MOLECULAR BASIS of SALT TOLERANCE in Physcomitrella Patens MODEL PLANT: POTASSIUM HOMEOSTASIS and PHYSIOLOGICAL ROLES of CHX TRANSPORTERS

UNIVERSIDAD POLITÉCNICA DE MADRID ESCUELA TÉCNICA SUPERIOR DE INGENIEROS AGRÓNOMOS DEPARTAMENTO DE BIOTECNOLOGÍA

MOLECULAR BASIS OF SALT TOLERANCE IN Physcomitrella patens MODEL PLANT: POTASSIUM HOMEOSTASIS AND PHYSIOLOGICAL ROLES OF CHX TRANSPORTERS.

Director: Alonso Rodríguez Navarro Profesor Emérito, E.T.S.I. Agrónomos Universidad Politécnica de Madrid

TESIS DOCTORAL

SHADY ABDEL MOTTALEB

MADRID, 2013

MOLECULAR BASIS OF SALT TOLERANCE IN Physcomitrella patens MODEL PLANT: POTASSIUM HOMEOSTASIS AND PHYSIOLOGICAL ROLES OF CHX TRANSPORTERS.

Memoria presentada por SHADY ABDEL MOTTALEB para la obtención del grado de Doctor por la Universidad Politécnica de Madrid

Fdo. Shady Abdel Mottaleb

VºBº Director de Tesis:

Fdo. Dr. Alonso Rodríguez-Navarro Profesor emérito Departamento de Biotecnología ETSIA- Universidad Politécnica de Madrid

Madrid, Septiembre 2013

To my family

ACKNOWLEDGEMENTS

This work would not have been possible without the collaboration and inspiration of many people. First of all, I would like to express my deep appreciation to my supervisor and mentor Alonso Rodríguez Navarro for giving me this once-in-a- lifetime opportunity to both studying what I like most and developing my professional career. For integrating me in his research group, introducing me to novel research topics and for his constant support at both personal and professional level. I owe a debt of gratitude to Rosario Haro for her unconditional help, ongoing support and care during my thesis. For teaching me to pay attention to the details of everything and for her immense efforts in supervising all the experiments of this thesis. I also feel grateful to Begoña Benito for her constant help and encouragement, for her useful advice and for always being available to answer my questions at anytime with enthusiasm and a pleasant smile. I would like to thank a lot Blanca Garciadeblás for her constant encouragement and help in many important experiments of this thesis… and for bearing my constant flow of silly (and not so silly) questions during the past four years! I would also like to thank Mª Antonia Bañuelos for her useful advices and for our interesting conversations.

I wish to express my appreciation to my colleagues in the research group. Their help, comments and unique expertise helped me to be accurate, thorough, and balanced throughout the course of this work. Thank you for the great help and for all the good laughs we shared together during the coffee breaks! Ana Claudia Ureta for her help and funny conversations. Marcel Veldhuizen for his help and advice on many useful shortcuts for experimental protocols! Angela Sáez for her support and understanding. Rocío Álvarez for her dedicated help in many experiments, her constant encouragement and for our many funny conversations. A special thanks to my dear friend Ana Fraile for teaching me her experience on the different experimental techniques I used in this work, for her constant help, advices and constructive critics. For being patient with my many questions and for all the after-work activities and trips shared together!

Thanks to our neighbor “Rhizobium lab.” members, professors Tomás Argüeso, Pepe Palacios, Juan Imperial, Luis Rey and Belén Brito… and thanks to Bea, Carmen, David, Laura, Marta, Mónica, Rosabel for their collaboration and good times shared during coffee and lunch breaks. A special thanks to my good friend and iii

“bus partner” Anabel for all the good times, laughs and conversations we had during our coffee and lunch breaks, in the bus, trips and everywhere!

I deeply appreciate the technical help and dedication of Dr. Pablo Melendi in the confocal microscopy experiments.

My gratitude is extended to all my friends in the “Plant virus lab.” Antolín, Jean-Michel, Manuel & Manuel, Nils and Pablo, and a special thanks to my “lunch partner” Miguel Ángel for all the good laughs and for teaching me so many aspects of the Spanish culture, typical foods, slang language, sayings and many more!

I would also like to thank a lot many people that I was lucky to know in the Centro de Biotecnología y genómica de plantas (CBGP), and had the opportunity to share with them many nice moments: Alessandro, Amir, Angela, Anja, Bahia, Carlos, César, Elena, Eva, Fátima, Jan, Julia, Inés, Irene, Mar, Nancy, Pilar, Ruth, Sara, Simon and Vivi.

Many thanks to my CBGP-Basketball team members for all the entertaining matches played under the burning sun, rain or snow! Thanks to Alejandro, Antonio, Chechu, Jorge, Laura, Miguel Ángel, Nacho and Nils.

I would also like to thank my housemates and friends in Madrid Alba, Ali, Amr, Basem, Chakib, Gemma, Haytham, Isabel, Jorge, Jose, Juan, Lurdes, Mohamed, Montse, Osama, Vicky and Yasser for keeping my morale high at hard times. It would have been very different without your care and generosity!

Finally, I feel extremely lucky to have an exceptional family that trusted me and was so patient and understanding even under all the negative circumstances they have suffered during my absence. Their love has been strong, unconditional, and selfless. They have made everything I’ve accomplished possible. I am forever in your debt!

RESUMEN El suelo salino impone un estrés abiótico importante que causa graves problemas en la agricultura ya que la mayoría de los cultivos se ven afectados por la salinidad debido a efectos osmóticos y tóxicos. Por ello, la contaminación y la escasez de agua dulce, la salinización progresiva de tierras y el aumento exponencial de la población humana representan un grave problema que amenaza la seguridad alimentaria mundial para las generaciones futuras. Por lo tanto, aumentar la tolerancia a la salinidad de los cultivos es un objetivo estratégico e ineludible para garantizar el suministro de alimentos en el futuro.

Mantener una óptima homeostasis de K+ en plantas que sufren estrés salino es un objetivo importante en el proceso de obtención de plantas tolerantes a la salinidad. Aunque el modelo de la homeostasis de K+ en las plantas está razonablemente bien descrito en términos de entrada de K+, muy poco se sabe acerca de los genes implicados en la salida de K+ o de su liberación desde la vacuola. En este trabajo se pretende aclarar algunos de los mecanismos implicados en la homeostasis de K+ en plantas. Para ello se eligió la briofita Physcomitrella patens, una planta no vascular de estructura simple y de fase haploide dominante que, entre muchas otras cualidades, hacen que sea un modelo ideal. Lo más importante es que no sólo P. patens es muy tolerante a altas concentraciones de Na+, sino que también su posición filogenética en la evolución de las plantas abre la posibilidad de estudiar los cambios claves que, durante el curso de la evolución, se produjeron en las diversas familias de los transportadores de K+.

Se han propuesto varios transportadores de cationes como candidatos que podrían tener un papel en la salida de K+ o su liberación desde la vacuola, especialmente miembros de la familia CPA2 que contienen las familias de transportadores KEA y CHX. En este estudio se intenta aumentar nuestra comprensión de las funciones de los transportadores de CHX en las células de las plantas usando P. patens, como ya se ha dicho. En esta especie, se han identificado cuatro genes CHX, PpCHX1-4. Dos de estos genes, PpCHX1 y PpCHX2, se expresan aproximadamente al mismo nivel que el gen PpACT5, y los otros dos genes muestran una expresión muy baja. La expresión de PpCHX1 y PpCHX2 en mutantes de Escherichia coli defectivos en el transporte de K+ restauraron el crecimiento de esta cepa en medios con bajo contenido de K+, lo que

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sugiere que la entrada de K+ es energizada por un mecanismo de simporte con H+. Por otra parte, estos transportadores suprimieron el defecto asociado a la mutación kha1 en Saccharomyces cerevisiae, lo que sugiere que podrían mediar un antiporte en K+/H+. La proteína PpCHX1-GFP expresada transitoriamente en protoplastos de P. patens co- localizó con un marcador de Golgi. En experimentos similares, la proteína PpCHX2- GFP localizó aparentemente en la membrana plasmática y tonoplasto. Se construyeron las líneas mutantes simples de P. patens ∆Ppchx1 y ∆Ppchx2, y también el mutante doble ∆Ppchx2 ∆Pphak1. Los mutantes simples crecieron normalmente en todas las condiciones ensayadas y mostraron flujos de entrada normales de K+ y Rb+; la mutación ∆Ppchx2 no aumentó el defecto de las plantas ∆Pphak1. En experimentos a largo plazo, las plantas ∆Ppchx2 mostraron una retención de Rb+ ligeramente superior que las plantas silvestres, lo que sugiere que PpCHX2 promueve la transferencia de Rb+ desde la vacuola al citosol o desde el citosol al medio externo, actuando en paralelo con otros transportadores. Sugerimos que transportadores de K+ de varias familias están involucrados en la homeostasis de pH de orgánulos ya sea mediante antiporte K+/H+ o simporte K+-H+.

