME is active or inactive at the very apex is still unclear. Nevertheless, it is clear that the presence of PME correlates with local weakening, leading to apical cell wall extension. Glycoproteins also may contribute substantially to cell wall integrity and, thus, polarized growth. Extensin18 (EXT18), a Hyp-rich glycoprotein, appears to direct pollen tube structure and growth (Choudhary et al., 2015). EXT18-null mutants are less fertile, which may result in part from reduced pollen tube growth but also from an increase in pollen tube bursting (Table I). The bursting phenotype lends support to the idea that EXT18 provides structural support to the growing pollen tube at the tip or is involved in the communication between the wall and the cytoplasm. Open in new tabDownload slide Open in new tabDownload slide More attention has been given to arabinogalactan proteins (AGPs), which have been shown to modulate growth in different tip-growing cells (Tan et al., 2012). AGPs consist of branched chain carbohydrates linked to a peptide backbone, which may be attached to the plasma membrane through a glycosylphosphatidyl inositol anchor. While ubiquitous, seemingly involved in growth across plant cell types, their specific functions are poorly understood (Ellis et al., 2010). To elucidate the functions of AGPs, many studies have used Yariv phenylglycoside, a dye that binds and stains AGPs as well as blocks their function. At low concentrations (1 μm), Yariv reagent blocks cell extension in pollen tubes (Roy et al., 1998, 1999; Mollet et al., 2002) and moss protonemata (Lee et al., 2005; Table I). Interestingly, the Yariv reagent inhibits cell extension in pollen tubes but not secretion. An electron microscopic analysis revealed that Yariv-treated cells build up wall material in the apical periplasmic space (Roy et al., 1999). In contrast to other inhibitors such as caffeine or cyanide, which degrade the apical calcium gradient, pollen tubes treated with Yariv have only mildly diminished calcium levels at their tip (Roy et al., 1999; Table I). Continued secretion and the presence of the calcium gradient suggest that these activities are likely linked. Genetic approaches also have implicated AGPs in tip growth. In Arabidopsis, RNAi of AGP6 and AGP11 resulted in plants with reduced fertility, which is partly due to impaired pollen tube growth (Levitin et al., 2008; Table I). Similarly, moss protonemata AGP1 knockout plants have reduced protonemal cell length (Table I). In root hairs, AGPs are reduced markedly in the Arabidopsis reb1-1 mutant. The root epidermal trichoblasts of reb1-1 exhibit morphological defects, including bulging and disorganized cortical microtubules (Andème-Onzighi et al., 2002; Driouich and Baskin, 2008; Table I). Given that REB1 is part of a gene family involved in sugar metabolism, it seems likely that REB1 is involved in the synthesis of AGPs rather than regulating microtubule organization directly. However, compromised AGPs at the plasma membrane and in the cell wall may affect the organization of cortical microtubules (Andème-Onzighi et al., 2002), ultimately affecting the direction of cell expansion (Ketelaar et al., 2003). While it is clear that AGPs are a major player in cell wall expansion, the underlying mechanism is less straightforward. AGPs could control the assembly or incorporation of new material into the matrix of the cell wall. Alternatively, given that AGPs bind calcium with high affinity, periplasmic AGPs could serve as a calcium capacitor, creating a significant calcium reservoir (Lamport and Várnai, 2013; Lamport et al., 2014). Given that calcium ions are essential for cell wall integrity, it is not clear how the AGP/calcium capacitor modulates the wall. Perhaps the capacitor competes for calcium throughout the wall and works together with the stressed pectin residues to weaken the wall (Boyer, 2009). Alternatively, AGPs might directly facilitate the intercalation of new pectin monomers deep into the wall matrix, where its bonding activities will weaken existing wall bonds (Ray, 1992; Hepler et al., 2013). If the AGP calcium reservoir also is available for intracellular signaling, then it may serve to link wall status to the cytoplasm. CONCLUSION In this Update, we have called attention to advances in our understanding of tip growth in pollen tubes, root hairs, and moss protonemata. To establish a mechanistic understanding of tip growth is to elucidate how intracellular events cause a local weakening of the apical cell wall, a prerequisite for turgor-driven cell expansion. The weakening, of course, must be balanced with the deposition of new wall material, or else tip growth fails (Table I). As discussed, localized ion gradients almost certainly play pivotal roles in different processes. For example, the turnover of F-actin in the apex, thought to occur through the ion-dependent activities of actin-binding proteins (Table II), might facilitate the dynamics of exocytic and endocytic vesicles (Bibeau et al., 2017; see Outstanding Questions). It is unsurprising, then, that dynamic ion gradients and cytoskeletal elements occur in the same place: the site of polarized expansion (Fig. 1). Moving forward, it is imperative to understand how the cell wall signals to the intracellular machinery responsible for its expansion (see Outstanding Questions). By continuing to investigate both the similarities and differences across tip-growing systems, we gain deep insights into fundamental and derived processes that sum together in a tip-growing plant cell. LITERATURE CITED Ambrose C , Wasteneys GO ( 2014 ) Microtubule initiation from the nuclear surface controls cortical microtubule growth polarity and orientation in Arabidopsis thaliana . Plant Cell Physiol 55 : 1636 – 1645 Google Scholar Crossref Search ADS PubMed WorldCat Andème-Onzighi C , Sivaguru M, Judy-March J, Baskin TI, Driouich A ( 2002 ) The reb1-1 mutation of Arabidopsis alters the morphology of trichoblasts, the expression of arabinogalactan-proteins and the organization of cortical microtubules . Planta 215 : 949 – 958 Google Scholar Crossref Search ADS PubMed WorldCat Augustine RC , Pattavina KA, Tüzel E, Vidali L, Bezanilla M ( 2011 ) Actin interacting protein1 and actin depolymerizing factor drive rapid actin dynamics in Physcomitrella patens . Plant Cell 23 : 3696 – 3710 Google Scholar Crossref Search ADS PubMed WorldCat Augustine RC , Vidali L, Kleinman KP, Bezanilla M ( 2008 ) Actin depolymerizing factor is essential for viability in plants, and its phosphoregulation is important for tip growth . Plant J 54 : 863 – 875 Google Scholar Crossref Search ADS PubMed WorldCat Bamburg JR ( 1999 ) Proteins of the ADF/cofilin family: essential regulators of actin dynamics . Annu Rev Cell Dev Biol 15 : 185 – 230 Google Scholar Crossref Search ADS PubMed WorldCat Baumberger N , Steiner M, Ryser U, Keller B, Ringli C ( 2003 ) Synergistic interaction of the two paralogous Arabidopsis genes LRX1 and LRX2 in cell wall formation during root hair development . Plant J 35 : 71 – 81 Google Scholar Crossref Search ADS PubMed WorldCat Berry EA , Tran ML, Dimos CS, Budziszek MJ Jr , Scavuzzo-Duggan TR, Roberts AW ( 2016 ) Immuno and affinity cytochemical analysis of cell wall composition in the moss Physcomitrella patens . Front Plant Sci 7 : 248 Google Scholar Crossref Search ADS PubMed WorldCat Bibeau JP , Kingsley JL, Furt F, Tüzel E, Vidali L ( 2018 ) F-actin meditated focusing of vesicles at the cell tip is essential for polarized growth . Plant Physiol 176 : 352 – 363 Google Scholar Crossref Search ADS Google Preview WorldCat COPAC Bibikova TN , Blancaflor EB, Gilroy S ( 1999 ) Microtubules regulate tip growth and orientation in root hairs of Arabidopsis thaliana . Plant J 17 : 657 – 665 Google Scholar Crossref Search ADS PubMed WorldCat Bibikova TN , Jacob T, Dahse I, Gilroy S ( 1998 ) Localized changes in apoplastic and cytoplasmic pH are associated with root hair development in Arabidopsis thaliana . Development 125 : 2925 – 2934 Google Scholar Crossref Search ADS PubMed WorldCat Boisson-Dernier A , Roy S, Kritsas K, Grobei MA, Jaciubek M, Schroeder JI, Grossniklaus U ( 2009 ) Disruption of the pollen-expressed FERONIA homologs ANXUR1 and ANXUR2 triggers pollen tube discharge . Development 136 : 3279 – 3288 Google Scholar Crossref Search ADS PubMed WorldCat Bosch M , Hepler PK ( 2005 ) Pectin methylesterases and pectin dynamics in pollen tubes . Plant Cell 17 : 3219 – 3226 Google Scholar Crossref Search ADS PubMed WorldCat Boyer JS ( 2009 ) Cell wall biosynthesis and the molecular mechanism of plant enlargement . Funct Plant Biol 36 : 383 – 394 Google Scholar Crossref Search ADS PubMed WorldCat Braidwood L , Breuer C, Sugimoto K ( 2014 ) My body is a cage: mechanisms and modulation of plant cell growth . New Phytol 201 : 388 – 402 Google Scholar Crossref Search ADS PubMed WorldCat Burkart GM , Baskin TI, Bezanilla M ( 2015 ) A family of ROP proteins that suppresses actin dynamics, and is essential for polarized growth and cell adhesion . J Cell Sci 128 : 2553 – 2564 Google Scholar PubMed OpenURL Placeholder Text WorldCat Caffall KH , Mohnen D ( 2009 ) The structure, function, and biosynthesis of plant cell wall pectic polysaccharides . Carbohydr Res 344 : 1879 – 1900 Google Scholar Crossref Search ADS PubMed WorldCat Candeo A , Doccula FG, Valentini G, Bassi A, Costa A ( 2017 ) Light sheet fluorescence microscopy quantifies calcium oscillations in root hairs of Arabidopsis thaliana . Plant Cell Physiol 58 : 1161 – 1172 Google Scholar Crossref Search ADS PubMed WorldCat Cárdenas L , Lovy-Wheeler A, Kunkel JG, Hepler PK ( 2008 ) Pollen tube growth oscillations and intracellular calcium levels are reversibly modulated by actin polymerization . Plant Physiol 146 : 1611 – 1621 Google Scholar Crossref Search ADS PubMed WorldCat Certal AC , Almeida RB, Carvalho LM, Wong E, Moreno N, Michard E, Carneiro J, Rodriguéz-Léon J, Wu HM, Cheung AY, et al. ( 2008 ) Exclusion of a proton ATPase from the apical membrane is associated with cell polarity and tip growth in Nicotiana tabacum pollen tubes . Plant Cell 20 : 614 – 634 Google Scholar Crossref Search ADS PubMed WorldCat Chen CY , Wong EI, Vidali L, Estavillo A, Hepler PK, Wu HM, Cheung AY ( 2002 ) The regulation of actin organization by actin-depolymerizing factor in elongating pollen tubes . Plant Cell 14 : 2175 – 2190 Google Scholar Crossref Search ADS PubMed WorldCat Choudhary P , Saha P, Ray T, Tang Y, Yang D, Cannon MC ( 2015 ) EXTENSIN18 is required for full male fertility as well as normal vegetative growth in Arabidopsis . Front Plant Sci 6 : 553 Google Scholar Crossref Search ADS PubMed WorldCat Cosgrove DJ ( 2018 ) Diffuse growth of plant cell walls . Plant Physiol 176 : 16 – 27 Google Scholar Crossref Search ADS PubMed WorldCat Cooper JA ( 1987 ) Effects of cytochalasin and phalloidin on actin . J Cell Biol 105 : 1473 – 1478 Google Scholar Crossref Search ADS PubMed WorldCat Damineli DSC , Portes MT, Feijó JA ( 2017 ) One thousand and one oscillators at the pollen tube tip: the quest for a central pacemaker revisited . In Obermeyer G, Feijo J, eds, Pollen Tip Growth. Springer International Publishing , New York, NY , pp 391 – 413 Google Scholar Crossref Search ADS Google Preview WorldCat COPAC Doonan JH , Cove DJ, Lloyed CW ( 1988 ) Microtubules and microfilaments in tip growth: evidence that microtubules impose polarity on protonemal growth in Physcomitrella patens . J Cell Sci 89 : 533 – 540 Google Scholar Crossref Search ADS WorldCat Driouich A , Baskin TI ( 2008 ) Intercourse between cell wall and cytoplasm exemplified by arabinogalactan proteins and cortical microtubules . Am J Bot 95 : 1491 – 1497 Google Scholar Crossref Search ADS PubMed WorldCat Dutta R , Robinson KR ( 2004 ) Identification and characterization of stretch-activated ion channels in pollen protoplasts . Plant Physiol 135 : 1398 – 1406 Google Scholar Crossref Search ADS PubMed WorldCat Ellis M , Egelund J, Schultz CJ, Bacic A ( 2010 ) Arabinogalactan-proteins: key regulators at the cell surface? Plant Physiol 153 : 403 – 419 Google Scholar Crossref Search ADS PubMed WorldCat Emons AMC , Ketelaar T ( 2009 ) Intracellular organization: a prerequisite for root hair elongation and cell wall deposition . In Emons AMC, Ketelaar T, eds, Root