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ABSTRACT Soil salinity is a major abiotic stress causing serious problems in agriculture as most crops are affected by it. Moreover, the contamination and shortage of freshwater, progressive land salinization and exponential increase of human population aggravates the problem implying that world food security may not be ensured for the next generations. Thus, a strategic and an unavoidable goal would be increasing salinity tolerance of plant crops to secure future food supply.

Maintaining an optimum K+ homeostasis in plants under salinity stress is an important trait to pursue in the process of engineering salt tolerant plants. Although the model of K+ homeostasis in plants is reasonably well described in terms of K+ influx, very little is known about the genes implicated in K+ efflux or release from the vacuole. In this work, we aim to clarify some of the mechanisms involved in K+ homeostasis in plants. For this purpose, we chose the bryophyte plant Physcomitrella patens, a nonvascular plant of simple structure and dominant haploid phase that, among many other characteristics, makes it an ideal model. Most importantly, not only P. patens is very tolerant to high concentrations of Na+, but also its phylogenetic position in land plant evolution opens the possibility to study the key changes that occurred in K+ transporter families during the course of evolution.

Several cation transporter candidates have been proposed to have a role in K+ efflux or release from the vacuole especially members of the CPA2 family which contains the KEA and CHX transporter families. We intended in this study to increase our understanding of the functions of CHX transporters in plant cells using P. patens, in which four CHX genes have been identified, PpCHX1-4. Two of these genes, PpCHX1 and PpCHX2, are expressed at approximately the same level as the PpACT5 gene, but the other two genes show an extremely low expression. PpCHX1 and PpCHX2 restored growth of Escherichia coli mutants on low K+-containing media, suggesting they mediated K+ uptake that may be energized by symport with H+. In contrast, these genes suppressed the defect associated to the kha1 mutation in Saccharomyces cerevisiae, which suggest that they might mediate K+/H+ antiport. PpCHX1-GFP protein transiently expressed in P. patens protoplasts co-localized with a Golgi marker. In similar experiments, the PpCHX2-GFP protein appeared to localize to tonoplast and plasma

membrane. We constructed the ∆Ppchx1 and ∆Ppchx2 single mutant lines, and the ∆Ppchx2 ∆Pphak1 double mutant. Single mutant plants grew normally under all the conditions tested and exhibited normal K+ and Rb+ influxes; the ∆Ppchx2 mutation did not increase the defect of ∆Pphak1 plants. In long-term experiments, ∆Ppchx2 plants showed a slightly higher Rb+ retention than wild type plants, which suggests that PpCHX2 mediates the transfer of Rb+ from either the vacuole to the cytosol or from the cytosol to the external medium in parallel with other transporters. We suggest that K+ transporters of several families are involved in the pH homeostasis of organelles by mediating either K+/H+ antiport or K+-H+ symport.

CONTENTS DEDICATION ...…………………………………….…………………………..……...i ACKNOWLEDGMENTS ...…………………………….……..…...... ……..………..iii RESUMEN ...…………………………………………………...... …...vii ABSTRACT ...……………..……………………………………….....….….…….…..ix CONTENTS ...………………………………………………..…………….……...... xi LIST OF ABBREVIATIONS ...………………………...………………………..…..xv 1. INTRODUCTION ...…………………………………………………….……..……3 1.1. Objectives ...……………….………………………………………….……5 2. REVIEW OF LITERATURE ...……………………...…………………...…...……9 2.1. Sodium toxicity and tolerance in plants …………………...... 9 2.1.1. Present situation of world population and food security ...... 9 2.1.2. Salt-affected soils …………………………………….……...... 10 2.1.3. Salinity stress: definition and negative consequences ..…..….11 2.1.3.1. Osmotic stress ……………….……...... …11 2.1.3.2. Sodium imbalance and toxicity stress .……….....…..13 2.1.3.3. Other effects of salinity stress ……….……..…...…...14 2.1.4. Sodium transport inside the plant .……………………..…...... 14 2.1.4.1. Sodium influx into the plant ………...……………....15 2.1.4.2. Sodium efflux …………..……………………….……17 2.1.4.3. Radial transport of sodium across the root ..…...…..18 2.1.4.4. Vacuolar sequestration of sodium …….….…...….…19 2.1.4.5. Xylem loading ……..……….……..………...………..20 2.1.4.6. Sodium retrieval from the xylem and removal from the shoot ...... 21 2.1.5. Can sodium be beneficial for plants? ...... 22 2.1.6. Salinity tolerance mechanisms in plants …….…….……...…..23 2.2. Potassium homeostasis …….……………………….………..…...... …...29 2.2.1. Physiological functions of potassium in plants …...…….....….29 2.2.2. Potassium deficiency symptoms …...…………….…...….…....30 2.2.3. Soil and plant concentrations of potassium …....….…..……...31 2.2.4. Potassium homeostasis and transport in plants …..…..…..….32 2.2.4.1. Long distance potassium transport …………..…...... 33 xi

2.2.4.1.1. Potassium influx …..……..…………………33 2.2.4.1.2. Potassium accumulation and release from vacuole ...…………………………...... 36 2.2.4.1.3. Potassium efflux ………..…...……...………40 2.2.4.1.4. Potassium loading to the xylem ……….…..42 2.2.4.1.5. Potassium unloading from the xylem, phloem loading/unloading and recirculation …………..…….43 2.2.4.2. Potassium homeostasis under salinity stress …….…43 2.2.5. Poorly studied cation transporter families: a new source of information on potassium homeostasis?...... 45 2.3. The model plant physcomitrella patens ………....………....…………....47 2.3.1. Biological and evolutionary significance of bryophytes …..…47 2.3.2. The moss Physcomitrella patens: a model plant for the new millennium ………………………………...………………...…...... ….49 3. MATERIALS AND METHODS …………...………...………………………...... 57 3.1. Biological materials …………...……………...……………………....….57 3.1.1. Bacterial strains and culture conditions …………..………….57 3.1.2. Yeast strains and culture conditions …………….…..……...... 58 3.1.3. Physcomitrella patens ecotype and culture conditions …….....60 3.1.3.1. Growth and propagation conditions of protonema and gametophores ……………………..……………….………….60 3.1.3.2. Culture conditions for obtaining sporangiophores…63 3.1.3.3. Conservation of Physcomitrella patens ……………...64 3.1.4. Plasmids …...……………………………………………...….…64 3.2. DNA manipulation techniques …….………………………………....…65 3.2.1. Bacterial plasmidic DNA purification …………………..……65 3.2.2. Plant genomic DNA purification …...…………..…...….…..…66 3.2.3. DNA Quantification …...………………...…….………….....…66 3.2.4. DNA amplification by PCR …..………………..………...... …67 3.2.5. Cloning and purification of PCR fragments ….……………...67 3.2.6. DNA Digestion and ligation of DNA fragments …..…...……..67 3.3. RNA manipulation techniques …………………..………………...... …68 3.3.1. Purification of total RNA of plants ..…………………….....…68 xii

3.3.2. RNA quantification …..……………….…………...……..…..68 3.3.3. cDNA synthesis by RT-PCR and study of the genic expression by quantitative real time PCR (qRT-PCR) ….…………….……..…69 3.3.4. Cloning of PpCHX1 and PpCHX2 cDNAs, and The in vitro construction of the PpCHX2.1 cDNA sequence …………...………..69 3.4. Methods of cellular transformation …..…………..………………...... 72 3.4.1. Transformation of bacteria and yeast ….……...…...…..….....72 3.4.2. Transformation of protoplasts of Physcomitrella ….……...…74 3.5. Construction of Physcomitrella knock-out mutants by genetic replacement (homologous recombination) …………………………..…...…78 3.5.1. Design of the construction for the disruption of PpCHX1 and PpCHX2 gene ………………………………..……………………....78 3.5.2. Screening and analysis of putative Physcomitrella knock out clones by PCR ………………….….…….…………...……………….80 3.6. Design of gene fusions with GFP to determine the subcellular localization of the transporters …………..………………..…………...…….82 3.7. Functional complementation “drop tests” of yeast and bacteria mutants ………………………………………………………………………………….84 3.8. Cation contents and cation uptake experiments in yeast …..……….…84 3.9. Cation fluxes tests in Physcomitrella plants …..………………...... ….....85 3.10. Software and bioinformatic Tools ………………………...... ………....86 4. RESULTS ……………….…………………...………………………………..……89 4.1. The CHX transporters of P. patens …………….……………….....……89 4.2. PpCHX1 and PpCHX2 gene expression and cDNAs cloning ….……....95 4.3. Alternative PpCHX2 proteins ……..….…….………………...... ……....98 4.4. Subcellular localization of PpCHX1-GFP, PpCHX2-GFP and PpCHX2.1-GFP protein fusions in yeast cells and Physcomitrella patens protoplasts…………………..…………………………………………..…..…99 4.5. Growth rescue of Escherichia coli mutants ……….……..…...……..101 4.6. PpCHX1 and PpCHX2 complement the kha1 mutation in yeast …....105 4.7. Functional analysis of PpCHXs in ∆Ppchx1 and ∆Ppchx2 plants …...113 5. DISCUSSION ………….……………..…………….……………………………..123

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5.1. Analysis in silico and cloning of CHX transporters of Physcomitrella patens ……….....………………………….………………….……...….…….123 5.2. Functional analysis of PpCHXs in heterlogous systems of expression ……………………………………………………………………………...…127 5.2.1. Escherichia coli ……………..……………………………..….127 5.2.2. Saccharomyces cerevisiae ….………..………………………..130 5.3. Proposed models of action for PpCHXs …………………...….…....…133 5.4. Functional analysis of PpCHXs in Physcomitrella patens ...…...……..134 6. CONCLUSIONS ....…….…………..…………………..……..…………...……...141 7. REFERENCES……………………….………….………...……………...…..…..145

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LIST OF ABREVIATIONS

ATPase Protein that transports a substrate driven by catalysis of ATP Amp Ampicillin AP Arginine phosphate medium ATP Adenosine triphosphate BLAST Basic local alignment search tool Bp Base pair cDNA Complementary DNA CHX Cation H+ exchanger CPA Cation proton antiporter family CTAB Cetyltrimethylammonium bromide C-terminus Carboxyl terminus d Days DNA Deoxyribonucleic acid EDTA Ethylenediaminetetraacetic acid EST Expressed sequence tag FAO Food and Agriculture Organization g Gr a ms GFP Green fluorescent protein h Hours HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid Hyg Hygromicin KEA K+ efflux antiporter KFM K+ free medium L Liter Km “Michaelis constant”: Substrate concentration at which the reaction rate is half of Vm a x LB Lysogeny Broth medium M Molar MES 2-(N-morpholino)ethanesulfonic acid min Minutes MM Minimal medium for bacteria mRNA Messenger RNA N-terminus Amino terminus OD Optical density ORF Open Reading frame PCR Polymerase chain reaction PEG Polyethylene glycol pH Negative logarithmic value of H+ concentration PM Plasma membrane PRM-B Protoplast regeneration medium for the bottom layer medium PRM-L Liquid protoplast regeneration medium PRM-T Protoplast regeneration medium for the top layer medium RNA Ribonucleic acid RNase Ribonuclease ROS Reactive oxygen species r.p.m Revolutions per minute RT-PCR Reverse transcription PCR xv

SD Synthetic defined minimal medium SDS Sodium dodecyl sulfate sec Seconds TAE Tris-acetic acid-EDTA buffer TAPS N-Tris(hydroxymethyl)methyl-3-aminopropanesulfonic acid TBE Tris-boric acid-EDTA buffer TE Tris-EDTA buffer TGN Trans-Golgi network Tris Tris(hydroxymethyl)aminomethane Vm a x Maximum rate achieved by the system at saturating substrate concentrations w/v Weight per volume YFP Yellow fluorescent protein YNB Yeast-Nitrogen-Base YPD Yeast-extract Peptone Dextrose medium Zeo Zeocin

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1. INTRODUCTION

1. INTRODUCTION Soil salinity is a major abiotic stress causing serious problems in agriculture. It is estimated that more than 800 million ha of land throughout the world are salt affected (FAO 2008) of which, 32 million ha are dryland agriculture farms and 45 million ha are irrigated lands (Munns and Tester 2008). Among the many types of salt-affected soils, those with high sodium chloride (NaCl) levels are the most common. NaCl disrupts metabolic processes in plants and hinders growth through four different mechanisms: osmotic stress, inhibition of potassium (K+) uptake, toxicity to cytosolic enzymes, and oxidative stress and cell death. Halophytes are adapted to high NaCl environments, but this adaptation does not apply to glycophytes among which most crop plants are included, and as a consequence crop productivity is greatly affected by NaCl (Flowers 2004). Moreover, the contamination and shortage of freshwater, progressive land salinization and exponential increase of human population only aggravates the problem implying that world food security is not ensured for the next generations. Thus, a strategic and an unavoidable goal would be increasing salinity tolerance of plant crops to secure future food supply (Flowers and Flowers 2005). Salinity tolerance in plants is a complex polygenic trait difficult to explain in simple biochemical or biological processes. It is predominately under control of additive effects with large environment x genotype interaction effects where an evident trade-off between yield and salinity tolerance is observed. Salinity tolerance in glycophytes depends upon plant morphology, ions compartmentalization and fluxes, production of compatible solutes, regulation of transpiration, control of ion movement and cytoplasmic tolerance (Flowers 2004). A special and important trait in salinity tolerance is the plants’ ability to maintain a high cytosolic K+/Na+ ratio. Evidence shows that among the effects of high Na+ concentration in the soil, the reduction of K+ uptake as a result of the direct competition between Na+ and K+ for uptake sites at the plasma membrane is one of the most aggressive effects. Furthermore, high Na+ levels induce plasma membrane (PM) depolarization, which in turn has two negative consequences on K+/Na+ ratio. First, at the normal K+ concentration in the soil solution, PM depolarization makes passive K+ uptake through inward-rectifying K+ channels impossible in thermodynamic terms and increases K+ efflux through outward-rectifying K+ channels, thus causing massive K+ loss from the cells (Shabala and Cuin 2008). Consequently, K+ homeostasis and K+/Na+ 3

ratio in the cell are perturbed reducing the optimal K+ concentrations required for the activation of many enzymes, lowering K+ availability as a cellular osmoticum for rapid cell expansion and altering several electrogenic transport processes. As a result, these K+ deficient plants show lower water content, impaired stomatal regulation, reduced transpiration, impaired phloem transport, and reduced photosynthetic capacity. Moreover, visible symptoms of K+ deficiency including scorching along the margins of older leaves, reduced growth and a greater susceptibility to abiotic stress, pests and diseases are observed (Mengel et al. 2001, Amtmann et al. 2008, Marschner 2012). Ion homeostasis of K+ and Na+ in the cytoplasm depends on several fluxes, the most important being K+ and Na+ influxes, effluxes, and accumulation in organelles and other internal membranes. Numerous K+ and Na+ transporters involved in maintaining K+ homeostasis and a suitable cytosolic K+/Na+ ratio have been described in plants. Among them, three gene families encoding K+ transporters have been identified in the model plant Arabidopsis thaliana: 1) KT/HAK/KUP, 2) TRK/HKT and 3) CPA, subdivided in CPA1 y CPA2 families (Mäser et al. 2001). Although the model of K+ homeostasis in plants is reasonably well described in terms of K+ influx, very little is known about the genes implicated in K+ efflux from the PM or release from the vacuole. Several cation transporter candidates have been proposed to have a role in K+ efflux or release from the vacuole especially members of the CPA2 family which contains the KEA and CHX gene families (Britto and Kronzucker 2006). The CPA2 family is considered the least studied transporter family in plants in contrast to CPA1 family, which comprises fairly well characterized transporters such as NHX1 and SOS1, and a limited number of works studying A. thaliana CPA2 family has been published in the past few years. The moss Physcomitrella patens is becoming increasingly used by the scientific community as a model plant for the study of plant cell biology and, by virtue of its basal position in land plant phylogeny, for comparative analysis of plant gene function and development (Rensing et al. 2008). Moreover, the tolerance of P.patens to high-salinity environments also makes it an ideal candidate for studying the molecular mechanisms by which plants respond to salinity stress (Benito and Rodríguez-Navarro 2003, Wang et al. 2008, Fraile-Escanciano et al. 2010). To our knowledge, no studies on CPA2 family of the model plant Physcomitrella patens have seen the light so far in contrast to those on A. thaliana. 4

1.1. OBJECTIVES In an attempt to further elucidate the K+ homeostasis model and particularly the possible transporters involved in K+ efflux and release from the vacuole, the general objective set in this thesis is the functional characterization of two CHX transporters, PpCHX1 and PpCHX2, of the CPA2 family of Physcomitrella patens. The following specific objectives have been pursued: 1. Bioinformatic analysis of CPA2 family in Physcomitrella patens with special emphasis on PpCHX gene family. 2. Analysis of PpCHX gene expression by quantitative real time PCR (qRT-PCR). 3. Cloning of PpCHX1 and PpCHX2 transporters and their functional characterization in different heterologous expression systems like Escherichia coli and Sacharomyces cereviseae. 4. Subcellular localization of PpCHX1-GFP and PpCHX2-GFP protein fusions in Sacharomyces cereviseae cells and Physcomitrella patens protoplasts. 5. Construction and characterization of ∆Ppchx1 and ∆Ppchx2 knock-out mutants of Physcomitrella patens.