Hairs. Springer , Berlin , pp 27 – 44 Google Scholar Crossref Search ADS Google Preview WorldCat COPAC Eng RC , Wasteneys GO ( 2014 ) The microtubule plus-end tracking protein ARMADILLO-REPEAT KINESIN1 promotes microtubule catastrophe in Arabidopsis . Plant Cell 26 : 3372 – 3386 Google Scholar Crossref Search ADS PubMed WorldCat Favery B , Ryan E, Foreman J, Linstead P, Boudonck K, Steer M, Shaw P, Dolan L ( 2001 ) KOJAK encodes a cellulose synthase-like protein required for root hair cell morphogenesis in Arabidopsis . Genes Dev 15 : 79 – 89 Google Scholar Crossref Search ADS PubMed WorldCat Feijó JA , Sainhas J, Hackett GR, Kunkel JG, Hepler PK ( 1999 ) Growing pollen tubes possess a constitutive alkaline band in the clear zone and a growth-dependent acidic tip . J Cell Biol 144 : 483 – 496 Google Scholar Crossref Search ADS PubMed WorldCat Finidori J , Friederich E, Kwiatkowski DJ, Louvard D ( 1992 ) In vivo analysis of functional domains from villin and gelsolin . J Cell Biol 116 : 1145 – 1155 Google Scholar Crossref Search ADS PubMed WorldCat Finka A , Schaefer DG, Saidi Y, Goloubinoff P, Zrÿd JP ( 2007 ) In vivo visualization of F-actin structures during the development of the moss Physcomitrella patens . New Phytol 174 : 63 – 76 Google Scholar Crossref Search ADS PubMed WorldCat Gao QF , Gu LL, Wang HQ, Fei CF, Fang X, Hussain J, Sun SJ, Dong JY, Liu H, Wang YF ( 2016 ) Cyclic nucleotide-gated channel 18 is an essential Ca2+ channel in pollen tube tips for pollen tube guidance to ovules in Arabidopsis . Proc Natl Acad Sci USA 113 : 3096 – 3101 Google Scholar Crossref Search ADS PubMed WorldCat Gibbon BC , Kovar DR, Staiger CJ ( 1999 ) Latrunculin B has different effects on pollen germination and tube growth . Plant Cell 11 : 2349 – 2363 Google Scholar Crossref Search ADS PubMed WorldCat Goode BL , Eck MJ ( 2007 ) Mechanism and function of formins in the control of actin assembly . Annu Rev Biochem 76 : 593 – 627 Google Scholar Crossref Search ADS PubMed WorldCat Gu Y , Li S, Lord EM, Yang Z ( 2006 ) Members of a novel class of Arabidopsis Rho guanine nucleotide exchange factors control Rho GTPase-dependent polar growth . Plant Cell 18 : 366 – 381 Google Scholar Crossref Search ADS PubMed WorldCat Gui CP , Dong X, Liu HK, Huang WJ, Zhang D, Wang SJ, Barberini ML, Gao XY, Muschietti J, McCormick S, et al. ( 2014 ) Overexpression of the tomato pollen receptor kinase LePRK1 rewires pollen tube growth to a blebbing mode . Plant Cell 26 : 3538 – 3555 Google Scholar Crossref Search ADS PubMed WorldCat Harries PA , Pan A, Quatrano RS ( 2005 ) Actin-related protein2/3 complex component ARPC1 is required for proper cell morphogenesis and polarized cell growth in Physcomitrella patens . Plant Cell 17 : 2327 – 2339 Google Scholar Crossref Search ADS PubMed WorldCat Hepler PK ( 2005 ) Calcium: a central regulator of plant growth and development . Plant Cell 17 : 2142 – 2155 Google Scholar Crossref Search ADS PubMed WorldCat Hepler PK ( 2016 ) The cytoskeleton and its regulation by calcium and protons . Plant Physiol 170 : 3 – 22 Google Scholar Crossref Search ADS PubMed WorldCat Hepler PK , Rounds CM, Winship LJ ( 2013 ) Control of cell wall extensibility during pollen tube growth . Mol Plant 6 : 998 – 1017 Google Scholar Crossref Search ADS PubMed WorldCat Hepler PK , Winship LJ ( 2015 ) The pollen tube clear zone: clues to the mechanism of polarized growth . J Integr Plant Biol 57 : 79 – 92 Google Scholar Crossref Search ADS PubMed WorldCat Hiwatashi Y , Sato Y, Doonan JH ( 2014 ) Kinesins have a dual function in organizing microtubules during both tip growth and cytokinesis in Physcomitrella patens . Plant Cell 26 : 1256 – 1266 Google Scholar Crossref Search ADS PubMed WorldCat Hoedemaekers K , Derksen J, Hoogstrate SW, Wolters-Arts M, Oh SA, Twell D, Mariani C, Rieu I ( 2015 ) BURSTING POLLEN is required to organize the pollen germination plaque and pollen tube tip in Arabidopsis thaliana . New Phytol 206 : 255 – 267 Google Scholar Crossref Search ADS PubMed WorldCat Hu C , Vogler H, Aellen M, Shamsudhin N, Jang B, Burri JT, Läubli N, Grossniklaus U, Pané S, Nelson BJ ( 2017 ) High precision, localized proton gradients and fluxes generated by a microelectrode device induce differential growth behaviors of pollen tubes . Lab Chip 17 : 671 – 680 Google Scholar Crossref Search ADS PubMed WorldCat Huang S , Blanchoin L, Chaudhry F, Franklin-Tong VE, Staiger CJ ( 2004 ) A gelsolin-like protein from Papaver rhoeas pollen (PrABP80) stimulates calcium-regulated severing and depolymerization of actin filaments . J Biol Chem 279 : 23364 – 23375 Google Scholar Crossref Search ADS PubMed WorldCat Huang S , Qu X, Zhang R ( 2015 ) Plant villins: versatile actin regulatory proteins . J Integr Plant Biol 57 : 40 – 49 Google Scholar Crossref Search ADS PubMed WorldCat Hwang JU , Gu Y, Lee YJ, Yang Z ( 2005 ) Oscillatory ROP GTPase activation leads the oscillatory polarized growth of pollen tubes . Mol Biol Cell 16 : 5385 – 5399 Google Scholar Crossref Search ADS PubMed WorldCat Ito K , Ren J, Fujita T ( 2014 ) Conserved function of Rho-related Rop/RAC GTPase signaling in regulation of cell polarity in Physcomitrella patens . Gene 544 : 241 – 247 Google Scholar Crossref Search ADS PubMed WorldCat Jásik J , Mičieta K, Siao W, Voigt B, Stuchlík S, Schmelzer E, Turňa J, Balu¡ka F ( 2016 ) Actin3 promoter reveals undulating F-actin bundles at shanks and dynamic F-actin meshworks at tips of tip-growing pollen tubes . Plant Signal Behav 11 : e1146845 Google Scholar Crossref Search ADS PubMed WorldCat Jones MA , Shen JJ, Fu Y, Li H, Yang Z, Grierson CS ( 2002 ) The Arabidopsis Rop2 GTPase is a positive regulator of both root hair initiation and tip growth . Plant Cell 14 : 763 – 776 Google Scholar Crossref Search ADS PubMed WorldCat Kang E , Zheng M, Zhang Y, Yuan M, Yalovsky S, Zhu L, Fu Y ( 2017 ) The microtubule-associated protein MAP18 affects ROP2 GTPase activity during root hair growth . Plant Physiol 174 : 202 – 222 Google Scholar Crossref Search ADS PubMed WorldCat Ketelaar T ( 2013 ) The actin cytoskeleton in root hairs: all is fine at the tip . Curr Opin Plant Biol 16 : 749 – 756 Google Scholar Crossref Search ADS PubMed WorldCat Ketelaar T , de Ruijter NCA, Emons AMC ( 2003 ) Unstable F-actin specifies the area and microtubule direction of cell expansion in Arabidopsis root hairs . Plant Cell 15 : 285 – 292 Google Scholar Crossref Search ADS PubMed WorldCat Khurana P , Henty JL, Huang S, Staiger AM, Blanchoin L, Staiger CJ ( 2010 ) Arabidopsis VILLIN1 and VILLIN3 have overlapping and distinct activities in actin bundle formation and turnover . Plant Cell 22 : 2727 – 2748 Google Scholar Crossref Search ADS PubMed WorldCat Klahre U , Becker C, Schmitt AC, Kost B ( 2006 ) Nt-RhoGDI2 regulates Rac/Rop signaling and polar cell growth in tobacco pollen tubes . Plant J 46 : 1018 – 1031 Google Scholar Crossref Search ADS PubMed WorldCat Klahre U , Kost B ( 2006 ) Tobacco RhoGTPase ACTIVATING PROTEIN1 spatially restricts signaling of RAC/Rop to the apex of pollen tubes . Plant Cell 18 : 3033 – 3046 Google Scholar Crossref Search ADS PubMed WorldCat Kost B , Lemichez E, Spielhofer P, Hong Y, Tolias K, Carpenter C, Chua NH ( 1999 ) Rac homologues and compartmentalized phosphatidylinositol 4,5-bisphosphate act in a common pathway to regulate polar pollen tube growth . J Cell Biol 145 : 317 – 330 Kost B , Spielhofer P, Chua NH ( 1998 ) A GFP-mouse talin fusion protein labels plant actin filaments in vivo and visualizes the actin cytoskeleton in growing pollen tubes . Plant J 16 : 393 – 401 Google Scholar Crossref Search ADS PubMed WorldCat Kovar DR , Drøbak BK, Staiger CJ ( 2000 ) Maize profilin isoforms are functionally distinct . Plant Cell 12 : 583 – 598 Google Scholar Crossref Search ADS PubMed WorldCat Kwon T , Sparks JA, Nakashima J, Allen SN, Tang Y, Blancaflor EB ( 2015 ) Transcriptional response of Arabidopsis seedlings during spaceflight reveals peroxidase and cell wall remodeling genes associated with root hair development . Am J Bot 102 : 21 – 35 Google Scholar Crossref Search ADS PubMed WorldCat Lamport DTA , Várnai P ( 2013 ) Periplasmic arabinogalactan glycoproteins act as a calcium capacitor that regulates plant growth and development . New Phytol 197 : 58 – 64 Google Scholar Crossref Search ADS PubMed WorldCat Lamport DTA , Varnai P, Seal CE ( 2014 ) Back to the future with the AGP-Ca2+ flux capacitor . Ann Bot 114 : 1069 – 1085 Google Scholar Crossref Search ADS PubMed WorldCat Lancelle SA , Hepler PK ( 1991 ) Association of actin with cortical microtubules revealed by immunogold localization in Nicotiana pollen tubes . Protoplasma 165 : 167 – 172 Google Scholar Crossref Search ADS WorldCat Lee KJ , Sakata Y, Mau SL, Pettolino F, Bacic A, Quatrano RS, Knight CD, Knox JP ( 2005 ) Arabinogalactan proteins are required for apical cell extension in the moss Physcomitrella patens . Plant Cell 17 : 3051 – 3065 Google Scholar Crossref Search ADS PubMed WorldCat Levitin B , Richter D, Markovich I, Zik M ( 2008 ) Arabinogalactan proteins 6 and 11 are required for stamen and pollen function in Arabidopsis . Plant J 56 : 351 – 363 Google Scholar Crossref Search ADS PubMed WorldCat Li S , Dong H, Pei W, Liu C, Zhang S, Sun T, Xue X, Ren H ( 2017 ) LlFH1-mediated interaction between actin fringe and exocytic vesicles is involved in pollen tube tip growth . New Phytol 214 : 745 – 761 Google Scholar Crossref Search ADS PubMed WorldCat Liu X , Qu X, Jiang Y, Chang M, Zhang R, Wu Y, Fu Y, Huang S ( 2015 ) Profilin regulates apical actin polymerization to control polarized pollen tube growth . Mol Plant 8 : 1694 – 1709 Google Scholar Crossref Search ADS PubMed WorldCat Lovy-Wheeler A , Kunkel JG, Allwood EG, Hussey PJ, Hepler PK ( 2006 ) Oscillatory increases in alkalinity anticipate growth and may regulate actin dynamics in pollen tubes of lily . Plant Cell 18 : 2182 – 2193 Google Scholar Crossref Search ADS PubMed WorldCat Lovy-Wheeler A , Wilsen KL, Baskin TI, Hepler PK ( 2005 ) Enhanced fixation reveals the apical cortical fringe of actin filaments as a consistent feature of the pollen tube . Planta 221 : 95 – 104 Google Scholar Crossref Search ADS PubMed WorldCat Malho R , Trewavas AJ ( 1996 ) Localized apical increases of cytosolic free calcium control pollen tube orientation . Plant Cell 8 : 1935 – 1949 Google Scholar Crossref Search ADS PubMed WorldCat McKenna ST , Kunkel JG, Bosch M, Rounds CM, Vidali L, Winship LJ, Hepler PK ( 2009 ) Exocytosis precedes and predicts the increase in growth in oscillating pollen tubes . Plant Cell 21 : 3026 – 3040 Google Scholar Crossref Search ADS PubMed WorldCat Mendrinna A , Persson S ( 2015 ) Root hair growth: it’s a one way street . F1000Prime Rep 7 : 23 Google Scholar Crossref Search ADS PubMed WorldCat Messerli MA , Créton R, Jaffe LF, Robinson KR ( 2000 ) Periodic increases in elongation rate precede increases in cytosolic Ca2+ during pollen tube growth . Dev Biol 222 : 84 – 98 Google Scholar Crossref Search ADS PubMed WorldCat Michard E , Lima PT, Borges F, Silva AC, Portes MT, Carvalho JE, Gilliham M, Liu LH, Obermeyer G, Feijó JA ( 2011 ) Glutamate receptor-like genes form Ca2+ channels in pollen tubes and are regulated by pistil D-serine . Science 332 : 434 – 437 Google Scholar Crossref Search ADS PubMed WorldCat Michard E , Simon AA, Tavares B, Wudick MM, Feijó JA ( 2017 ) Signaling with ions: the keystone for apical cell growth and morphogenesis in pollen tubes . Plant Physiol 173 : 91 – 111 Google Scholar Crossref Search ADS PubMed WorldCat Mollet JC , Kim S, Jauh GY, Lord EM ( 2002 ) Arabinogalactan proteins, pollen tube growth, and the reversible effects of Yariv phenylglycoside . Protoplasma 219 : 89 – 98 Google Scholar Crossref Search ADS PubMed WorldCat Monshausen GB , Bibikova TN, Messerli MA, Shi C, Gilroy S ( 2007 ) Oscillations in extracellular pH and reactive oxygen species modulate tip growth of Arabidopsis root hairs . Proc Natl Acad Sci USA 104 : 20996 – 21001 Google Scholar Crossref Search ADS PubMed WorldCat Monshausen GB , Messerli MA, Gilroy S ( 2008 ) Imaging of the Yellow Cameleon 3.6 indicator reveals that elevations in cytosolic Ca2+ follow oscillating increases in growth in root hairs of Arabidopsis . Plant Physiol 147 : 1690 – 1698 Google Scholar Crossref Search ADS PubMed WorldCat Nakaoka Y , Kimura A, Tani T, Goshima G ( 2015 ) Cytoplasmic nucleation and atypical branching nucleation generate endoplasmic microtubules in Physcomitrella patens . Plant Cell 27 : 228 – 242 Google Scholar Crossref Search ADS PubMed WorldCat Nishida E , Iida K, Yonezama N, Koyasu S, Yahara I, Sakai H ( 1987 ) Cofilin is a component of intranuclear and cytoplasmic actin rods induced in cultured cells . Proc Natl Acad Sci USA 84 : 5262 – 5266 Olyslaegers G , Verbelen JP ( 1998 ) Improved staining of F-actin and co-localization of mitochondria in plant cells . J Microsc 192 : 73 – 77 Google Scholar Crossref Search ADS WorldCat Park E , Nebenführ A ( 2013 ) Myosin XIK of Arabidopsis thaliana accumulates at the root hair tip and is required for fast root hair growth . PLoS ONE 8 : e76745 Google Scholar Crossref Search ADS PubMed WorldCat Peaucelle A , Braybrook S, Höfte H ( 2012 ) Cell wall mechanics and growth control in plants: the role of pectins revisited . Front Plant Sci 3 : 121 Google Scholar Crossref Search ADS PubMed WorldCat Peaucelle A , Braybrook SA, Le Guillou L, Bron E, Kuhlemeier C, Höfte H ( 2011 ) Pectin-induced changes in cell wall mechanics underlie organ initiation in Arabidopsis . Curr Biol 21 : 1720 – 1726 Google Scholar Crossref Search ADS PubMed WorldCat Peaucelle A , Wightman R, Höfte H ( 2015 ) The control of growth symmetry breaking in the Arabidopsis hypocotyl . Curr Biol 25 : 1746 – 1752 Google Scholar Crossref Search ADS PubMed WorldCat Perroud PF , Quatrano RS ( 2006 ) The role of ARPC4 in tip growth and alignment of the polar axis in filaments of Physcomitrella patens . Cell Motil Cytoskeleton 63 : 162 – 171 Google Scholar Crossref Search ADS PubMed WorldCat Perroud PF , Quatrano RS ( 2008 ) BRICK1 is required for apical cell growth in filaments of the moss Physcomitrella patens but not for gametophore morphology . Plant Cell 20 : 411 – 422 Google Scholar Crossref Search ADS PubMed WorldCat Qu X , Zhang H, Xie Y, Wang J, Chen N, Huang S ( 2013 ) Arabidopsis villins promote actin turnover at pollen tube tips and facilitate the construction of actin collars . Plant Cell 25 : 1803 – 1817 Google Scholar Crossref Search ADS PubMed WorldCat Qu X , Zhang R, Zhang M, Diao M, Xue Y, Huang S ( 2017 ) Organizational innovation of apical actin filaments drives rapid pollen tube growth and turning . Mol Plant 10 : 930 – 947 Google Scholar Crossref Search ADS PubMed WorldCat Ray PM ( 1992 ) Mechanisms of wall loosening for cell growth . Current Topics in Plant Biochemistry and Physiology 11 : 18 – 41 Riedl J , Crevenna AH, Kessenbrock K, Yu JH, Neukirchen D, Bista M, Bradke F, Jenne D, Holak TA, Werb Z, et al. ( 2008 ) Lifeact: a versatile marker to visualize F-actin . Nat Methods 5 : 605 – 607 Google Scholar Crossref Search ADS PubMed WorldCat Röckel N , Wolf S, Kost B, Rausch T, Greiner S ( 2008 ) Elaborate spatial patterning of cell-wall PME and PMEI at the pollen tube tip involves PMEI endocytosis, and reflects the distribution of esterified and de-esterified pectins . Plant J 53 : 133 – 143 Google Scholar Crossref Search ADS PubMed WorldCat Rounds CM , Hepler PK, Winship LJ ( 2014 ) The apical actin fringe contributes to localized cell wall deposition and polarized growth in the lily pollen tube . Plant Physiol 166 : 139 – 151 Google Scholar Crossref Search ADS PubMed WorldCat Rounds CM , Lubeck E, Hepler PK, Winship LJ ( 2011 ) Propidium iodide competes with Ca2+ to label pectin in pollen tubes and Arabidopsis root hairs . Plant Physiol 157 : 175 – 187 Google Scholar Crossref Search ADS PubMed WorldCat Roy S , Jauh GY, Hepler PK, Lord EM ( 1998 ) Effects of Yariv phenylglycoside on cell wall assembly in the lily pollen tube . Planta 204 : 450 – 458 Google Scholar Crossref Search ADS PubMed WorldCat Roy SJ , Holdaway-Clarke TL, Hackett GR, Kunkel JG, Lord EM, Hepler PK ( 1999 ) Uncoupling secretion and tip growth in lily pollen tubes: evidence for the role of calcium in exocytosis . Plant J 19 : 379 – 386 Google Scholar Crossref Search ADS PubMed WorldCat Sheahan MB , Staiger CJ, Rose RJ, McCurdy DW ( 2004 ) A green fluorescent protein fusion to actin-binding domain 2 of Arabidopsis fimbrin highlights new features of a dynamic actin cytoskeleton in live plant cells . Plant Physiol 136 : 3968 – 3978 Google Scholar Crossref Search ADS PubMed WorldCat Sieberer BJ , Ketelaar T, Esseling JJ, Emons AMC ( 2005 ) Microtubules guide root hair tip growth . New Phytol 167 : 711 – 719 Google Scholar Crossref Search ADS PubMed WorldCat Stephan OOH ( 2017 ) Actin fringes of polar cell growth . J Exp Bot 68 : 3303 – 3320 Google Scholar Crossref Search ADS PubMed WorldCat Su H , Zhu J, Cai C, Pei W, Wang J, Dong H, Ren H ( 2012 ) FIMBRIN1 is involved in lily pollen tube growth by stabilizing the actin fringe . Plant Cell 24 : 4539 – 4554 Google Scholar Crossref Search ADS PubMed WorldCat Suarez C , Carroll RT, Burke TA, Christensen JR, Bestul AJ, Sees JA, James ML, Sirotkin V, Kovar DR ( 2015 ) Profilin regulates F-actin network homeostasis by favoring formin over Arp2/3 complex . Dev Cell 32 : 43 – 53 Google Scholar Crossref Search ADS PubMed WorldCat Suwińska A , Wasąg P, Zakrzewski P, Lenartowska M, Lenartowski R ( 2017 ) Calreticulin is required for calcium homeostasis and proper pollen tube tip growth in Petunia . Planta 245 : 909 – 926 Google Scholar Crossref Search ADS PubMed WorldCat Synek L , Vuka¡inović N, Kulich I, Hála M, Aldorfová K, Fendrych M, Žárský V ( 2017 ) EXO70C2 is a key regulatory factor for optimal tip growth of pollen . Plant Physiol 174 : 223 – 240 Google Scholar Crossref Search ADS PubMed WorldCat Szymanski D , Staiger CJ ( 2018 ) The actin cytoskeleton: functional arrays for cytoplasmic organization and cell shape control . Plant Physiol 176 : 106 – 118 Google Scholar Crossref Search ADS PubMed WorldCat Tan L , Showalter AM, Egelund J, Hernandez-Sanchez A, Doblin MS, Bacic A ( 2012 ) Arabinogalactan-proteins and the research challenges for these enigmatic plant cell surface proteoglycans . Front Plant Sci 3 : 140 Google Scholar Crossref Search ADS PubMed WorldCat Tan X , Cao K, Liu F, Li Y, Li P, Gao C, Ding Y, Lan Z, Shi Z, Rui Q, et al. ( 2016 ) Arabidopsis COG complex subunits COG3 and COG8 modulate Golgi morphology, vesicle trafficking homeostasis and are essential for pollen tube growth . PLoS Genet 12 : e1006140 Google Scholar Crossref Search ADS PubMed WorldCat Tucker EB , Lee M, Alli S, Sookhdeo V, Wada M, Imaizumi T, Kasahara M, Hepler PK ( 2005 ) UV-A induces two calcium waves in Physcomitrella patens . Plant Cell Physiol 46 : 1226 – 1236 Google Scholar Crossref Search ADS PubMed WorldCat Van Bruaene N , Joss G, Van Oostveldt P ( 2004 ) Reorganization and in vivo dynamics of microtubules during Arabidopsis root hair development . Plant Physiol 136 : 3905 – 3919 Google Scholar Crossref Search ADS PubMed WorldCat van der Honing HS , Kieft H, Emons AMC, Ketelaar T ( 2012 ) Arabidopsis VILLIN2 and VILLIN3 are required for the generation of thick actin filament bundles and for directional organ growth . Plant Physiol 158 : 1426 – 1438 Google Scholar Crossref Search ADS PubMed WorldCat van der Honing HS , van Bezouwen LS, Emons AMC, Ketelaar T ( 2011 ) High expression of Lifeact in Arabidopsis thaliana reduces dynamic reorganization of actin filaments but does not affect plant development . Cytoskeleton (Hoboken) 68 : 578 – 587 Google Scholar Crossref Search ADS PubMed WorldCat Vazquez LAB , Sanchez R, Hernández-Barrera A, Zepeda-Jazo I, Sanchez F, Quinto C, Torres LC ( 2014 ) Actin polymerization drives polar growth in Arabidopsis root hair cells . Plant Signal Behav 9 : e29401 – e29404 Google Scholar Crossref Search ADS PubMed WorldCat Vidali L , Augustine RC, Kleinman KP, Bezanilla M ( 2007 ) Profilin is essential for tip growth in the moss Physcomitrella patens . Plant Cell 19 : 3705 – 3722 Google Scholar Crossref Search ADS PubMed WorldCat Vidali L , Burkart GM, Augustine RC, Kerdavid E, Tüzel E, Bezanilla M ( 2010 ) Myosin XI is essential for tip growth in Physcomitrella patens . Plant Cell 22 : 1868 – 1882 Google Scholar Crossref Search ADS PubMed WorldCat Vidali L , Hepler PK ( 1997 ) Characterization and localization of profilin in pollen grains and tubes of Lilium longiflorum . Cell Motil Cytoskeleton 36 : 323 – 338 Google Scholar Crossref Search ADS PubMed WorldCat Vidali L , McKenna ST, Hepler PK ( 2001 ) Actin polymerization is essential for pollen tube growth . Mol Biol Cell 12 : 2534 – 2545 Google Scholar Crossref Search ADS PubMed WorldCat Vidali L , Rounds CM, Hepler PK, Bezanilla M ( 2009 a ) Lifeact-mEGFP reveals a dynamic apical F-actin network in tip growing plant cells . PLoS ONE 4 : e5744 Google Scholar Crossref Search ADS PubMed WorldCat Vidali L , van Gisbergen PAC, Guérin C, Franco P, Li M, Burkart GM, Augustine RC, Blanchoin L, Bezanilla M ( 2009 b ) Rapid formin-mediated actin-filament elongation is essential for polarized plant cell growth . Proc Natl Acad Sci USA 106 : 13341 – 13346 Google Scholar Crossref Search ADS PubMed WorldCat Vogler F , Konrad SSA, Sprunck S ( 2015 ) Knockin’ on pollen’s door: live cell imaging of early polarization events in germinating Arabidopsis pollen . Front Plant Sci 6 : 246 Google Scholar Crossref Search ADS PubMed WorldCat Wang X , Zhu L, Liu B, Wang C, Jin L, Zhao Q, Yuan M ( 2007 ) Arabidopsis MICROTUBULE-ASSOCIATED PROTEIN18 functions in directional cell growth by destabilizing cortical microtubules . Plant Cell 19 : 877 – 889 Google Scholar Crossref Search ADS PubMed WorldCat Wang YF , Fan LM, Zhang WZ, Zhang W, Wu WH ( 2004 ) Ca2+-permeable channels in the plasma membrane of Arabidopsis pollen are regulated by actin microfilaments . Plant Physiol 136 : 3892 – 3904 Google Scholar Crossref Search ADS PubMed WorldCat Winship LJ , Rounds C, Hepler PK ( 2016 ) Perturbation analysis of calcium, alkalinity and secretion during growth of lily pollen tubes . Plants (Basel) 6 : 3 Google Scholar Crossref Search ADS WorldCat Wu G , Gu Y, Li S, Yang Z ( 2001 ) A genome-wide analysis of Arabidopsis Rop-interactive CRIB motif-containing proteins that act as Rop GTPase targets . Plant Cell 13 : 2841 – 2856 Google Scholar PubMed OpenURL Placeholder Text WorldCat Wu J , Qu H, Jin C, Shang Z, Wu J, Xu G, Gao Y, Zhang S ( 2011 ) cAMP activates hyperpolarization-activated Ca2+ channels in the pollen of Pyrus pyrifolia . Plant Cell Rep 30 : 1193 – 1200 Google Scholar Crossref Search ADS PubMed WorldCat Wu SZ , Bezanilla M ( 2014 ) Myosin VIII associates with microtubule ends and together with actin plays a role in guiding plant cell division . eLife 3 : 3895 Google Scholar OpenURL Placeholder Text WorldCat Yang Q , Wang S, Wu C, Zhang Q, Zhang Y, Chen Q, Li Y, Hao L, Gu Z, Li W, et al. ( 2017 ) Malus domestica ADF1 severs actin filaments in growing pollen tubes . Funct Plant Biol 44 : 455 – 463 Google Scholar Crossref Search ADS PubMed WorldCat Zhang M , Zhang R, Qu X, Huang S ( 2016 ) Arabidopsis FIM5 decorates apical actin filaments and regulates their organization in the pollen tube . J Exp Bot 67 : 3407 – 3417 Google Scholar Crossref Search ADS PubMed WorldCat Zhang S , Pan Y, Tian W, Dong M, Zhu H, Luan S, Li L ( 2017 ) Arabidopsis CNGC14 mediates calcium influx required for tip growth in root hairs . Mol Plant 10 : 1004 – 1006 Google Scholar Crossref Search ADS PubMed WorldCat Zhu L , Zhang Y, Kang E, Xu Q, Wang M, Rui Y, Liu B, Yuan M, Fu Y ( 2013 ) MAP18 regulates the direction of pollen tube growth in Arabidopsis by modulating F-actin organization . Plant Cell 25 : 851 – 867 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 M.B. is supported by a grant from the National Science Foundation (grant no. MCB-1330171) and the David and Lucille Packard Foundation. C.S.B. received support from the Plant Biology Graduate Program at the University of Massachusetts. 2 Address correspondence to [email protected]. C.S.B., P.K.H., and M.B. wrote the article; C.S.B. drew the figure and compiled the information in the tables. [OPEN] Articles can be viewed without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.17.01466 © 2018 American Society of Plant Biologists. All Rights Reserved. © The Author(s) 2018. Published by Oxford University Press on behalf of American Society of Plant Biologists. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.