2. REVIEW OF LITERATURE

2. REVIEW OF LITERATURE 2.1. SODIUM TOXICITY AND TOLERANCE IN PLANTS 2.1.1. Present situation of world population and food security Over 800 million people are under-nourished in the developing world. Of them, 232 million are in India, 200 million in sub-Saharan Africa, 112 million in China, 152 million in Asia and the Pacific, 56 million in Latin America and 40 million in the Near East and North Africa (UN Millennium Project 2003). Also, world population is expected to increase over 8 billion by the year 2020 and it is predicted that at least 10 billion people will be hungry and malnourished by the end of this century (FAO 2003). Furthermore, world cereal demand will likely increase by 50%, driven strongly by rapidly growing animal feed use and meat consumption (Borlaug 2007) as well as the rapidly increasing use of cereals in biofuel production. This increase in food demands is not a new situation; 50 years ago, population growth threatened to overtake food production. Thus, to reduce the food insecurity, crop production would have to be doubled and produced in more environmentally sustainable ways (Borlaug and Dowswell 2005). This can be achieved by expanding cultivable land area or by increasing per hectare crop productivity. However, the enhancement in crop production due to expansion in growing area was only observed in the first half of the twentieth century. Also, during the second half of the past century the increase in per hectare crop productivity was due to improving the yield potential. At that point, it was discovered that semi-dwarf mutants of wheat produced much more grain than their taller relatives. Selective breeding for this and other important traits over the past half century has led to steady annual increases in grain production, the Green Revolution. Nevertheless, food security is still facing additional major challenges. About 80% of the earth freshwater is currently consumed by irrigated agriculture, and this level of consumption will not be sustainable into the future. The predicted population growth will require that more of the available water resource be used for domestic, municipal, industrial, and environmental needs. Furthermore, plants rarely grow in optimum conditions and are usually subjected to environmental biotic and abiotic stress, where it is commonly believed that different kinds of abiotic stress are the main source of yield reduction (Reynolds and Tuberosa 2008). These estimated yield losses are 17% due to drought, 20% due to salinity, 40% due to high temperature, 15% due to low

temperature and 8% by other stress factors (Ashraf et al. 2008). The increasing environmental stress and climate change impose, thus, not only the challenge of stabilizing yields, but increasing them as well (Tester and Langridge 2010). Paradoxically, the semi-dwarfism trait that increased grain yields in the past, makes wheat in many cases more vulnerable to certain kinds of stress (Kantar et al. 2011). Therefore, the next Green Revolution must develop plant varieties that produce high yields under environmental stress in order to improve, or at least avoid, the worsening of the current scenario of food insecurity.

2.1.2. Salt-affected soils Soil properties have substantial influence on the life conditions of plants and crops, where specific relationships exist between a particular soil and the vegetation cover of that specific soil. Thus, any unfavorable soil conditions such as a high salt content or an inadequate nutrient supply will have an adverse effect on the life of the plants, sometimes seriously hindering their effective production. Salt-affected soils, in simple terms, are those that contain a high percentage of soluble salts, where one or more of these salt components adversely affect plant growth. Nearly 10% of the total land surface is affected by salt and no continent in our planet is free from it (Pessarakli and Szabolcs 2010). Salt-affected soils are divided into five groups. 1) Saline soils, containing high levels of mainly sodium chloride (NaCl) and sodium sulfate (Na2SO4) that occur mainly in arid and semiarid regions and form a major part of all the salt-affected soils of the world. 2) Sodic (alkali) soils, with high levels of sodium carbonate (Na2CO3) and sodium bicarbonate (NaHCO3) extended in practically all the climatic regions from the humid tropics to beyond the polar circles, and their total salt content is usually lower than that of saline soils. 3) Gypsiferous soils, with high levels of calcium sulfate (CaSO4) mainly found in the arid and semiarid regions of North America, North Africa, the Near East, Middle East, and Far East, and also in Australia. 4) High magnesium content soils, heavy texture soils occurring in arid, semiarid, and even semi-humid regions. 5) Acid-sulfate soils, with salt contents composed mainly of aluminum sulfate Al2(SO4)3 and ferric sulfate Fe2(SO 4)3 (Pessarakli and Szabolcs 2010 and references therein). The aforementioned five types of salt-affected soils developed due to natural soil-forming processes and are called primary salt-affected soils (primary salinity). All 10

these different groups have diverse physicochemical and biological properties besides having high electrolyte content. Although some agronomic and engineering practices ma y address this form of salinity, genetic improvement of crops provides the ma i n way to increase productivity on these lands. Another group of salt-affected soils exists; the secondary salt-affected soils (secondary salinity). These soils have been salinized due to ma n -made factors, mainly as a consequence of improper methods of irrigation and drainage causing water tables to rise and concentrate the salts in the root zone. Also, other anthropogenic factors such as overgrazing, deforestation, soil chemical contamination and accumulation of airborne or waterborne salts contribute to the formation of secondary salt-affected soils (Pessarakli and Szabolcs 2010 and references therein). Although these soil conditions significantly reduce the yield of most crops that are glycophytes, high soil salinity conditions are a clearly defined natural ecosystem for halophytes which are adequately well-adapted to this natural flora (Flowers et al. 2010).

2.1.3. Salinity stress: definition and negative consequences As mentioned earlier, saline soils are the most common and widespread salt- affected soil type. These soils contain excessive salinity with electrical conductivity of saturation extracts values (ECe) of > 4 dS/m (US Salinity Laboratory 1954; Tanji 2002), which is equivalent to roughly 40 mM NaCl generating an osmotic potential of approximately –0.2 MPa (Munns and Tester 2008). The measurement of soil salinity is challenging because of the strong influence of soil moisture content, among many other factors. Moreover, the soil salinity criterion is a relative one because there are substantial differences in salt tolerance among plants as will be discussed later on in this chapter (see Tanji 2002 and FAO webpage http://www.fao.org/docrep/005/y4263e/y4263e0e.htm/). In all cases, the harmful effects of saline soils on most plants are brought about generally by a combination of two kinds of stress: an osmotic stress and a Na+ imbalance and toxicity stress.

2.1.3.1. Osmotic stress The aforementioned value of osmotic potential generated by saline soils is calculated from the concentration of the salt solution by the equation (derived from

Van’t Hoff’s law) Ψs = –CRT, where Ψs is the osmotic potential (expressed in MPa), C

is the osmolality or salt concentration of the solution (moles of dissolved solute per liter of water), R is the universal gas constant (8.32 J mol–1 K–1), T is the absolute temperature (in degrees Kelvin) and the minus sign indicates that the dissolved salts reduce the water potential of a solution relative to the reference state of pure water (Sperelakis 2001, Taiz and Zeiger 2002). This high concentrations of NaCl lowers the soil osmotic potential affecting the general water balance of plants. At the cellular level, o exposure of plant cells to the low water potential of a saline environment (Ψ w) results i in the equilibration of the internal cell water potential (Ψ w) by cell-water loss and an i accompanying decrease of the cell osmotic potential (Ψ π) and turgor (Ψp) according to o i i the equation Ψ w = Ψ w = Ψ π + Ψp (Salisbury and Ross 1992). The water loss is also inevitably accompanied by a decrease of cell volume. Consequently, the overall effect of water loss and turgor decrease, negatively affects cell expansion and, hence, growth ceases. At the whole plant level, the general water balance of plants is affected because leaves need to develop an even lower water potential to maintain a “downhill” gradient of water potential between the soil and the leaves (Taiz and Zeiger 2002). Stomatal closure together with leaf growth inhibition is among the earliest responses to water deficit, protecting the plants from extensive water loss (Cornic 2000). The detailed mechanisms of stomatal response to osmotic stress are complex and are mediated by chemical signals involving mainly abscisic acid hormone (ABA) and SLAC1 anion channel that mediates ABA-induced stomatal closure (Brandt et al. 2012). The stomatal closure reduces the stomatal conductance of CO2 limiting the rate of photosynthesis, which together with the slower formation of photosynthetic leaf area reduces the flow of assimilates to the meristematic and growing tissues of the plant including both leaves and roots (Chaves et al. 2009). Within hours, however, cells regain their original volume and turgor owing to osmotic adjustment as will be discussed later (Passioura and Munns 2000). For a moderate salinity stress, an inhibition of lateral shoot development becomes apparent over weeks, and over months there are effects on reproductive development, such as early flowering or a reduced number of flowers. During this time, a number of older leaves may die, however, production of younger leaves continues. In all cases, the decreased rate of leaf growth after an increase in soil salinity is primarily due to the osmotic effect of the salt around the roots and not due to NaCl toxicity in the plant. This was confirmed using other single salts such as KCl, and

non-ionic solutes such as mannitol or polyethylene glycol (PEG) which had similar negative effects as NaCl on leaf expansion (Yeo et al. 1991).