Bascom, Carlisle S.; Hepler, Peter K.; Bezanilla, Magdalena

doi: 10.1104/pp.17.01466pmid: 29138353

Bidhendi, Amir J.; Geitmann, Anja

doi: 10.1104/pp.17.01684pmid: 29229695

Plant cells come in a striking variety of different shapes. Shape formation in plant cells is controlled through modulation of the cell wall polymers and propelled by the turgor pressure. Understanding the shaping aspects of plant cells requires knowledge of the molecular players and the biophysical conditions under which they operate. Mechanical modeling has emerged as a useful tool to correlate cell wall structure, composition, and mechanics with cell and organ shape. The finite element method is a powerful numerical approach employed to solve problems in continuum mechanics. This Update critically analyzes studies that have used finite element analysis for the mechanical modeling of plant cells. Focus is on models involving single cell morphogenesis or motion. Model design, validation, and predictive power are analyzed in detail to open future avenues in the field. The cell wall, a polysaccharide-rich extracellular matrix, gives plant cells their shape at the expense of constraining their growth and movement. All cellular growth processes and shape changes involve a deformation of this extracellular matrix and are controlled by it. This control is exerted by modulating the mechanical properties of the matrix, which, in turn, are regulated by the polymers present in the wall and the state of linkages between them. The main polysaccharides of the primary cell wall are pectins, cellulose microfibrils, and xyloglucans (Bidhendi and Geitmann, 2016; Cosgrove, 2016). Cellulose microfibrils are recognized as the primary load-bearing component limiting cellular expansion (Baskin, 2005; Geitmann and Ortega, 2009). However, an increasing amount of evidence points at pivotal functions of pectins and hemicelluloses in defining the mechanics of the cell wall (Parre and Geitmann, 2005; Peaucelle et al., 2011; Palin and Geitmann, 2012; Braybrook and Peaucelle, 2013; Amsbury et al., 2016; Bidhendi and Geitmann, 2016; Torode et al., 2017). To understand how modulation of the plant cell wall affects and regulates the change of cell shape, the biomechanical context must be considered; for instance, see the Update in this issue on wall structure, mechanics, and growth (Cosgrove, 2018) or previous reviews (Geitmann and Ortega, 2009; Bidhendi and Geitmann, 2016). Open in new tabDownload slide Open in new tabDownload slide Biological experimentation with the goal to identify the crucial players in determining cell mechanics is challenging. Mutational or pharmacological modifications of the cell wall biochemistry often result in pleiotropic effects through feedback mechanisms that alter other cellular processes. Therefore, mechanical modeling has proven useful to guide biological experimentation by focusing on mechanical aspects of the behavior. Most modeling approaches in plant cell mechanics are based on the premise that the cell wall is a deformable material and that the deforming force is the turgor pressure, uniformly applied within the compartment of a single cell. This concept applies both to irreversible shape changes (cell growth) and reversible shape changes (stress generation or turgor-regulated motion). Since turgor is a scalar, for nonspherical cell shapes to develop during differentiation, the cell wall mechanical behavior must differ between subcellular regions. This can be achieved through the variation of wall thickness or heterogenous distribution of the material properties (Green, 1962; Sanati Nezhad and Geitmann, 2015). The mechanical aspects of shaping or deformation processes can be explored using a variety of mathematical approaches (Dyson and Jensen, 2010). The finite element (FE) method is one of the available computational techniques particularly suitable for the analysis of problems in continuum mechanics with a high degree of geometrical details or material complexity (Box 1). This Update analyzes examples in which this numerical tool is applied to evaluate the growth and elastic deformations of individual plant cells. Open in new tabDownload slide Open in new tabDownload slide The uses of FE modeling for cell or tissue studies can be categorized as forward or inverse approaches. The forward use of a model describes a deformation behavior, reversible or irreversible, inherent to the cell, such as a growth or shaping process. The purpose is to predict or explain the mechanical behavior arising from wall properties and turgor pressure (Fig. 1). Models used for an inverse approach are employed for the identification of material parameters from experiments such as indentation measurements (Bolduc et al., 2006; Bidhendi and Korhonen, 2012; Forouzesh et al., 2013; Sanati Nezhad et al., 2013). In this Update, we take a critical look at selected forward modeling studies of single plant cells (Fig. 1). Figure 1. Open in new tabDownload slide A, A closed cylindrical shell with hemispherical caps generated by the rotation of a line (orange) around a symmetry axis (yellow). The thin-shelled closed vessel is constrained on its right half by two nondeforming rigid, flat plates. B, The cylinder is meshed using three-dimensional quadrilateral shell elements (curved shell). The image on the left shows the first-order elements defined by four nodes (purple) used to discretize the geometry. Additional nodes (blue) would formulate the second-order elements. The elements can be regularly shaped or skewed. Excessively skewed element shapes are to be avoided. C, Boundary conditions are applied to the model. The rigid plates are fixed to prevent their rotation or displacement. Displacement boundary conditions are applied to prevent the cylinder moving freely in the space. The turgor pressure is applied uniformly to the internal surfaces. A sliding frictionless contact property is defined between the rigid plates and the deformable cylinder to prevent the penetration of one body into the other, while allowing their relative displacement. D, The isotropic closed cylinder deforms toward a spherical shape where it is not constricted by the plates. The heat map represents the von Mises stress distribution. E, First-order (left) and second-order (right) elements used around a discontinuity. F, Graph depicting the results obtained in a mesh convergence study. A value such as stress in a critical region is plotted against the total number of elements representing the structure to verify the independence of results from the mesh quality and the number of elements. Figure 1. Open in new tabDownload slide A, A closed cylindrical shell with hemispherical caps generated by the rotation of a line (orange) around a symmetry axis (yellow). The thin-shelled closed vessel is constrained on its right half by two nondeforming rigid, flat plates. B, The cylinder is meshed using three-dimensional quadrilateral shell elements (curved shell). The image on the left shows the first-order elements defined by four nodes (purple) used to discretize the geometry. Additional nodes (blue) would formulate the second-order elements. The elements can be regularly shaped or skewed. Excessively skewed element shapes are to be avoided. C, Boundary conditions are applied to the model. The rigid plates are fixed to prevent their rotation or displacement. Displacement boundary conditions are applied to prevent the cylinder moving freely in the space. The turgor pressure is applied uniformly to the internal surfaces. A sliding frictionless contact property is defined between the rigid plates and the deformable cylinder to prevent the penetration of one body into the other, while allowing their relative displacement. D, The isotropic closed cylinder deforms toward a spherical shape where it is not constricted by the plates. The heat map represents the von Mises stress distribution. E, First-order (left) and second-order (right) elements used around a discontinuity. F, Graph depicting the results obtained in a mesh convergence study. A value such as stress in a critical region is plotted against the total number of elements representing the structure to verify the independence of results from the mesh quality and the number of elements. IRREVERSIBLE SHAPE FORMATION IN GROWING PLANT CELLS Plant cell growth involves an irreversible stretching of the cell wall and an increase in cell volume and surface that can be substantial in certain cell types. Biologically, this is accompanied by the continuous insertion of new cell wall material to the existing wall. Simulating the resulting large deformations and the concomitant addition of material is a challenge for mechanical modeling that can be tackled in several ways. Typically, when using FE modeling, small, pressure-induced deformations are simulated repeatedly, and between the increments, a remeshing is performed. The geometrical structure resulting from the previous deformation step is meshed again to replace the often distorted mesh from the previous step. Stretching of the cell wall due to loading results in a reduction of cell wall thickness. To account for the addition of new material, the thickness of the wall is readjusted (e.g. set to the initial value to maintain a constant wall thickness). This principle has been used, for example, to simulate tip growth in the pollen tube, the delivery organ for the sperm cells in plants (Fayant et al., 2010). FE Modeling of Tip Growth Tip-growing cells such as pollen tubes, root hairs, and fungal hyphae feature a spatially confined expansion zone allowing these cells to perform invasive behavior (Sanati Nezhad et al., 2013; Sanati Nezhad and Geitmann, 2015; Bascom et al., 2018). The profile of the growing tip is radially symmetrical and remains self-similar when moving forward. Mutations or pharmacological treatments interfering with cytoskeletal functioning or cell wall composition can affect the self-similarity, resulting in either a tapered or a swollen phenotype (Kost et al., 1999; Hwang et al., 2005; Klahre et al., 2006; Kost, 2008). Modeling of tip growth has been addressed using a variety of approaches (Goriely and Tabor, 2003; Dumais et al., 2006; Kroeger et al., 2008; Campàs and Mahadevan, 2009; Kroeger and Geitmann, 2011, 2012), one of which focused on the cell wall mechanical properties using the FE approach. FE modeling was used specifically to simulate how the generation of aberrant pollen tube phenotypes is mediated by changes in the cell wall mechanics (Fayant et al. (2010). The wall of the growing pollen tube was represented with a shell of uniform thickness (Box 2) but spatially varying material properties. The apical dome was divided into hoop-shaped subregions in which the elastic modulus (Box 3; Boudaoud, 2010; Dumais, 2013; Niklas, 1992; Huang et al., 2012; Julkunen et al., 2007; Kha et al., 2010; Sun, 2006; Zerzour et al., 2009) could be assigned independently (Fig. 2A). Load application was performed repeatedly, and after each step, the structure was remeshed, and the wall thickness was reset to the initial value to counter thinning and simulate the addition of cell wall material (Fig. 2B). The goal was to predict the spatial distribution of material properties in the cell wall that would generate perfectly cylindrical and self-similar growth patterns and to identify those that result in deviations such as swelling and tapering. Open in new tabDownload slide Open in new tabDownload slide Open in new tabDownload slide Open in new tabDownload slide Figure 2. Open in new tabDownload slide A, A pollen tube modeled as a hollow shell with uniform thickness. The apical dome is divided into subregions, allowing for the elastic properties to be adjusted in each region independently. B, Several key points are followed on the FE model upon each loading cycle and remeshing to mimic growth. C, The stiffness gradient predicted by the FE model to produce a self-similar tube closely matches the deesterification pattern of pectin. Images were adopted from Fayant et al. (2010). Figure 2. Open in new tabDownload slide A, A pollen tube modeled as a hollow shell with uniform thickness. The apical dome is divided into subregions, allowing for the elastic properties to be adjusted in each region independently. B, Several key points are followed on the FE model upon each loading cycle and remeshing to mimic growth. C, The stiffness gradient predicted by the FE model to produce a self-similar tube closely matches the deesterification pattern of pectin. Images were adopted from Fayant et al. (2010). Two types of biological constraints were used to validate the biological relevance of the simulations. To represent a normally growing tube, the model had to produce a self-similar shape profile, and the strain pattern resulting from the deformation of the wall had to reproduce those observed experimentally (Rojas et al., 2011). Two key mechanical parameters were analyzed for their ability to shape the tube: (1) the profile of the elastic modulus gradient from tip to shank; and (2) the degree of material anisotropy expressing how different the material responds to loads applied in different directions. The validation suggested that isotropic behavior combined with a characteristic increase in elastic modulus, gradual from the tip to the flank and sudden from the flank to the shank, most accurately reproduce the pollen tube growth pattern. The mechanical gradients incorporated in this model were implemented as varying the elastic modulus between discrete regions rather than as a continuous and smooth function, and the values extracted were rather approximate. Despite these approximations, the model results matched well with mechanical and biochemical observations, suggesting that the simplification was not detrimental to the purpose of the study. Crucially, the gradient in elastic modulus predicted by the FE model correlates well with the biochemical composition of the pollen tube wall, notably the distribution of esterified and acidic pectin (Fig. 2C). This result is consistent with the effect of the pectinase-mediated digestion of pectin, which results in a dramatic apical swelling of the pollen tube (Parre and Geitmann, 2005), presumably through the loss of the modulus gradient. It also accords well with mechanical measurements revealing that the cell wall at the tip of the growing pollen tube is softer and modulates its properties to generate an oscillatory growth pattern (Zerzour et al., 2009). The reciprocation between predictions made by the in silico FE model, experimental validation, refinement of the model, and guidance toward further biological experimentation illustrates the value of FE in the predictive modeling of cell development. FE Modeling of Diffuse Growth during Trichome Branch Morphogenesis A similar yet distinct modeling approach was carried out to investigate the growth mechanics in trichomes. These epidermal cells (Fig. 3A) come in many shapes, sizes, and metabolic functions. They can be branched or unbranched, glandular or nonglandular (Tissier, 2012), and can be single-cell entities or comprise multiple cells. Trichome shape is intimately linked to their respective function, such as defense, pollination, or moisture retention (Oelschlägel et al., 2009; Amada et al., 2017). The unicellular trichome in Arabidopsis (Arabidopsis thaliana) forms a stellate body with three or four branches and is an excellent cell type in which to investigate the mechanics underlying complex cell morphogenesis. Actin is involved in the diffuse growth of plant cells (see Szymanski and Staiger, 2018). Disruption of cytoskeletal components is associated with the loss of branching, a needle-like phenotype, or swelling (Mathur et al., 1999; Szymanski et al., 1999; Mathur, 2004), phenotypes that can be studied using FE modeling. Branch morphogenesis in Arabidopsis trichomes was investigated using FE modeling in conjunction with live-cell imaging to understand the mechanics of branch growth (Yanagisawa et al., 2015). Similar to modeling of the pollen tube, the two constraints on the model to match were the shape of the branch and the growth (strain) pattern of the wall. Time-lapse imaging demonstrated that, unlike the self-similar pollen tube, the trichome tip radius tapers off while the radius of the base of the branch remains relatively constant (Fig. 3B). Fiducial markers were used to track the local growth pattern. To justify a choice of material model, the authors visualized the alignment of cellulose synthase (CESA) complexes and microtubules (see Elliott and Shaw, 2018), which were oriented transversely to the branch axis. Motivated by this preferential orientation inferred for cellulose that is commonly regarded as the major load-bearing polymer of the cell wall, transverse isotropy (Box 3) was incorporated in the elastic model using the Holzapfel-Gasser-Ogden hyperelastic material behavior (Gasser et al., 2006). The critical parameters in this material model are the dispersion, fiber angle, and fiber-to-matrix stiffness ratio, which needs to be sufficiently large for anisotropy to emerge (Huang et al., 2012). The FE simulations indicated that strong transverse alignment of fibers results in the anisotropic, axial growth of the shell. The random orientation of the fibers implemented in this model produced a spherical bulge instead. Figure 3. Open in new tabDownload slide A, Epidermal cells on the adaxial surface of an Arabidopsis leaf feature three cell types: trichomes (brown), stomatal guard cells (red), and pavement cells (green). B, Development of the trichome branch embodies reduction of the tip radius of curvature, while the radius at the base of the branch remains constant. L, branch length; RT, branch radius at the tip; RB, branch radius at the base. C, The growth and thickness of the cell wall in a trichome branch are correlated and exhibit a gradient toward the tip of the branch. Microtubules and CESA trajectories are oriented transversely to the long axis of the branch, while the tip exhibits a microtubule-depleted zone. Image redrawn after Yanagisawa et al. (2015). Figure 3. Open in new tabDownload slide A, Epidermal cells on the adaxial surface of an Arabidopsis leaf feature three cell types: trichomes (brown), stomatal guard cells (red), and pavement cells (green). B, Development of the trichome branch embodies reduction of the tip radius of curvature, while the radius at the base of the branch remains constant. L, branch length; RT, branch radius at the tip; RB, branch radius at the base. C, The growth and thickness of the cell wall in a trichome branch are correlated and exhibit a gradient toward the tip of the branch. Microtubules and CESA trajectories are oriented transversely to the long axis of the branch, while the tip exhibits a microtubule-depleted zone. Image redrawn after Yanagisawa et al. (2015). As the tip apex of the trichomes was observed to be depleted of microtubules, it was concluded that the cell wall at the tip should be isotropic. Varying the size of the isotropic zone and comparing the evolution of the tip radius of curvature against the branch length, the FE results indicated that the size of the isotropic apical zone should vary over time to reproduce the experimentally observed tapering. This conclusion was drawn because no single value of the tip isotropic zone could produce results that fit the experimental curve. It is not clear to what extent such detailed results depend on the choice of material model, and whether a different hyperelastic model would bear a different conclusion remains to be investigated. However, the transverse isotropy per se could not reproduce the growth gradient toward the tip observed experimentally (Fig. 3C). Therefore, it was suggested that a thickness or elastic modulus gradient should exist along the trichome branch. Transmission electron microscopy and light microscopy confirmed this predicted attenuation of cell wall thickness toward the tip, with a value close to but less dramatic than that predicted by the FE model. Therefore, a combination of both thickness and elastic modulus gradient parameters might be employed by the cell, although the FE model used here produced unrealistic results when combining the two. As remarked earlier, it would be interesting to investigate the dependency of simulation results on the material model employed. In the Arabidopsis mutant arpc2/distorted2, trichome branch growth is hampered, the tip radius of curvature remains high compared with the wild type, and the stalk swells (Kotchoni et al., 2009). Intriguingly, wall thickness and growth gradients were both absent in these aberrant trichome branches, further corroborating a correlation between wall thickness and growth rate variations. However, whether the relationship is causal, and if it is, which is the cause or effect, remain unclear. While a thinner wall in the model can translate into a lower rigidity to reproduce a higher strain, a higher strain in the absence of reinforcing new wall material results in wall thinning. Likewise, the absence of a thickness gradient in arpc2 may result from a failure to grow and a consequent lack of wall thinning. This study suggests that, while similarities exist, the growth behavior particular to trichome branches is distinct from tip growth, as the tip radius attenuates, wall thickness is not preserved, and growth occurs in the whole branch rather than a confined apical zone. Yanagisawa et al. (2015) report that the cell wall of arpc2 is enriched in well-aligned cellulose, and microtubules are transversely oriented. Cellulose orientation was used as a proxy for transverse isotropy bringing about anisotropic (axial) growth in mutant branches. Although trichome branches in arpc2 do not grow considerably, comparing the time data provided for the branch length and tip radius for the wild type and mutant shows that, for an equal length (e.g. at 40-µm branch length), the mutant branch has a larger tip radius of curvature, indicating that at least some degree of swelling occurs in the branch too, besides the general swelling in the stalk (see Figs. 1B and 3C in Yanagisawa et al., 2015). Whether this apparent swelling is a result of changes in the tip isotropic zone or changes in other wall polymers is not clear. While the fibrillar texture of the cell walls between the wild type and the mutant seems unaltered, the quality of the fibers and their linkages might have undergone changes. Furthermore, other wall polymers may have been affected as a result of mutation. This emphasizes the need for experimental studies to characterize and compare the changes in mechanical properties in a wide array of Arabidopsis mutants; the mechanical changes due to mutations can be manifested in ways that go beyond what can be assessed readily by visually tracing cellulose orientation. In the studies that have modeled the shaping process of plant cells using the FE method, the geometry is loaded incrementally while cell wall thickness is adjusted to account for the addition of material. The drawback of this strategy is that this process results in the elimination of stresses that develop in each increment. While it is possible to retain the stress information and transfer it to the next increment, it might differ from the quality of wall stresses during the insertion of wall-building materials. Attempts to describe cell growth were also been done by using viscoelastic behavior (Huang et al., 2015). However, loading to deform an elastic or viscoelastic structure develops stresses that cleavage of chemical bonds and insertion of new materials might not. In fact, currently, we have little information on changes occurring in cell wall stress during growth or cell deformation. Additionally, in most of the available modeling studies, stress information is presented only in relative form. Absolute stress values could serve as a useful parameter to further validate the quality and relevance of a model’s predictions. However, reporting absolute values based on models that are inevitably greatly simplified requires considerable experimental support, as explained below. Stress Development in Plant Cells Correlates with Morphogenetic Phenomena A subset of forward FE simulations of plant cells are the stress analysis models. The main parameter investigated in these models is generally the stress developed in a single step of turgor application. The stress analysis models of plant cells or tissues published so far do not fall under irreversible FE models and are static, since they do not involve remeshing, wall modification, stress update, or otherwise introduction of a form of permanent deformation. While these models do not explicitly simulate irreversible material deformation, since they are sometimes employed to investigate the link between the mechanics and a physiological or morphogenetic response, such as cytoskeletal patterning, that can be linked to an irreversible biological response, we categorize them under the irreversible use of FE models. Several studies use static stress analysis to correlate the mechanical aspects with a morphogenetic problem such as gene expression, hormonal activity, and ontogeny at tissue scale (Bassel et al., 2014; Bozorg et al., 2014; Boudon et al., 2015). To illustrate the concept, however, here we will discuss those that focus primarily on cell shape, namely the shape of epidermal pavement cells in relation to microtubule organization and wall biochemistry. Pavement cells in the leaf epidermis of eudicotyledons form interlocking patterns similar to pieces of a jigsaw puzzle (Fig. 3A). There have been numerous hypotheses, such as cuticle stiffening or cells being squeezed physically during growth, to explain the peculiar shaping phenomenon in these cells (Korn, 1976; Armour et al., 2015). The potential role of mechanics underlying the shaping process and the potential advantages of such cell shape for the epidermis or leaf have remained elusive (Jacques et al., 2014). Since pavement cells, as opposed to trichome branches, are tightly connected to neighboring cells, they allow studying the mechanics of cell-tissue interaction. Studies have suggested the involvement of the cytoskeleton in the shaping process downstream of an auxin-dependent pathway (Fu et al., 2005; Xu et al., 2010; Zhang et al., 2011; Lin et al., 2013). Microtubules are suggested to be associated with regions of indentation (often termed the neck), putatively resulting in anisotropic reinforcement of these regions by guiding CESA complexes and preferential deposition of cellulose microfibrils (Panteris and Galatis, 2005; Belteton et al., 2018). For simpler cell shapes, it is known that microtubules reorient in the direction of maximal mechanical stress (Williamson, 1990; Hamant et al., 2008). Whether the microtubule arrays experimentally observed in pavement cells equally correlate with stress patterns, however, was unknown for lack of information on stress distribution. FE Modeling of Turgor-Induced Stresses in the Periclinal Pavement Cell Walls The correlation between cell shape, mechanical stress, and microtubule alignment can be easily verified for simpler cell geometries, such as the tubular shapes of trichome branches or root and shoot epidermal cells. In a pressurized thin-walled cylindrical vessel with hemispherical caps, the pressure-induced transverse stress that arises due to cell shape is twice the longitudinal component. However, in the case of pavement cells, predicting the local distribution of stress is not as straightforward. Sampathkumar et al. (2014) used FE modeling to study the effect of cell shape and tissue-level stresses on microtubule arrangement in pavement cells. The static model developed in this study focuses on cell-level stress development and how it correlates with tissue-level mechanical and physiological responses. The FE model consists of the outer periclinal walls (the horizontal walls parallel to the plane of leaf) of pavement cells, extracted from confocal microscopy stacks, modeled as thin shells. At the borders of periclinal walls, the anticlinal walls (the vertical side walls) were modeled as 1D beam elements adding stiffness to borders. Tensile stress was generated in the cell walls upon the application of pressure to the inner face of the walls. Whether an isotropic or transversely isotropic hyperelastic material was used, the result indicated a higher stress at the indentation side of the wavy borders. The location and pattern of stress from the shell FE model matched well with the anisotropic alignment of microtubules, with a preference for bundling at sites of indentation (Fu et al., 2005; Zhang et al., 2011; Armour et al., 2015). These data suggest that stress resulting from cell shape at the subcellular scale might act as a mechanical cue for the cytoskeletal arrangement, even in cells with complex shapes. Sampathkumar et al. (2014) used the FE model of the pavement cells further, to predict the microtubule organization based on tissue-level stress patterns and their alterations upon the application of external forces or cell ablation. The FE model suggested that, upon laceration, or more subtly, cell ablation, the stress pattern becomes circumferential around the vacant region (Fig. 4, A and C). Cell removal was reproduced in the model by gradual reduction in turgor pressure and cell wall stiffness in compromised cells. Time-lapse imaging reported an increase in microtubule bundling and a change in organization hours after laceration. Previous observations also had reported a reorientation of microtubules due to externally applied mechanical stress in Arabidopsis leaves (Jacques et al., 2013). The authors propose that this indicates that changes in tissue-wide mechanical stresses can affect microtubule organization despite the initial cell-level order imposed on microtubular arrangement. However, to what degree the perturbation of mechanical stresses at tissue scale can override the cell-level control of microtubule arrangement is disputable. First, the FE model developed by Sampathkumar et al. (2014) predicts that, upon laceration or cell removal, circumferential stress patterns would be produced in pavement cells in a region of tissue spanning over multiple cells (Fig. 4A). Conversely, observation of a fluorescently tagged microtubule line seems to suggest that microtubule arrangement is affected more strongly only in cells near the afflicted region (Fig. 4B). Second, the study reports that the response of microtubules to changes in mechanical stress depends upon the magnitude of stress. To demonstrate this, the FE model was used to reproduce the ablation of only a few pavement cells. Similar to the case of laceration but with a lower magnitude, the FE model predicts a circumferential rearrangement of microtubules adjacent to the location of the perished cells (Fig. 4C). However, the experimental observation does not seem to closely match the FE prediction (Fig. 4, C and D). The authors argue that this observation demonstrates a stress-magnitude dependency of the microtubule rearrangement and that ablation of only a few cells does not seem to be able to strongly rearrange the microtubules circumferentially in neighboring cells. However, a closer look reveals microtubules in neighboring cells to be oriented predominantly parallel to the long axes of the cells, rather than featuring a circumferential orientation around the site of ablation (Fig. 4D). This might indicate that the rearrangement of microtubules due to cell ablation might at least partially occur due to other cues, such as a wound response, a mechanism shown previously (Geitmann et al., 1997). Furthermore, the microtubule response might depend upon a competition between the subcellular and supercellular cues, with a weight factor for each that depends on the homeostasis within the cell and tissue, rather than being dominated consistently by one. Figure 4. Open in new tabDownload slide A, FE model suggesting a stress pattern in pavement cells circumferential to the site of laceration. B, Fluorescence-tagged microtubules demonstrate hyperbundling and a seemingly circumferential pattern around the wound site. White arrowheads show examples of noncircumferential microtubules in cells away from the site of laceration. The white arrow shows that, even in cells adjacent to laceration, there seems to be a local competition between the cell shape-dictated microtubule organization and the putative circumferential reorientation of microtubules due to tissue-level stress. C, FE model suggesting a stress pattern in pavement cells circumferential to removed cells. D, Microtubules demonstrate a change in bundling and orientation upon the small-scale wound. However, their orientations seem longitudinal to cell axes rather than being circumferential to the site of the wound. White arrows indicate examples of cells with microtubules oriented parallel to the long cell axis, inconsistent