2.1.3.2. Sodium imbalance and toxicity stress Na+ and K+ are similar in their physicochemical properties such as ionic radius and ion hydration energy and, thus, Na+ inevitably competes with K+ for major binding sites in key metabolic processes in the cytoplasm, such as enzymatic reactions, protein synthesis and ribosomal functions. Concentrations higher than 0.4 M inhibit most enzymes because of disturbances to the hydrophobic–electrostatic balance that is necessary to maintain the protein structure (Wyn Jones and Pollard, 1983). Other in vitro studies showed that Na+ inhibit some sensitive enzymes at concentrations lower than 100 mM (Flowers and Dalmond 1992). Also, given the fact that over 50 cytoplasmic enzymes are activated by K+, the disruption of metabolism is severe in all plant organs due to Na+ toxicity (Marschner 2012). The presence of Na+ in the soil affects the ability of the plant to acquire and metabolize other essential nutrients. For example, it reduces three-fold the activity of Ca2+ making it less available for plants. Also, it directly competes at uptake sites with many essential cations such as K+, Mg2+ + + or NH4 . In the case of K for instance, it may affect its influx via both low affinity nonselective cation channels (NSCC) and high affinity K+ transporters (HAK). In addition, when positively charged Na+ crosses the plasma membrane of root cells, a major membrane depolarization occurs, which in turn leads into two important negative effects. First it makes passive uptake of many essential cations thermodynamically impossible, and it increases efflux of some essential cations at the same time, such as K+, which leak through depolarization-activated outward-rectifying K+ channels (KOR). On the other hand, plant cells synthesize various compatible solutes used for osmoprotection under high Na+ concentrations as will be mentioned later on in this chapter. This process is metabolically ‘expensive’ as it reduces the available ATP pool and consequently reducing H+-ATPase activity. This not only makes high-affinity cation influx even more difficult to achieve, but also significantly reduces uptake of − −3 some important anions such as NO3 and PO4 , as their transport is energized by the H+-ATPase activity (Shabala and Cuin 2008).

2.1.3.3. Other effects of salinity stress Stomatal closure and accumulation of high levels of toxic Na+ in the cytosol under saline conditions impair the photosynthetic machinery and reduce plant’s capacity to fully utilize light absorbed by photosynthetic pigments. This leads to the formation of chemically reactive oxygen species (ROS) in green tissues having a toxic effect on biological structures (oxidative stress). ROS cause lipid peroxidation in cellular membranes, DNA damage, protein denaturation, carbohydrate oxidation, pigment breakdown and an impairment of enzymatic activity not only in leaves, but in roots as well (Mittler 2002, Apel and Hirt 2004, Miller et al. 2008). ROS directly activate Ca2+- permeable plasma membrane channels triggering a rapid Ca2+ uptake which increases cytosolic Ca2+ concentrations activating NADPH oxidase and causes additional increase 2+ in (Ca )cyt via positive feedback mechanisms (Lecourieux et al. 2002, Demidchik et al. 2003). Also, ROS activate a certain class of K+-permeable NSCC channels resulting in a massive K+ leak from the cytosol (Shabala et al. 2006). This constant elevation in cytosolic Ca2+ together with the depletion of the K+ pool activate caspase-like proteases and trigger programmed cell death (PCD) (Shabala 2009).

2.1.4. Sodium transport inside the plant Ion transport across cellular membranes depends on metabolically coupled proton pumps that generate an electrical and pH potential gradient across these cellular membranes, the proton motive force. This is accomplished by two major electrogenic proton pumps: 1) H+-ATPase (present at both the plasma membrane and tonoplast), and

2) PPi-ase (present in tonoplast only), that hydrolyze ATP and inorganic pyrophosphate

(PPi), respectively. This proton motive force drives the secondary transport processes of many ions against their electrochemical gradient. These secondary active transport mechanisms include: 1) antiport, a coupled transport in which the downhill movement of protons drives the uphill transport of a cation in the opposite direction; 2) symport, influx of a cation alongside the inward movement of H+. On the other hand, ion channels are integral membrane proteins facilitating ion movement along the electrochemical gradient (i.e. passive transport). These channels do not stay open for long periods, but have "gates" that open and close the channel pore in response to external signals, such as voltage, pH, cytosolic calcium etc. Many channels show a

steep rectification, e.g. are capable of conducting ions only in a certain range of membrane potentials and/or in a particular direction. Inward-rectifying channels mediate current flow into the cell, and outward-rectifiers - out of the cell. Ion currents through such channels may be either time-dependent (e.g. having current that increases over a period of several hundred milliseconds) upon a change in the membrane voltage, or time-independent. Outward-rectifiers are opened upon membrane depolarization, resulting in a loss of ions from the cell, while inward-rectifiers are activated by hyperpolarization of the membrane potential resulting in the movement of ions into the cell. Within a plant, the uptake of ions from the growth medium and their transport within the plant utilizes both secondary transport processes (antiporters or symporters) and channels, as will be discussed below. Na+ transport processes in plants have been extensively reviewed in the past few years (Tester and Davenport 2003, Horie and Schroeder 2004, Pardo et al. 2006, Rodríguez-Navarro and Rubio 2006, Apse and Blumwald 2007, Plett and Møller 2010, Zhang et al. 2010, Kronzucker and Britto 2011), and a detailed explanation of these processes is beyond the scope of this introduction. However, a short summary will be made covering the most important aspects of Na+ transport based on the aforementioned reviews, whereas chloride (Cl-) transport can be reviewed in Teakle and Tyerman (2010).

2.1.4.1. Sodium influx into the plant Like all other cations, Na+ enters into the plant through the epidermal cells of roots. In a similar way to K+, Na+ shows two modes of uptake: a rapidly saturating system that shows high affinity transport for Na+ (discussed later on in this chapter), and a low affinity transport system (Epstein 1966). Plasma membrane of plant cells is relatively impermeable to ions and, thus, Na+ influx from the soil and its subsequent transport within the plant is accomplished, as described above, via channels and transporters embedded in the lipid bilayer of the plasma membrane. In saline soils, Na+ enters the cytoplasm of root cells passively favored by both the concentration gradient and voltage differential across the plasma membrane (Cheeseman 1982). Also, Na+ influx appears to be mediated by Ca2+-sensitive and insensitive processes, as it was found that the presence of up to 10 mM Ca2+ in the soil (solution) inhibits the majority

of unidirectional Na+ influx (Cramer 2002). Until this very moment, the exact identity of the genes encoding the main site(s) of Na+ entry in roots is still uncertain. Regarding Ca2+-sensitive Na+ influx, a consensus has developed on the implication of non selective cation channels (NSCCs) in catalyzing Na+ influx across the plasma membranes of plant cells. Many candidates of NSCCs, especially those in the voltage insensitive NSCCs category, appear to be involved in Na+ influx across the plasma membrane (see evidence critically reviewed by Kronzucker and Britto 2011). These include cyclic nucleotide-gated NSCCs (CNGCs), amino-acid-gated NSCCs (AAG-NSCCs), and reactive-oxygen-species-activated NSCCs (ROS-NSCCs) (Davenport 2002, Demidchik and Maathuis 2007, Kaplan et al. 2007). In all cases, more studies are still needed to support that NSCCs are the main pathways of Na+ influx across the plasma membrane. Another candidate for mediating Na+ influx is the low affinity cation transporter (LCT1) identified from wheat (Schachtman et al. 1997, Amtmann et al. 2001).TaLCT1 has been shown to transport Na+, among other cations, when expressed heterologously in yeast cells. LCT1 of wheat is also sensitive to Ca2+ in a similar way to NSCCs. On the other hand, no sequences of significant identity to TaLCT1 exist in Arabidopsis or rice and thus, LCT1 may be unique to wheat. NSCCs and LCT1 are apparently doubtful to be the major pathways of Na+ influx, since in most soils calcium levels are high enough to inhibit Na+ uptake through NSCCs and LCT1 considerably. Thus, several other transporters and channels might potentially mediate this influx. In this regard, members of the HKT2 subfamily have been clearly shown to be involved in primary Na+ uptake in rice roots (Horie et al. 2007). However, the only HKT gene present in Arabidopsis (AtHKT1;1) localizes in the stele and not in the root tissues, and it was recently shown to be involved in Na+ retrieval from the xylem (Davenport et al. 2007) and not Na+ influx, as was originally proposed by Rus et al. (2001). In contrast to the single HKT gene involved in Na+ transport present in Arabidopsis and possibly many other species like Physcomitrella (Haro et al. 2010), rice has nine HKT genes (Platten et al. 2006). The presence of a numerous number of HKTs could also be the case in other cereals and monocotyledonous plants (Haro et al. 2010), and could be implicated in distinct Na+ transport activities in different plant tissues (table 1 and 2 in Zhang et al. 2010). The functions of HKT-type transporters are thus very complex and much more work is required to properly elucidate the functions of this family. 16

On the other hand, the implication of s o me K+ transporters families such as KT ⁄HAK⁄KUP and AKT families in Na+ uptake has been considered in early reviews (Blumwald et al. 2000). However, many of the results concerning these K+ transporter families seem to be contradictory in many cases (Kronzucker and Britto 2011 and references therein). Further investigation is needed to elucidate certain aspects of Na+ influx such as the possible implication of the unknown third system of K+ influx (Alemán et al. 2011) in Na+ uptake. Also, the possibility that Na+ uptake occurs through specific Na+ influx transporters in conditions of salinity is still poorly studied and needs to be thoroughly investigated (Benito et al. 2012). The varying number of HAK and HKT genes among species also raises many questions regarding the shared functions that they may play in plants regarding Na+ transport in general and influx in particular.