with the hypothesized circumferential orientation. The star indicates ablated cells. Numbers indicated individual cells. All these images are reprinted from Sampathkumar et al. (2014) with permission from the authors. Bars = 25 µm (B) and 50 µm (D). Figure 4. Open in new tabDownload slide A, FE model suggesting a stress pattern in pavement cells circumferential to the site of laceration. B, Fluorescence-tagged microtubules demonstrate hyperbundling and a seemingly circumferential pattern around the wound site. White arrowheads show examples of noncircumferential microtubules in cells away from the site of laceration. The white arrow shows that, even in cells adjacent to laceration, there seems to be a local competition between the cell shape-dictated microtubule organization and the putative circumferential reorientation of microtubules due to tissue-level stress. C, FE model suggesting a stress pattern in pavement cells circumferential to removed cells. D, Microtubules demonstrate a change in bundling and orientation upon the small-scale wound. However, their orientations seem longitudinal to cell axes rather than being circumferential to the site of the wound. White arrows indicate examples of cells with microtubules oriented parallel to the long cell axis, inconsistent with the hypothesized circumferential orientation. The star indicates ablated cells. Numbers indicated individual cells. All these images are reprinted from Sampathkumar et al. (2014) with permission from the authors. Bars = 25 µm (B) and 50 µm (D). FE Modeling of Wave Formation in Anticlinal Walls of Pavement Cells While the model by Sampathkumar et al. (2014) was useful to predict how stresses in fully differentiated pavement cells are distributed, it does not address the generation of the cell undulations. Majda et al. (2017) proposed an FE model to explain the underlying mechanics of wave formation in pavement cells. Motivated by their undulating shapes, the model focuses exclusively on the anticlinal cell walls. The authors propose that wave formation results from bending of the anticlinal walls due to their stretching combined with a particular spatial distribution of mechanical properties. The FE model developed in this study purportedly demonstrates that, if segments with high and low elastic moduli are laid alternatingly along and across the anticlinal walls, stretching this structure forms bends resembling the protrusions and indentations of the pavement cell wall. A main result of the model is that the indentation side of the bend of the anticlinal wall is associated with the softer material. While this result seems corroborated by the atomic force microscopy (AFM) stiffness measurements reported in this study, it is challenged by available data on the mechanics of the pavement cell wall. First, as mentioned before, well-aligned microtubules are associated with indentation sides in anticlinal and periclinal walls (Fu et al., 2005; Zhang et al., 2011; Armour et al., 2015), although whether these antecede the initiation of undulations warrants further investigation (Belteton et al., 2018). It was hypothesized that this microtubular array leads to the deposition of well-aligned cellulose enrichment, thus preventing further expansion in these areas of the periclinal walls (Panteris and Galatis, 2005). This local stiffening in the periclinal wall of differentiated cells is corroborated by AFM stiffness mapping (Sampathkumar et al., 2014). Moreover, using fiducial markers on the surface of growing pavement cells, Armour et al. (2015) demonstrated that cell wall expansion is more pronounced on the protrusion side of undulations while the opposing indentation sides seemed restricted in their growth. This further corroborates an added stiffness at the indentation side, all of which seems to be difficult to reconcile with the results of the anticlinal wall FE model developed by Majda et al. (2017). While the two scenarios are not mutually exclusive, reconciling them would necessitate allowing for a drastic and sudden change in the biochemical and biomechanical makeup at the border between adjacent cell wall regions. An indentation would have to feature a stiff periclinal wall (Panteris and Galatis, 2005; Sampathkumar et al., 2014; Armour et al., 2015) directly neighboring a soft anticlinal wall (Majda et al., 2017). Only detailed analysis of the local wall biochemistry and mechanical behavior will provide conclusive answers. An additional consideration is the geometry. Given that the very narrow band of the anticlinal wall is bordered by two large sheets of periclinal wall, potentially representing significant boundary conditions limiting the freedom of displacement, one wonders whether the former can dominate the latter and whether modeling the anticlinal wall while entirely neglecting the periclinal walls can be a justifiable simplification. We posit that an FE model that represents the entire 3D geometry of the cell, including all its load-bearing walls, is warranted to address the challenge of wavy pavement cell morphogenesis. REVERSIBLE SHAPE CHANGES IN PLANT CELLS Reversible shape changes in plant organs and cells can be generated by modulation of the turgor pressure. As opposed to growth-related deformations, these remain mostly in the elastic range and do not involve dynamic modification of the cell wall material and biochemistry. However, these reversible movements are still governed by the mechanical properties of the cell wall material and the geometry of the cell or tissue. At the tissue level, the opening of the Venus flytrap (Dionaea muscipula) and processes enabling seed dispersal are examples of the exploitation of turgor modulation, cell shape, and wall mechanics to accomplish actuation (Forterre et al., 2005; Geitmann, 2016; Hofhuis et al., 2016). Similarly, motion at the single cell level, such as stomatal opening and closing, is powered hydraulically. To illustrate the application of FE modeling to reversible shape changes, the example of stomatal guard cells is examined here. Guard cells form pores in the leaf epidermis that are specialized to optimize gas exchange between the plant and the environment. The ability of these epidermal valves to respond efficiently to various stimuli, such as light and aridity, is crucial for photosynthesis, water retention, and, thus, survival. Stomatal opening is driven by an increase in turgor pressure in the guard cells. The pressure in subsidiary cells, the specialized epidermal cells immediately surrounding the guard cells, antagonizes this process, together regulating the stomatal dynamics (Von Mohl, 1856; Edwards et al., 1976; Franks and Farquhar, 1998). The shape and structure of guard cells vary among species. Stomata in graminaceous monocots have guard cells that are typically narrow and dumbbell shaped, whereas those of dicots are kidney shaped. Here, we focus on the latter. The increase in the width of the stomatal aperture by inflation of the guard cells is suggested to occur in two stages. The guard cells in the closed state of the pore display a nearly elliptical cross-section that becomes circular when pressurized (Fig. 5A). The first stage of the pore opening or increase in volume of guard cells is suggested to be governed mainly by an inflation-driven change in the cross-sectional shape of guard cells with little stretch in the wall. The further increase in cell volume is attributed to stretching of the cell wall accompanied by its thinning and expansion at the poles (Sharpe and Wu, 1978). How the swelling of the two guard cells, their specific design, and their wall mechanics enable pore opening has been the subject of multiple studies with somewhat antithetical outcomes (DeMichele and Sharpe, 1973; Raschke, 1975; Cooke et al., 1976, 2008; Sharpe and Wu, 1978; Amsbury et al., 2016; Carter et al., 2017; Woolfenden et al., 2017). Several cell features have been hypothesized to be critical for pore opening when guard cell pressure rises: (1) the increased thickness of the ventral walls of guard cells (Fig. 5A); (2) the radial reinforcement by cellulose microfibrils, resulting in anisotropy of the cell wall (Fig. 5B); (3) the elliptical cross-section of guard cells under low turgor pressure; and (4) the constraint on polar expansion of guard cells due to pectin deesterification. Figure 5. Open in new tabDownload slide A, Cross-sectional view of guard cells composed of ventral wall (VW), dorsal wall (DW), inner wall (IW), and outer wall (OW). Ra and Rb refer to the horizontal and vertical radii of the elliptical cross-section, respectively. Only the outer ledge (OL) is shown. Inflation of the guard cells causes a change of the elliptical cross-section to circular and then to an ellipse with the major axis perpendicular to the plane of the leaf. B, Confocal micrograph of guard cells in an Arabidopsis cotyledon, stained with Calcofluor White to reveal cellulose. Bar = 15 µm. Figure 5. Open in new tabDownload slide A, Cross-sectional view of guard cells composed of ventral wall (VW), dorsal wall (DW), inner wall (IW), and outer wall (OW). Ra and Rb refer to the horizontal and vertical radii of the elliptical cross-section, respectively. Only the outer ledge (OL) is shown. Inflation of the guard cells causes a change of the elliptical cross-section to circular and then to an ellipse with the major axis perpendicular to the plane of the leaf. B, Confocal micrograph of guard cells in an Arabidopsis cotyledon, stained with Calcofluor White to reveal cellulose. Bar = 15 µm. Reversible Changes in Guard Cell Cross-Sectional Shape May Underlie Stomatal Pore Opening FE studies by Cook and colleagues were among the first to consider a realistic closed-cell geometry for guard cells. The comprehensive analyses carried out in these studies produced results that remain cogent to date (Cooke et al., 1976, 2008; Lee, 1986; more information is available under Stomatal Control System [hdl.handle.net/1813/45423]). Cooke et al. (1976) modeled a generic stomate with a doubly elliptical toroidal shell by rotating an ellipse forming the transverse cross-section about another ellipse that lies in the horizontal plane (Fig. 5A; videos demonstrating the data for the guard cell model and results, for shell and solid, can be accessed at the following links: hdl.handle.net/1813/43793 and hdl.handle.net/1813/43794). The parameters investigated were the effects of guard cell geometry, wall thickness, radial cellulose reinforcement, and the turgor pressure in both guard and subsidiary cells. Thickening of the ventral wall has long been proposed to underlie pore opening (Meidner and Mansfield, 1968). Considering a nonuniform wall thickness, Cooke et al. (1976) found that opening of aperture width is virtually the same as in a model with uniform wall thickness, suggesting the wall thickness gradient to be insubstantial for stomatal opening. These results indicated, however, that the geometry of guard cells is paramount in their function. If the torus was set to be doubly circular in this model or if the cross-section was defined to be elliptical with the major axis perpendicular to the leaf (dotted ellipse in Fig. 5A), the pore was shown to close upon pressure application unless some physiologically implausible criteria were enforced. In these cases, the guard cell deformation majorly consisted of cell wall stretching rather than change in cell cross-sectional shape. In contrast to other geometries, the authors determined that a doubly elliptical geometry opens upon the application of turgor pressure and disturbs the neighboring cells minimally when inflated. The simulations indicated that, during deformation and pore opening, the cells bulge out of the leaf plane. While the width of the pair increases as a result of pore opening, the width of a guard cell can decrease slightly because of the out-of-plane bulge of the inner and outer walls. Interestingly, the aperture length was observed to remain virtually constant during pore opening, without necessitating a displacement restraint to be imposed a priori on guard cell poles. Parametric studies carried out by Cooke et al. (1976) revealed that the aperture width is a multilinear function of the pressures in guard cells and subsidiary cells. An antagonism ratio was defined to express the contribution of each cell type in aperture opening. The radial orientation of cellulose microfibrils in mature guard cells (Fig. 4B) has long been known (Ziegenspeck, 1955), and the resulting transverse isotropy has been considered a crucial feature promoting stomatal opening. An early study even made a physical model of stomatal opening by radially reinforcing a pair of elongated balloons using adhesive tape (Aylor et al., 1973). In contrast, simulations by Cooke et al. (1976) suggest that circumferential cellulose reinforcement acts as a hindrance to aperture opening driven by guard cell inflation. Ceteris paribus, their model, predicted that increasing the elastic modulus of the wall in the radial direction, representing a higher anisotropy ratio by radial cellulose bundles, diminishes the effect of a unit increase in guard cell pressure on pore opening while it increases the contribution of a unit increase in the pressure of subsidiary cells in closing it. From this, the authors concluded that radial cellulose anisotropy is not a mechanism to open but, conversely, a leverage to close the pore. Interestingly, a recent study by Rui and Anderson (2016) has demonstrated that guard cells in a mutant with reduced cellulose content and anisotropy exhibit a wider aperture, which seems to, at least partly, support the findings of the model by Cooke et al. (1976), although assessing the dynamics of the pore opening and the effect of subsidiary cells will require further investigation. The emergence of such complex and nonlinear control on the Watergate by relying only on geometry and mechanics is a spectacularly simple strategy and may be, in part, how plants can respond rapidly and reliably to environmental cues (Roelfsema and Hedrich, 2005; Franks and Farquhar, 2007; Raven, 2014). Together, Cooke et al. (1976, 2008) and Lee (1986) conclude that opening of the stomatal pore is influenced saliently by the elliptical geometry of the guard cell cross-section. They suggest that nonuniform wall thickening or anisotropic material properties are not required per se, although they might regulate the dynamic response of guard cells. Reassessing the Contribution of Cellulose-Induced Radial Anisotropy to Stomatal Opening The role of cellulose orientation and cell wall anisotropy was reassessed in a recent study by Woolfenden et al. (2017), who used nonlinear elasticity with a transversely isotropic material behavior to represent the guard cell wall, similar to studies by Cooke et al. (1976). The authors observed that, using isotropic material properties, increasing the turgor pressure causes the stomatal pore to close rather than open. A radially reinforced version of their model engendered pore opening. They concluded that circumferentially oriented cellulose microfibrils are crucial for stomatal opening. However, these results were based on a structure with an idealized circular cross-section for the guard cells, whereas the base model in the study by Cooke et al. (1976) was elliptical in cross-section (Fig. 5A). While Woolfenden et al. (2017) did simulate this situation as well, they stated that, at higher ‘correct’ pressures, the pore closes in the absence of anisotropic properties. The study does not explicitly state a caveat that cannot be neglected, however. Many of the model inputs, including the cell wall thickness, cross-sectional shape, and material model parameters including the elastic moduli, are significantly simplified, and idealized or arbitrary values are used, as is inevitable in the absence of biomechanical data. Therefore, it is hardly possible to expect the model to reliably predict anything more than general tendencies, even if biologically relevant absolute pressure values are used. A small change in, say, the assumed Young’s modulus of the cell wall or the use of a different nonlinear elastic model has the potential to significantly alter the observed threshold values or even the trends at which the model switches from open to closed. Arguing that one model more accurately reflects the reality over another, if neither uses better quality input parameters, warrants substantiation. Similar caveats apply to attempts aimed at quantitatively identifying material constants from such models; many approaches are best suited to remain qualitative. This points to a limitation of modeling in plant cell mechanics in general. In the absence of detailed quantitative information, many parameters required to define the model must be input based on educated guesses, and whether the predictions made by the model hold in experimental conditions remains to be shown. That a given combination of material and geometrical parameters produces results resembling the biological situation is seducing but does not prove this combination to be the one reflecting the reality. Other solutions often are possible. This caveat should at least be acknowledged but is not always done. Another example is the observation that the aperture is least sensitive to turgor pressure at higher width (the curve moves toward a plateau; Franks and Farquhar, 1998). This led Woolfenden et al. (2017) to conclude that a strain stiffening must occur in the cell wall matrix. While this explanation is reasonable, it may not be the only one. Annexing increasing numbers of parameters to the model to match an observed behavior, without proper substantiation, risks appearing opportunistic if not arbitrary. The nonlinear behavior observed in experiments (Franks and Farquhar, 1998) also could be caused by a two-step opening mechanism, similar to the one described before and demonstrated in the FE model by Cooke et al. (1976, 2008). This would entail an initial turgor-induced change in geometry from elliptical to circular (inflation), followed by an accommodation of any further increase in pressure by cell wall stretching, a process that requires higher turgor differentials to produce visible results. Therefore, the matrix strain-stiffening conjecture proffered by Woolfenden et al. (2017), while consistent with polymer chemistry (Bidhendi and Geitmann, 2016), could be a secondary mechanism responsible for the apparent low sensitivity of aperture opening at high pressures. These concepts certainly merit further validation. Woolfenden et al. (2017), furthermore, did not extensively explore the effect of subsidiary cells in their model. They suggested that, in the presence of cellulose-mediated anisotropy in the guard cell walls, the effect of pressure in the subsidiary cells becomes negligible. This result is in clear contrast to that of Cooke et al. (1976), who reported the guard cell anisotropy to augment the effect of subsidiary cells in closing the stomata. Further investigations must address this discrepancy. Correlating the Mechanics and Phenotype of Genotypes: The Devil May Lie in the Ultrastructural Details While cell wall mechanical studies often focus on cellulose orientation, parameters such as cell wall thickness, cross-section shape, and other cell wall polymers seem to have the potential to influence the mechanics and function of guard cells. However, since a multitude of parameters may interplay, isolating the role of each may not always be straightforward. It has been reported that pectin chemistry is correlated with the ability of guard cells to function correctly (Merced and Renzaglia, 2014; Amsbury et al., 2016). pme6-1 was shown to have a decreased dynamic range and a defect in stomatal opening. Interestingly, the results of immunolabeling suggest that, in wild-type Arabidopsis, unesterified pectin (antibody, LM19) is present in all cell wall regions of guard cells. Highly methylated pectin (antibody, LM20), on the other hand, is absent from guard cell walls, as also reported previously by Merced and Renzaglia (2014), and is limited to cell junctions, as is calcium-bridged pectin (antibody, 2F4). This means that unesterified pectin in the guard cell walls, although negatively charged, does not seem to be gelated by calcium ions. In pme6-1, the pectin distribution was reversed: highly esterified pectin was reported in the entire guard cell wall. The authors concluded that pectin determines the mechanics of the cell wall and that lack of a functioning pectin methylesterase causes the wall to become too rigid and lose the deformability required to open the pore (Amsbury et al., 2016). While this is a reasonable hypothesis (Bidhendi and Geitmann, 2016), the study provides no evidence for the fact that the weakly esterified pectin was gelated by calcium or whether the mechanical properties of the cell wall materials were altered in any way. The antibody 2F4 was not used on the mutant, let alone micromechanical testing of cell wall properties. More importantly, the study fails to ascertain that no other parameters are changed in the mutant. While the authors claim a lack of any noticeable difference in ultrastructure between the wild type and the mutant, transmission electron microscopy cross-section micrographs of the guard cells of pme6-1 and the wild type provided in their study seem to reveal an interesting, yet conflicting, phenomenon (see Supplemental Fig. S2, G and H, of Amsbury et al., 2016). In cross-sections, the guard cell walls of the mutant appear considerably thicker than those of the wild type, relative to the area of the lumen. This seems especially the case for the inner walls. Whether the images in this paper are representative remains open. However, compensation mechanisms are common when a normal process of the cell is disturbed and can cause a chain of events. Here, in response to a disturbed function of pectin methylesterase, an increase in cell rigidity may have occurred through abnormal wall thickening. If this were confirmed, the change in cell wall dimensions or cross-sectional shape, rather than biochemical processes, would be the factor altering the mechanics of the unit surface of the wall (see Supplemental Fig. S2, G and H, of Amsbury et al., 2016). The irregular cell shape as viewed in the cross-section and inner wall thickness may explain the anomaly in stomatal opening, which requires further investigation through FE modeling. It should be noted that an earlier study on the chemical alteration of the guard cell wall during development in Funaria hygrometrica by Merced and Renzaglia (2014) reserves a special role for rhamnogalacturonan I distribution in guard cell walls, in addition to the role of homogalacturonan. Further assessment of the potential contribution of other types of pectin in guard cell mechanics may provide answers to some of the outstanding questions. The Roles of Pectin-Induced Stiffening and Adjacent Subsidiary Cells in the Polar Prevention of Guard Cell Elongation and Stomatal Opening Pectin biochemistry was also the focus of a modeling study that suggests deesterification of pectin at poles of guard cells to underlie pore opening (Carter et al., 2017). Ventral walls of guard cells are generally thicker compared with dorsal walls (Meidner and Mansfield, 1968; Renzaglia et al., 2017). Using AFM stiffness measurements, Carter et al. (2017) suggested that ventral wall thickening does not occur in young guard cells, while the stomata are still as functional as in mature guard cells. This is consistent with predictions made by an FE model developed in the same article, similar to the model by Woolfenden et al. (2017), suggesting the effect of ventral wall thickening to be minimal in stomatal opening, as also proposed by Cooke et al. (1976). Interestingly, Carter et al. (2017) showed that pectin is unesterified in guard cell poles and that the application of polygalacturonases rendered guard cells incapable of opening the aperture. Probing the stiffness of the enzyme-treated guard cells with AFM, the authors suggested that the guard cells’ dysfunction arises from the removal of polar stiffening due to polygalacturonase treatment. However, from the image provided, it seems that, rather than the removal of stiffness from poles, the enzyme treatment had caused the relative apparent stiffness to spread over a broader region of the guard cell walls (see Fig. 4I of Carter et al., 2017). Furthermore, it should be considered that the enzyme treatment also might change the turgor pressure in the guard cells or subsidiary cells. Neither of these possible collateral effects nor any possible changes in cell ultrastructure resulting from pectin modification were verified. The simulations predicted that, whether or not the cells are fixed at the poles, a threshold pressure (1 MPa) is required to initiate aperture opening. A threshold pressure has indeed been observed in biological samples. This lag in response was found to be due to the antagonizing effect of turgid subsidiary cells (Franks and Farquhar, 1998). As the pressure in these cells approaches zero, for example due to damage, the threshold pressure vanishes and the pore opens at guard cell pressures close to zero. The role of subsidiary cells is not spelled out in the model by Carter et al. (2017), but the modeling equivalent of an external constraining obstacle was nevertheless incorporated, only motivated by an unrelated biological feature. Carter et al. (2017) used the experimental finding of the stiffened guard cell poles to hypothesize that polar stiffening augments the pore opening at a given turgor pressure. They argued that the stiffened polar cell wall fixes the cell ends in place, and this concept was implemented by adding a boundary condition to the Woolfenden et al. (2017) FE model, on which the model by Carter et al. (2017) is based. This boundary condition consisted of fixing the poles in place. The problem of this translation of a biological concept into the FE model is that, to replicate the polar stiffening due to pectin deesterification, it should have been implemented as a property (e.g. locally elevated Young’s modulus) of the guard cell wall. Instead, the constrained displacement boundary condition removes displacement degrees of freedom at the poles, which biologically can only reflect an external constraint such as the above-mentioned surrounding subsidiary cells. Therefore, while the simulations are consistent with the findings by Franks and Farquhar (1998), the biological justification used by Carter et al. (2017) to implement the boundary condition merits reassessing. It is important to note that, as mentioned before, the model developed by Cooke et al. (1976; videos are available under hdl.handle.net/1813/43793 and hdl.handle.net/1813/43794) did not demonstrate a considerable polar expansion even though the poles were free to displace. Clearly, the choice of the model geometry is a crucial step in model construction. Open in new tabDownload slide Open in new tabDownload slide Future studies to address these questions, specifically the effects of cell cross-sectional shape and subsidiary cells on the stomatal complex, have the potential to further elucidate the functioning of pectin in stomatal mechanics. Suffice it to say that the scenarios proposed by various groups, even if seemingly inconsistent, provide food for further thought. We wonder whether guard cells use different mechanisms redundantly, or distinctly at different stages of development. An observation reinforcing this hypothesis may be the change from semicircular to elliptical cross-section between early-stage and mature guard cells (Merced and Renzaglia, 2014). REMARK FE modeling is a powerful tool that has been employed successfully to simulate the behavior of geometrically complex plant cells. The modeling technique has been used to localize and predict stress and strain in cells with the purpose to understand the underlying biological mechanisms, and it has been applied to both reversible and irreversible processes, such as guard cell movement and cell growth events, respectively. Despite, or because, the rapid adoption of FE modeling by the plant cell community, care must be taken in interpreting FE results. FE modeling, as with any modeling strategy, is subject to a dependency on the quality of inputs; flawed inputs result in flawed outcomes. Oversimplification or misrepresentation of the model components, ranging from the geometry, material behavior, or boundary conditions, has the potential to bear misleading results or to reinforce a bias as a self-fulfilling prophecy (see Outstanding Questions). Good modeling practice is to experiment with and eliminate the parameters that may affect the outcome before accepting the remaining ones. It is the responsibility of the user to ascertain that the inputs agree well with the physics of the problem and that the output is biologically relevant, which requires a proper understanding of both the physics and the biology of the problem. LITERATURE CITED Amada G , Onoda Y, Ichie T, Kitayama K ( 2017 ) Influence of leaf trichomes on boundary layer conductance and gas‐exchange characteristics in Metrosideros polymorpha (Myrtaceae) . Biotropica 49 : 482 – 492 Google Scholar Crossref Search ADS WorldCat Amsbury S , Hunt L, Elhaddad N, Baillie A, Lundgren M, Verhertbruggen Y, Scheller HV, Knox JP, Fleming AJ, Gray JE ( 2016 ) Stomatal function requires pectin de-methyl-esterification of the guard cell wall . Curr Biol 26 : 2899 – 2906 Google Scholar Crossref Search ADS PubMed WorldCat Armour WJ , Barton DA, Law AM, Overall RL ( 2015 ) Differential growth in periclinal and anticlinal walls during lobe formation in Arabidopsis cotyledon pavement cells . Plant Cell 27 : 2484 – 2500 Google Scholar Crossref Search ADS PubMed WorldCat Aylor DE , Parlange JY, Krikorian AD ( 1973 ) Stomatal mechanics . Am J Bot 60 : 163 – 171 Google Scholar Crossref Search ADS WorldCat Bascom CS , Hepler PK, Bezanilla M ( 2018 ) Interplay between ions, the cytoskeleton, and cell wall properties during tip growth . Plant Physiol 176 : 28 – 40 Google Scholar Crossref Search ADS PubMed WorldCat Baskin TI ( 2005 ) Anisotropic expansion of the plant cell wall . Annu Rev Cell Dev Biol 21 : 203 – 222 Google Scholar Crossref Search ADS PubMed WorldCat Bassel GW , Stamm P, Mosca G, Barbier de Reuille P, Gibbs DJ, Winter R, Janka A, Holdsworth MJ, Smith RS ( 2014 ) Mechanical constraints imposed by 3D cellular geometry and arrangement modulate growth patterns in the Arabidopsis embryo . Proc Natl Acad Sci USA 111 : 8685 – 8690 Google Scholar Crossref Search ADS PubMed WorldCat Belteton S , Sawchuk MG, Donohoe BS, Scarpella E, Szymanski DB ( 2018 ) Reassessing the roles of PIN proteins and anticlinal microtubules during pavement cell morphogenesis . Plant Physiol 176 : 432 – 449 Google Scholar Crossref Search ADS PubMed WorldCat Bidhendi AJ , Geitmann A ( 2016 ) Relating the mechanics of the primary plant cell wall to morphogenesis . J Exp Bot 67 : 449 – 461 Google Scholar Crossref Search ADS PubMed WorldCat Bidhendi AJ , Korhonen RK ( 2012 ) A finite element study of micropipette aspiration of single cells: effect of compressibility . Comput Math Methods Med 2012 : 192618 Google Scholar PubMed OpenURL Placeholder Text WorldCat Bolduc JE , Lewis LJ, Aubin CÉ, Geitmann A ( 2006 ) Finite-element analysis of geometrical factors in micro-indentation of pollen tubes . Biomech Model Mechanobiol 5 : 227 – 236 Google Scholar Crossref Search ADS PubMed WorldCat Boudaoud A ( 2010 ) An introduction to the mechanics of morphogenesis for plant biologists . Trends Plant Sci 15 : 353 – 360 Google Scholar Crossref Search ADS PubMed WorldCat Boudon F , Chopard J, Ali O, Gilles B, Hamant O, Boudaoud A, Traas J, Godin C ( 2015 ) A computational framework for 3D mechanical modeling of plant morphogenesis with cellular resolution . PLoS Comput Biol 11 : e1003950 Google Scholar Crossref Search ADS PubMed WorldCat Bozorg B , Krupinski P, Jönsson H ( 2014 ) Stress and strain provide positional and directional cues in development . PLoS Comput Biol 10 : e1003410 Google Scholar Crossref Search ADS PubMed WorldCat Braybrook SA , Peaucelle A ( 2013 ) Mechano-chemical aspects of organ formation in Arabidopsis thaliana: the relationship between auxin and pectin . PLoS ONE 8 : e57813 Google Scholar Crossref Search ADS PubMed WorldCat Campàs O , Mahadevan L ( 2009 ) Shape and dynamics of tip-growing cells . Curr Biol 19 : 2102 – 2107 Google Scholar Crossref Search ADS PubMed WorldCat Carter R , Woolfenden H, Baillie A, Amsbury S, Carroll S, Healicon E, Sovatzoglou S, Braybrook S, Gray JE, Hobbs J, et al. ( 2017 ) Stomatal opening involves polar, not radial, stiffening of guard cells . Curr Biol 27 : 2974 – 2983 Google Scholar Crossref Search ADS PubMed WorldCat Cooke JR , De Baerdemaeker JG, Rand RH, Mang HA ( 1976 ) A finite element shell analysis of guard cell deformations . Trans ASAE 19 : 1107 – 1121 Google Scholar Crossref Search ADS WorldCat Cooke JR , Rand RH, Mang HA, De Baerdemaeker JG, Lee JY ( 2008 ) Shell analysis of elliptical guard cells in higher plants: a review . In JF Abel , JR Cooke , eds, International Conference on Computation of Shell and Spatial Structures. IASS-IACM 2008: “Spanning Nano to Mega,” Cornell University, Ithaca, NY, USA, pp. 723 – 726 Cosgrove DJ ( 2016 ) Plant cell wall extensibility: connecting plant cell growth with cell wall structure, mechanics, and the action of wall-modifying enzymes . J Exp Bot 67 : 463 – 476 Google Scholar Crossref Search ADS PubMed WorldCat Cosgrove DJ ( 2018 ) Diffuse growth of plant cell walls . Plant Physiol 176 : 16 – 27 Google Scholar Crossref Search ADS PubMed WorldCat DeMichele DW , Sharpe PJ ( 1973 ) An analysis of the mechanics of guard cell motion . J Theor Biol 41 : 77 – 96 Dumais J ( 2013 ) Modes of deformation of walled cells . J Exp Bot 64 : 4681 – 4695 Google Scholar Crossref Search ADS PubMed WorldCat Dumais J , Shaw SL, Steele CR, Long SR, Ray PM ( 2006 ) An anisotropic-viscoplastic model of plant cell morphogenesis by tip growth . Int J Dev Biol 50 : 209 – 222 Google Scholar Crossref Search ADS PubMed WorldCat Dyson R , Jensen O ( 2010 ) A fibre-reinforced fluid model of anisotropic plant cell growth . J Fluid Mech 655 : 472 – 503 Google Scholar Crossref Search ADS WorldCat Edwards M , Meidner H, Sheriff D ( 1976 ) Direct measurements of turgor pressure potentials of guard cells. II. The mechanical advantage of subsidiary cells, the spannungsphase, and the optimum leaf water deficit . J Exp Bot 27 : 163 – 171 Google Scholar Crossref Search ADS WorldCat Elliott A , Shaw SL ( 2018 ) Update. Plant cortical microtubule arrays . Plant Physiol 176 : 94 – 105 Google Scholar Crossref Search ADS PubMed WorldCat Fayant P , Girlanda O, Chebli Y, Aubin CÉ, Villemure I, Geitmann A ( 2010 ) Finite element model of polar growth in pollen tubes . Plant Cell 22 : 2579 2593 Google Scholar Crossref Search ADS PubMed WorldCat Forouzesh E , Goel A, Mackenzie SA, Turner JA ( 2013 ) In vivo extraction of Arabidopsis cell turgor pressure using nanoindentation in conjunction with finite element modeling . Plant J 73 : 509 – 520 Google Scholar Crossref Search ADS PubMed WorldCat Forterre Y , Skotheim JM, Dumais J, Mahadevan L ( 2005 ) How the Venus fly trap snaps . Nature 433 : 421 – 425 Google Scholar Crossref Search ADS PubMed WorldCat Franks P , Farquhar G ( 1998 ) A study of stomatal mechanics using the cell pressure probe . Plant Cell Environ 21 : 94 – 100 Google Scholar Crossref Search ADS WorldCat Franks PJ , Farquhar GD ( 2007 ) The mechanical diversity of stomata and its significance in gas-exchange control . Plant Physiol 143 : 78 – 87 Google Scholar Crossref Search ADS PubMed WorldCat Fu Y , Gu Y, Zheng Z, Wasteneys G, Yang Z ( 2005 ) Arabidopsis interdigitating cell growth requires two antagonistic pathways with opposing action on cell morphogenesis . Cell 120 : 687 – 700 Google Scholar Crossref Search ADS PubMed WorldCat Gasser TC , Ogden RW, Holzapfel GA ( 2006 ) Hyperelastic modelling of arterial layers with distributed collagen fibre orientations . J R Soc Interface 3 : 15 – 35 Google Scholar Crossref Search ADS PubMed WorldCat Geitmann A ( 2016 ) Actuators acting without actin . Cell 166 : 15 – 17 Google Scholar Crossref Search ADS PubMed WorldCat Geitmann A , Hush J, Overall R ( 1997 ) Inhibition of ethylene biosynthesis does not block microtubule re-orientation in wounded pea roots . Protoplasma 198 : 135 – 142 Google Scholar Crossref Search ADS WorldCat Geitmann A , Ortega JK ( 2009 ) Mechanics and modeling of plant cell growth . Trends Plant Sci 14 : 467 – 478 Google Scholar Crossref Search ADS PubMed WorldCat Goriely A , Tabor M ( 2003 ) Self-similar tip growth in filamentary organisms . Phys Rev Lett 90 : 108101 Google Scholar Crossref Search ADS PubMed WorldCat Green PB ( 1962 ) Mechanism for plant cellular morphogenesis . Science 138 : 1404 – 1405 Google Scholar Crossref Search ADS PubMed WorldCat Hamant O , Heisler MG, Jönsson H, Krupinski P, Uyttewaal M, Bokov P, Corson F, Sahlin P, Boudaoud A, Meyerowitz EM, et al. ( 2008 ) Developmental patterning by mechanical signals in Arabidopsis . Science 322 : 1650 – 1655 Google Scholar Crossref Search ADS PubMed WorldCat Hofhuis H , Moulton D, Lessinnes T, Routier-Kierzkowska AL, Bomphrey RJ, Mosca G, Reinhardt H, Sarchet P, Gan X, Tsiantis M, et al. ( 2016 ) Morphomechanical innovation drives explosive seed dispersal . Cell 166 : 222 – 233 Google Scholar Crossref Search ADS PubMed WorldCat Huang R , Becker A, Jones I ( 2012 ) Modelling cell wall growth using a fibre-reinforced hyperelastic-viscoplastic constitutive law . J Mech Phys Solids 60 : 750 – 783 Google Scholar Crossref Search ADS WorldCat Huang R , Becker AA, Jones IA ( 2015 ) A finite strain fibre-reinforced viscoelasto-viscoplastic model of plant cell wall growth . J Eng Math 95 : 121 – 154 Google Scholar Crossref Search ADS WorldCat Hwang JU , Gu Y, Lee YJ, Yang Z ( 2005 ) Oscillatory ROP GTPase activation leads the oscillatory polarized growth of pollen tubes . Mol Biol Cell 16 : 5385 – 5399 Google Scholar Crossref Search ADS PubMed WorldCat Jacques E , Verbelen JP, Vissenberg K ( 2013 ) Mechanical stress in Arabidopsis leaves orients microtubules in a ‘continuous’ supracellular pattern . BMC Plant Biol 13 : 163 Google Scholar Crossref Search ADS PubMed WorldCat Jacques E , Verbelen JP, Vissenberg K ( 2014 ) Review on shape formation in epidermal pavement cells of the Arabidopsis leaf . Funct Plant Biol 41 : 914 – 921 Google Scholar Crossref Search ADS PubMed WorldCat Julkunen P , Kiviranta P, Wilson W, Jurvelin JS, Korhonen RK ( 2007 ) Characterization of articular cartilage by combining microscopic analysis with a fibril-reinforced finite-element model . J Biomech 40 : 1862 – 1870 Google Scholar Crossref Search ADS PubMed WorldCat Kha H , Tuble SC, Kalyanasundaram S, Williamson RE ( 2010 ) WallGen, software to construct layered cellulose-hemicellulose networks and predict their small deformation mechanics . Plant Physiol 152 : 774 – 786 Google Scholar Crossref Search ADS PubMed WorldCat Klahre U , Becker C, Schmitt AC, Kost B ( 2006 ) Nt-RhoGDI2 regulates Rac/Rop signaling and polar cell growth in tobacco pollen tubes . Plant J 46 : 1018 – 1031 Google Scholar Crossref Search ADS PubMed WorldCat Korn RW ( 1976 ) Concerning the sinuous shape of leaf epidermal cells . New Phytol 77 : 153 – 161 Google Scholar Crossref Search ADS WorldCat Kost B ( 2008 ) Spatial control of Rho (Rac-Rop) signaling in tip-growing plant cells . Trends Cell Biol 18 : 119 – 127 Google Scholar Crossref Search ADS PubMed WorldCat Kost B , Lemichez E, Spielhofer P, Hong Y, Tolias K, Carpenter C, Chua NH ( 1999 ) Rac homologues and compartmentalized phosphatidylinositol 4,5-bisphosphate act in a common pathway to regulate polar pollen tube growth . J Cell Biol 145 : 317 – 330 Google Scholar Crossref Search ADS PubMed WorldCat Kotchoni SO , Zakharova T, Mallery EL, Le J, El-Assal Sel-D, Szymanski DB ( 2009 ) The association of the Arabidopsis actin-related protein2/3 complex with cell membranes is linked to its assembly status but not its activation . Plant Physiol 151 : 2095 – 2109 Google Scholar Crossref Search ADS PubMed WorldCat Kroeger J , Geitmann A ( 2011 ) Modeling pollen tube growth: feeling the pressure to deliver testifiable predictions . Plant Signal Behav 6 : 1828 – 1830 Google Scholar Crossref Search ADS PubMed WorldCat Kroeger JH , Geitmann A ( 2012 ) Pollen tube growth: getting a grip on cell biology through modeling . Mech Res Commun 42 : 32 – 39 Google Scholar Crossref Search ADS WorldCat Kroeger JH , Geitmann A, Grant M ( 2008 ) Model for calcium dependent oscillatory growth in pollen tubes . J Theor Biol 253 : 363 – 374 Google Scholar Crossref Search ADS PubMed WorldCat Lee JY ( 1986 ) A finite element for shell analysis and its application to biological objects. PhD thesis. Cornell University, Ithaca, NY Lin D , Cao L, Zhou Z, Zhu L, Ehrhardt D, Yang Z, Fu Y ( 2013 ) Rho GTPase signaling activates microtubule severing to promote microtubule ordering in Arabidopsis . Curr Biol 23 : 290 – 297 Google Scholar Crossref Search ADS PubMed WorldCat Majda M , Grones P, Sintorn IM, Vain T, Milani P, Krupinski P, Zagórska-Marek B, Viotti C, Jönsson H, Mellerowicz EJ, et al. ( 2017 ) Mechanochemical polarization of contiguous cell walls shapes plant pavement cells . Dev Cell 43 : 290 – 304 Mathur J ( 2004 ) Cell shape development in plants . Trends Plant Sci 9 : 583 – 590 Google Scholar Crossref Search ADS PubMed WorldCat Mathur J , Spielhofer P, Kost B, Chua N ( 1999 ) The actin cytoskeleton is required to elaborate and maintain spatial patterning during trichome cell morphogenesis in Arabidopsis thaliana . Development 126 : 5559 – 5568 Google Scholar Crossref Search ADS PubMed WorldCat Meidner H , Mansfield TA ( 1968 ) Physiology of Stomata. Tata McGraw-Hill , Bombay Google Scholar Google Preview OpenURL Placeholder Text WorldCat COPAC Merced A , Renzaglia K ( 2014 ) Developmental changes in guard cell wall structure and pectin composition in the moss Funaria: implications for function and evolution of stomata . Ann Bot 114 : 1001 – 1010 Google Scholar Crossref Search ADS PubMed WorldCat Niklas KJ ( 1992 ) Plant biomechanics: an engineering approach to plant form and function. PhD thesis. University of Chicago, Chicago Oelschlägel B , Gorb S, Wanke S, Neinhuis C ( 2009 ) Structure and biomechanics of trapping flower trichomes and their role in the pollination biology of Aristolochia plants (Aristolochiaceae) . New Phytol 184 : 988 – 1002 Google Scholar Crossref Search ADS PubMed WorldCat Palin R , Geitmann A ( 2012 ) The role of pectin in plant morphogenesis . Biosystems 109 : 397 – 402 Google Scholar Crossref Search ADS PubMed WorldCat Panteris E , Galatis B ( 2005 ) The morphogenesis of lobed plant cells in the mesophyll and epidermis: organization and distinct roles of cortical microtubules and actin filaments . New Phytol 167 : 721 – 732 Google Scholar Crossref Search ADS PubMed WorldCat Parre E , Geitmann A ( 2005 ) Pectin and the role of the physical properties of the cell wall in pollen tube growth of Solanum chacoense . Planta 220 : 582 – 592 Google Scholar Crossref Search ADS PubMed WorldCat Peaucelle A , Braybrook SA, Le Guillou L, Bron E, Kuhlemeier C, Höfte H ( 2011 ) Pectin-induced changes in cell wall mechanics underlie organ initiation in Arabidopsis . Curr Biol 21 : 1720 – 1726 Google Scholar Crossref Search ADS PubMed WorldCat Raschke K ( 1975 ) Stomatal action . Annu Rev Plant Physiol 26 : 309 – 340 Google Scholar Crossref Search ADS WorldCat Raven JA ( 2014 ) Speedy small stomata? J Exp Bot 65 : 1415 – 1424 Google Scholar Crossref Search ADS PubMed WorldCat Renzaglia KS , Villarreal JC, Piatkowski BT, Lucas JR, Merced A ( 2017 ) Hornwort stomata: architecture and fate shared with 400 million year old fossil plants without leaves . Plant Physiol 174 : 788–797 Google Scholar Crossref Search ADS Google Preview WorldCat COPAC Roelfsema MRG , Hedrich R ( 2005 ) In the light of stomatal opening: new insights into ‘the Watergate’ . New Phytol 167 : 665 – 691 Google Scholar Crossref Search ADS PubMed WorldCat Rojas ER , Hotton S, Dumais J ( 2011 ) Chemically mediated mechanical expansion of the pollen tube cell wall . Biophys J 101 : 1844 – 1853 Google Scholar Crossref Search ADS PubMed WorldCat Rui Y , Anderson CT ( 2016 ) Functional analysis of cellulose and xyloglucan in the walls of stomatal guard cells of Arabidopsis . Plant Physiol 170 : 1398 – 1419 Google Scholar Crossref Search ADS PubMed WorldCat Sampathkumar A , Krupinski P, Wightman R, Milani P, Berquand A, Boudaoud A, Hamant O, Jönsson H, Meyerowitz EM ( 2014 ) Subcellular and supracellular mechanical stress prescribes cytoskeleton behavior in Arabidopsis cotyledon pavement cells . eLife 3 : e01967 Google Scholar Crossref Search ADS PubMed WorldCat Sanati Nezhad A , Geitmann A ( 2015 ) Tip growth in walled cells: cellular expansion and invasion mechanisms . In C Cuerrier , A Pelling , eds, Cells, Forces and the Microenvironment. Pan Stanford Publishing , pp 335 – 356 Google Scholar Google Preview OpenURL Placeholder Text WorldCat COPAC Sanati Nezhad A , Naghavi M, Packirisamy M, Bhat R, Geitmann A ( 2013 ) Quantification of the Young’s modulus of the primary plant cell wall using Bending-Lab-On-Chip (BLOC) . Lab Chip 13 : 2599 – 2608 Google Scholar Crossref Search ADS PubMed WorldCat Sharpe PJ , Wu HI ( 1978 ) Stomatal mechanics: volume changes during opening . Plant Cell Environ 1 : 259 – 268 Google Scholar Crossref Search ADS WorldCat Sun EQ ( 2006 ) Shear locking and hourglassing in MSC Nastran, ABAQUS, and ANSYS . In MSC Software Corporation’s 2006 Americas Virtual Product Development Conference: Evolution to Enterprise Simulation. MSC Software , Santa Ana, CA , presentation 27, pp. 1 – 9 Szymanski DB , Marks MD, Wick SM ( 1999 ) Organized F-actin is essential for normal trichome morphogenesis in Arabidopsis . Plant Cell 11 : 2331 – 2347 Google Scholar Crossref Search ADS PubMed WorldCat Szymanski DB , Staiger CJ ( 2018 ) The actin cytoskeleton: functional arrays for cytoplasmic organization and cell shape control . Plant Physiol 176 : 106 – 118 Google Scholar Crossref Search ADS PubMed WorldCat Tissier A ( 2012 ) Glandular trichomes: what comes after expressed sequence tags? Plant J 70 : 51 – 68 Google Scholar Crossref Search ADS PubMed WorldCat Torode TA , O’Neill RE, Marcus SE, Cornuault V, Pose-Albacete S, Lauder RP, Kracun SK, Gro Rydahl M, Andersen MCF, Willats WGT, et al. ( 2018 ) Branched pectic galactan in phloem-sieve-element cell walls: implications for cell mechanics . Plant Physiol (in press) https://doi.org/10.1104/pp.17.01568 Google Scholar OpenURL Placeholder Text WorldCat Von Mohl H ( 1856 ) Welche Ursachen bewirken die Erweiterung und Verengung der Spaltöffnungen . Bot Z 14 : 697 – 704 Google Scholar OpenURL Placeholder Text WorldCat Williamson RE ( 1990 ) Alignment of cortical microtubules by anisotropic wall stresses . Funct Plant Biol 17 : 601 – 613 Google Scholar Crossref Search ADS WorldCat Woolfenden HC , Bourdais G, Kopischke M, Miedes E, Molina A, Robatzek S, Morris RJ ( 2017 ) A computational approach for inferring the cell wall properties that govern guard cell dynamics . Plant J 92 : 5 – 18 Google Scholar Crossref Search ADS PubMed WorldCat Xu T , Wen M, Nagawa S, Fu Y, Chen JG, Wu MJ, Perrot-Rechenmann C, Friml J, Jones AM, Yang Z ( 2010 ) Cell surface- and rho GTPase-based auxin signaling controls cellular interdigitation in Arabidopsis . Cell 143 : 99 – 110 Google Scholar Crossref Search ADS PubMed WorldCat Yanagisawa M , Desyatova AS, Belteton SA, Mallery EL, Turner JA, Szymanski DB ( 2015 ) Patterning mechanisms of cytoskeletal and cell wall systems during leaf trichome morphogenesis . Nat Plants 1 : 15014 Yi H , Puri VM ( 2012 ) Architecture-based multiscale computational modeling of plant cell wall mechanics to examine the hydrogen-bonding hypothesis of the cell wall network structure model . Plant Physiol 160 : 1281 – 1292 Google Scholar Crossref Search ADS PubMed WorldCat Zerzour R , Kroeger J, Geitmann A ( 2009 ) Polar growth in pollen tubes is associated with spatially confined dynamic changes in cell mechanical properties . Dev Biol 334 : 437 – 446 Google Scholar Crossref Search ADS PubMed WorldCat Zhang C , Halsey LE, Szymanski DB ( 2011 ) The development and geometry of shape change in Arabidopsis thaliana cotyledon pavement cells . BMC Plant Biol 11 : 27 Google Scholar Crossref Search ADS PubMed WorldCat Ziegenspeck A ( 1955 ) Das Vorkommen von Fila in radialer Anordnung in den Schliesszellen . Protoplasma 44 : 385 – 388 Google Scholar Crossref Search ADS WorldCat Author notes 1 Research in the Geitmann lab is funded by Discovery and Accelerator Grants from the National Science and Engineering Research Council of Canada and by the Canada Research Chair program. 2 Address correspondence to [email protected]. A.J.B. and A.G. wrote the manuscript; A.J.B. prepared the figures. [OPEN] Articles can be viewed without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.17.01684 © 2018 American Society of Plant Biologists. All Rights Reserved. © The Author(s) 2018. Published by Oxford University Press on behalf of American Society of Plant Biologists. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.

Bidhendi, Amir J.; Geitmann, Anja

doi: 10.1104/pp.17.01684pmid: 29229695