2.1.4.2. Sodium efflux The ideal situation for salt-sensitive plants growing in a saline soil is to keep Na+ out of the cells by Na+ exclusion (efflux). This would prevent the problems arising from Na+ getting into the cytoplasm or into the xylem stream discussed earlier. The electrochemical gradients that make Na+ influx into the root passive, make the efflux of Na+ from the cell an active process. In higher plants, evidence implies that efflux of Na+ depends only on an electroneutral secondary active transport process at the plasma membrane mediated by Na+/H+ antiporters(s) of stoichiometry 1:1 costing an ATP for each Na+ ion extruded. This is mediated in higher (flowering) plants, at least partially, by SOS1 Na+/H+ antiporter (Shi et al. 2000) which forms part of a complex pathway and activation mechanism (Mahajan et al. 2008, Quintero et al. 2011). Curiously, despite the numerous studies on SOS1 in the past few years, its function is still ambiguous, and evidence supports its contribution to many other direct and indirect functions inside the plant beside Na+ efflux (Kronzucker and Britto 2011 and references therein). Uncharacterized Na+/H+ antiporters implicated in Na+ efflux are yet to be uncovered, probably in less studied transporters such as the CHX transporter family, among others. Non vascular plants, on the other hand, posses P-type Na+-ATPases homologous to those present in many fungi, which are absent in higher plants (Garciadeblás et al. 2001, Benito and Rodríguez-Navarro 2003). For example, the moss Physcomitrella patens has three ENA Na+-ATPases (Fraile-Escanciano et al. 2009), where PpENA1 17

and PpSOS1 are complementary systems of Na+ efflux. PpENA1 is relevant exclusively at high pH values, where PpSOS1 is inactive, but is unnecessary at low pH values where PpSOS1 is active (Fraile-Escanciano et al. 2010). The absence of this system in vascular land plants indicates the occurrence of an evolutionary loss of this system after plants colonized land (Garciadeblás et al. 2001, Fraile-Escanciano et al. 2010). The increasing body of knowledge regarding salinity tolerance built over the past years suggests that an ideal salt tolerant plant should posses a serial of traits related to the minimization of Na+ toxicity in it. Na+ influx processes need be minimized in the epidermal and outer cortical cells. However, to attain this urgent goal as discussed earlier, the exact gene identities involved in Na+ influx must be first discovered. In addition, Na+ efflux should be maximized specifically in the outer root cells to improve Na+ extrusion. On one hand, this could be possibly achieved by overexpressing Na+/H+ antiporters such as SOS1. Although constitutive overexpression of SOS1 is generally successful in enhancing salt tolerance (Shi et al. 2003, Yang et al. 2009) apparently by maintaining a higher K+/Na+ ratio (Yue et al. 2012), a cell type-specific overexpression of SOS1 targeted towards the plasma membrane of root epidermal and cortical tissues is needed and is expected to be successful. On the other hand, maximizing Na+ efflux can also be attained by overexpression of Na+ ATPases such as ENA1 (Benito and Rodríguez-Navarro 2003). This was recently studied by overexpressing PpENA1 of the moss P. patens in rice, which led to more salt tolerance and produced greater biomass than controls (Jacobs et al. 2011). Curiously, although they observed that the protein localizes to plasma membrane in onion cells, as previously observed in P. patens protoplasts (Fraile-Escanciano et al. 2009), the enhanced salinity tolerance of the transgenic rice overexpressing PpENA1 was found not to be due to an increase in Na+ exclusion. In all cases, future studies are needed to untangle these apparent contradictions.

2.1.4.3. Radial transport of sodium across the root If the difference between unidirectional Na+ influx and efflux (i.e. the net influx) is not directed into the vacuoles of the epidermal and cortical root cells (as will be discussed later), Na+ is transported across the root via symplastic and apoplastic pathways (see below) from the epidermis until reaching the xylem. Symplastic transport of Na+ occurs between the cytoplasm of root cells via plasmodesmata. Also, Na+ is 18

transported transcellularly across the plasma membrane of adjacent root cells via transporters and channels, although the identity of these is still unknown (Plett and Møller et al. 2010). In any case, the mechanism of control of cell type specific patterns of Na+ accumulation is yet to be elucidated, given the dramatic differences in Na+ cellular concentrations of different cell types across the roots (figure 4 in Munns and Tester 2008). Na+ is also transported radially through the apoplast of the root cells until reaching the endodermis, where it stops due to the casparian strip. The Casparian strip of the endodermis forms a suberized barrier to apoplastic movement of water and many solutes. At this point, any further radial movement of Na+ towards the xylem will be restricted to the symplastic route through the plasma membranes of these cells. In many plants, however, interruptions in the endodermis lead to an unimpeded flow of Na+ into the xylem stream via the cell wall in a process called apoplastic bypass (Yeo et al. 1987, Faiyue et al. 2010). This indicates the presence of leaks in the endodermis or that the endodermis is permeable to some extent. Curiously, although apoplastic bypass flow in rice is relatively high, it is minimal in other species like Arabidopsis (Essah et al. 2003). The localization of certain plasma membrane transporters in the root endodermis, such as AtCHX21, might imply that they play a role in Na+ transport from the endodermal cells to the stele. Although the AtCHX21 mechanism of action in these suberized cells is mysterious, evidence in Atchx21 mutants confirm its possible role showing a significantly lower concentration of Na+ in the xylem and leaf sap compared with the wild type plants (Hall et al. 2006). Knowing this, an important salinity tolerance strategy would be to decrease symplastic Na+ transport in the root towards the xylem. Hence, minimizing the expression or even knocking-out the gene of AtCHX21 and other transporters involved in the symplastic Na+ transport might decrease the Na+ accumulation in the shoots and probably contributing to salinity tolerance.

2.1.4.4. Vacuolar sequestration of sodium Another useful trait of crucial importance in Na+ accumulation within the plant, is its partitioning within the cell. To achieve this, Na+ is sequestered in vacuoles of all plant tissue cells in order to avoid accumulation of levels of Na+ that are toxic to proteins in the cytoplasm. A considerable body of literature highlights the importance of vacuolar sequestration by experiments in which constitutive overexpression of vacuolar 19

transporters was claimed to greatly increase salinity tolerance in several species. Until very recently this function was attributed to tonoplast localized NHX-like antiporters that were thought implicated in sequestering Na + into vacuoles to avert ion toxicity in the cytosol of plants under salinity stress, where the necessary H+ gradient is maintained by both vacuolar H+-ATPase and H+-pyrophosphatase (Gaxiola et al. 2001). Salinity tolerance enhancement by overexpressing NHX1 was reported for Arabidopsis thaliana (Apse et al. 1999), tomato (Zhang and Blumwald 2001), Brassica napus (Zhang et al. 2001), wheat (Xue et al. 2004), maize (Yin et al. 2004), tobacco (Wu et al. 2004), cotton (He et al. 2005) and upland rice (Chen et al. 2007a). On the other hand, subsequent works failed to notice any enhancement of salinity tolerance in transgenic Arabidopsis overexpressing NHX1 compared to controls (Yang et al. 2009). This was also the case with transgenic tomato overexpressing AtNHX1which showed larger K+ vacuolar pools but no consistent enhancement of Na+ accumulation under salt stress. (Leidi et al. 2010). Indeed, the model of Na+ sequestration into vacuoles by NHX1 was shown later on to be incorrect, giving rise to a new model in Arabidopsis stating that NHX1 together with its isoform NHX2 are essential for active K+ accumulation at the tonoplast, for turgor regulation, and for stomatal function rather than mainly sequestering Na+ in the vacuole (Barragán et al. 2012). The discovery of the exact identity of key transporters responsible for the specific sequestration of Na+ in vacuoles and their subsequent overexpression together with vacuolar pyrophosphatase genes (Gaxiola et al. 2001, Brini et al. 2007) in both root and shoot cells is expected to be a reasonable strategy for enhancing salinity tolerance in plants.

2.1.4.5. Xylem loading Once Na+ reaches the stelar root cells, it is released (loaded) into the xylem. Some authors suggest that Na+ loading could be a passive process at high external Na+ concentrations based on reports that describe stelar cytosolic Na+ levels of 100 mM and xylem Na+ contents of 2 mM (Harvey 1985, Munns 1985, Shi et al. 2002). However, evidence from recent reports strongly suggests that it is improbable that Na+ loads passively (leaks) into the xylem. The xylem content of Na+ ranges between 10 and 30 mM (Shabala et al. 2010) and xylem parenchyma cells are usually hyperpolarized having membrane potential values of around −120 to −140 mV that become depolarized by −40 to −60 mV, under salinity conditions (Wegner et al. 2011). Thus, plants must 20

have more than 300 mM Na+ in the parenchyma’s cell cytosol to get Na+ loaded into the xylem passively, which is unlikely to occur. In all cases, at least under mild saline conditions, active Na+ loading is the most probable mechanism to occur in plants. The most likely candidate for this active loading is the Na+/H+ antiporter SOS1,which is preferentially expressed at the xylem symplast boundary of roots (Shi et al. 2002). Interestingly, being an active process, scientists investigated the reason behind the paradox of why plants apparently "waste" energy to pump Na+ into the xylem and then try to minimize its accumulation in the shoot at same time. As will be discussed later on this chapter, Na+ can be used as a “cheap” osmoticum to maintain cell turgor, supposing it will be efficiently sequestered in the cell vacuoles by the tonoplast Na+/H+ exchangers and that the loaded Na+ will be accompanied by a parallel loading of sufficient amounts of K+ into the xylem to maintain the optimal xylem K+/Na+ ratio. Although this process is apparently necessary in certain cases, however, under high salinity conditions excessive concentrations of Na+ will inevitably be loaded into the xylem.

2.1.4.6. Sodium retrieval from the xylem and removal from the shoot The Na+ loaded to the xylem will be transported to the shoots via transpiration. In order to reduce Na+ accumulation in the shoots, to prevent damaging the photosynthetic machinery, Na+ must be retrieved from the xylem before reaching the shoots. This retrieval occurs along the plant at root and shoot vasculature, and at the base of the shoot. The retrieved Na+ would primarily be sequestered in root vacuoles, especially in the pericycle and xylem parenchyma cells (figure 4 in Munns and Tester 2008). However, as in the case of Na+ loading, it is uncertain whether the process of Na+ removal from the xylem is a thermodynamically active or passive process depending on the growth and salinity conditions surrounding the plant. Several candidates have been proposed to carry out this process like inwardly rectifying channels (Wegner and Raschke 1994). Nevertheless, it is currently widely accepted that AtHKT1;1, located in the plasma membrane of the xylem parenchyma (Sunarpi et al. 2005), has a clear role in controlling retrieval of Na+ from the xylem, at least in the case of Arabidopsis (Davenport et al. 2007). Also, some HKTs in other species like rice (Ren et al. 2005) and wheat (James et al. 2006) seem to be implicated in this process as well. When salinity in the soil is high, Na+ will inevitably reach the leaves, where it is often removed and recirculated to roots through the phloem. However, some authors 21

think that the amounts of recirculated Na+ are negligible (Tester and Davenport 2003). Nevertheless, among leaves, it is widely documented that the highest concentrations of Na+ accumulate in the oldest leaves in both monocot and dicot species, that will be later shed (Yeo and flowers 1982). Also, Na+ is preferably accumulated in the leaf epidermal cells, to be as far as possible from bundle sheath cells that are the most photosynthetically active cells in the plant (Karley et al. 2000). The genetic basis that control these Na+ partitioning processes in the leaves must be investigated in future studies.

2.1.5. Can sodium be beneficial for plants? One of the most noteworthy features of Na+ is the remarkable difference among species in its importance. Na+ is an essential element for animals (including humans) and must be present in relatively large amounts in the diet. It plays an important role in maintaining the ionic balance of body tissues and fluids; its osmotic characteristics are utilized in the blood stream for regulating osmotic pressure within the cells and body fluids, where it protects against excessive loss of water (Harrison 1991). In contrast to animals, high Na+ levels is detrimental to plants as mentioned earlier. Although Na+ and K+ are physicochemically and structurally similar monovalent cations, plants exhibit a strong preference for K+ as will be discussed in a subsequent chapter. However, numerous studies provide evidence that Na+ is beneficial and in certain cases essential for plants. For example, Na+ appears to play a critical role in chlorophyll synthesis and the regeneration of phosphoenolpyruvate (PEP) in mesophyll chloroplasts of some C4 plants (Ando and Oguchi 1990, Murata et al. 1992). Also, K+ seems to be partially replaceable with Na+ in stomatal function (Marschner 1971), while under limited K+ supply, Na+ can replace K+ in the vacuole as an alternative inorganic osmoticum (Flowers and Lauchli 1983, Rodríguez-Navarro 2000). The evidence on the importance of Na+ for plants under certain conditions, may be strengthened by the presence of high affinity Na+ uptake in plants; plants can take up Na+ from external concentrations as low as those at which K+ can be taken up (Rains and Epstein 1967, Rodríguez-Navarro and Rubio 2006). Recently, this type of transport was confirmed to be present in many plant species including several flowering plants and bryophytes (Haro et al. 2010). Na+ transport from low external concentrations displays saturable Michaelis–Menten properties with low Km values, characteristic of 22

high affinity systems, while at the same time showing pharmacological sensitivities typical of ion channels, even at external [Na+] of only 10 µM (Schulze et al. 2012). Studies have proved that OsHKT1 is the micromolar Na+ uptake system of rice (Garciadeblás et al. 2003, Horie et al. 2007) while TaHKT1 and HvHKT1 mediate high affinity Na+ uptake in wheat and barley respectively (Haro et al. 2005). However, it seems that the HKT-dependent high affinity Na+ transport does not apply to all species (see critical discussion in Rodríguez-Navarro and Rubio 2006, Kronzucker and Britto 2011). In fact, the HKT transporter of Arabidopsis does not mediate Na+ influx in roots as stated earlier (Berthomieu et al. 2003, Essah et al. 2003, Davenport et al. 2007). Also, it was demonstrated that high affinity Na+ uptake in P. patens is not mediated by PpHKT1 because Pphkt1 mutant plants maintained normal Na+ (and K+) influx (Haro et al. 2010). Instead, a recent study shows that high affinity Na+ uptake in P. patens seems to be dependent on PpHAK13 (Benito et al. 2012). This latter result opens up a new line of research involving HAK transporters in high affinity Na+ uptake in plants. In all cases, the physiological basis and significance of a high affinity Na+ transport system in higher plants is still poorly understood. Some fundamental questions about the system’s characteristics remain unanswered such as the ecological significance of this system that functions primarily under unrealistic conditions such as the near absence of major soil + 2+ + nutrients like K , Ca , and NH4 , that usually don’t occur in soils (Schulze et al. 2012).

2.1.6. Salinity tolerance mechanisms in plants Plant responses to salt stress are complex, extremely variable, mutually linked, and include a wide range of effects at the molecular, cellular, tissue and whole-plant level. Furthermore, many problems with the methodology of studying salinity stress are common and widespread among different research groups leading to erroneous results and inappropriate conclusions. For example, discrepancies in responses to salinity were found between barley plants grown in soil and hydroponic systems (Tavakkoli et al. 2010). Other important aspects of salinity conditions to be taken in consideration in salinity tolerance experiments include the types of salts to be used, at which stage of plant development to start the salinity treatment and at what concentration. Experiments should also be conducted over a sufficiently long period, as plant responses in the short term are primarily due to osmotic stress, while the toxic concentrations in leaves and salt-specific effects are seen in longer term. Recently, a group of renowned researchers 23

from different groups published a joint review discussing in depth the different aspects and considerations regarding growing plants for experimental purposes (Poorter et al. 2012 and references therein). All in all, there is an urgent need to associate traits for salinity tolerance to the existing “real-life” salinity scenarios, as many spectacular results obtained in one stress scenario may have a limited interest for improving food security in other geographical areas affected by that stress (Tardieu 2012). It is therefore unlikely in the near future, to have simple answers and solutions related to the problem of salinity tolerance. Further knowledge on salt-inducible genes, genetic control of salinity responses and signaling pathways offers a chance for creating a clearer picture of plant responses. There is hope that many aspects of salinity stress would be resolved, particularly through the knowledge from molecular biology, biotechnology and bioinformatics. Salinity tolerance in plants is a complex polygenic trait difficult to explain in simple biochemical or biological processes. It is predominately under control of additive effects with large environment x genotype interaction effects where an evident trade-off between yield and salinity tolerance is observed. Salinity tolerance genes in glycophytes fall under several categories and traits including plant morphology, ions compartmentalization and fluxes, production of compatible solutes, regulation of transpiration, control of ion movement and cytoplasmic tolerance (Flowers 2004). Nevertheless, in simple terms, salinity tolerance is generally assessed as “the percent of biomass production in saline versus control conditions over a prolonged period of time” (Munns 2002). Species vary greatly in their capacity to tolerate salinity. For example, among food crops, barley, cotton and sugar beet are the most tolerant; bread wheat is moderately tolerant, while rice and most legume species are sensitive. A broad survey of the variation in salt tolerance between crop, pasture forage and horticultural species was given by Maas and Hoffman (1977). Most land plants and especially crops are glycophytes and a small minority are halophytes (i.e. salt-loving plants). These halophytes usually are native to saline areas and require some salt to grow well; some need 10–50 mM NaCl to reach maximum growth, others like Atriplex nummularia grow best at around 200 mM NaCl, while many other halophytes can grow at salinities near those of seawater or even higher. Most halophytes accumulate high concentrations of NaCl in their tissues, which in itself contributes substantially to the dry weight of the plant. The salt concentration in the leaf tissues of halophytes can be greater than 500 24

mM, which is about the maximum concentration found in other glycophyte species. As mentioned earlier, enzymes cannot function in such a high NaCl concentration, and thus, special cellular features that allow metabolism of halophytes to continue exist. Nevertheless, we will be focusing here only on the salinity tolerance mechanisms of glycophytes, and those specific to halophytes can be reviewed elsewhere (Flowers and Colmer 2008). Based on the harmful effects of the salinity stress components discussed earlier in this chapter, plants possess several physiological mechanisms that confer salinity tolerance. Regarding the osmotic stress component imposed by salinity, to maintain a normal growth, plants must therefore readjust themselves to the increased external osmolality (a mechanism also known as “osmotic adjustment”). The strategy of this mechanism mainly includes the accumulation of a variety of molecules in the cytoplasm to counteract the external osmotic pressure. This is achieved by the accumulation of organic osmolytes (compatible solutes) either by their direct uptake from the external media or by de novo synthesis inside the plant (Munns and Tester 2008). These are non- toxic (thus termed “compatible”) small water-soluble molecules that may be accumulated in cells at high concentrations without affecting metabolic reactions inside the cell. Their accumulation in the cells result in an increase in cellular osmolarity leading to the influx of water into, or at least reduced efflux from cells, thus, providing the turgor necessary for cell expansion. Compatible solutes fall under four classes: sugars, polyols, amino acids and quaternary ammonium compounds (Delauney and Verma 1993). Although the ideal strategy for plants is to directly uptake these osmolytes from the soil, the concentration of these organic osmolytes in soils is extremely low ,and thus, this option is not practically viable. On the other hand, the de novo synthesis of these osmolytes, although more common, is not a very efficient strategy either. The synthesis of these solutes, besides being a very slow process and the low concentrations synthesized (usually do not exceed several mM) do not produce a significant osmotic adjustment of whole cells or tissues, the synthesis of these molecules comes at extremely high metabolic cost; usually 30-80 moles of ATP are used to synthesize one mole of compatible solutes (Shabala and Shabala 2011). Therefore, a feasible alternative is osmotic adjustment by means of inorganic ions like K+, Na+ and Cl−. K+ is the most suitable and routinely used ion for osmotic adjustment as its accumulation does not interfere with the cell metabolism. However, as described 25

earlier in this chapter, the electrochemical gradients induces K+ loss under high salinity conditions, and thus, Na+ and Cl− are used as cheap osmoticum to maintain normal cell turgor. Unfortunately, both ions are toxic at high concentrations and cause severe disruptions to cell metabolism. Thus, efficient sequestration in the vacuole is absolutely essential. Efficient Na+ sequestration in the vacuole is energetically the most favorable and efficient way to achieve osmotic adjustment under saline conditions. Pumping 1 mole of Na+ against the electrochemical gradient into the vacuole takes only 3.5 moles of ATP, which is only 1/10 of the amount required to produce 1 mole of organic osmolyte (Munns and Tester 2008),where the energetic benefits are obvious. Several tolerance mechanisms implemented by plants to overcome the ionic imbalance and toxic effects produced by high concentrations of Na+ ion have been documented in the literature. Two of these, Na+ exclusion (efflux) and sequestration have been discussed earlier in this chapter emphasizing the very essential role of plasma membrane and tonoplast Na+/H+ exchangers as a component of plant salinity tolerance mechanisms. These two processes are important in all plant tissues whether immediately after Na+ has entered the roots or after it has been unloaded from the xylem. However, in the case of Na+ sequestration in the vacuole, it is worth mentioning that efficient Na+ sequestration relies not only on transport across the tonoplast, but also on retention of ions within vacuoles, as Na+ may easily leak back given the difference in Na+ concentrations between the vacuole and cytosol, unless some efficient mechanisms are in place to prevent this process (Shabala and Mackay 2011). Also, besides Na+ sequestration in the vacuole, tissue-specific Na+ sequestration is a common salinity tolerance mechanism. As a general rule, Na+ concentrations are always lowest in the growing youngest tissue, as these are fed by the phloem which has very low Na+ concentrations. Also, when comparing fully grown leaves on a plant, Na+ concentrations are lower in the younger than older leaves because they have been transpiring for shorter periods of time. Moreover, many species can retrieve Na+ from the xylem as it flows through a leaf base towards the leaf tip or through an elongated stem towards the shoot apex. This retrieved Na+ was reported to be stored in the leaf sheath of some monocotyledonous species (Wolf et al. 1990, Davenport et al. 2005). It was empirically observed that a wide range of plant species that tolerate moderately saline environments, have a greater ability to exclude Na+ from the shoot and/or leaf blades, and thus maintain high levels of K+. Several studies documenting 26

this observation were published for varieties of Hordeum (Greenway 1962, Garthwaite et al. 2005), rye and triticale (Gorham 1990), durum wheat (Munns et al. 2000), and rice (Zhu et al. 2001). As a result, this led to the formation of a paradigm on salt tolerance in plants, stating the existence of a negative correlation between salinity tolerance and Na+ accumulation in leaves; the lower Na+ concentration in the shoots, the more tolerant the plant is to salinity. However, this paradigm seems to be an inadequate generalization more than a rule as it doesn't stand up to the numerous exceptions in other species. For example, in certain subspecies of tetraploid wheat (Triticum turgidum) differences in salt tolerance do not correlate with differences in Na+ exclusion and a lower content of Na+ in shoots (Munns and James 2003). This was also the case in a study made on bread wheat (Triticum aestivum) where no negative correlation was found either (Genc et al 2007). As for Arabidopsis, the relationship between Na+ exclusion and salinity tolerance also appears to be complex, and no correlation was observed (Møller and Tester 2007, Jha et al. 2010). In fact, Rus et al. (2006) research on Ts-1 and Tsu-1 Arabidopsis accessions revealed that under salinity stress, although both accessions had a higher shoot Na+ concentrations than Col-0 (control accession), both Ts-1 and Tsu-1 accessions were more tolerant to salinity stress and survived longer than Col-0. Recently, it was also shown that the overexpression of HvHKT2;1 in barley led to an increase of Na+ concentrations in the leaves but paradoxically resulting in more salt tolerance in barley (Mian et al. 2011). These results, on the other hand, do not undermine the trait of Na+ exclusion from shoots as an important trait in salinity tolerance, but only indicate that in many species, other mechanisms may be important as well. Regarding the oxidative injury caused by salinity, a number of enzymes and low molecular weight compounds capable of detoxifying ROS, without themselves undergoing conversion to a destructive radical, act as antioxidant molecules (Mittler 2002). These enzymatic and non-enzymatic components include: 1) superoxide dismutase (found in all cellular compartments); 2) water–water cycle (in chloroplasts); 3) the ascorbate–glutathione cycle (in chloroplasts, mitochondria, cytosol and apoplastic space); 4) glutathione peroxidase; and 5) catalase (both in peroxisomes). It has been widely documented that plant salinity tolerance may be achieved not only by cytosolic Na+ exclusion but also by efficient cytosolic K+ retention. The next chapter will briefly describe the essentiality of K+ to plants, its transport, as well as the

importance of K+ retention and maintaining a high cytosolic K+/Na+ ratio and homeostasis under normal and saline conditions.

2.2. POTASSIUM HOMEOSTASIS 2.2.1. Physiological functions of potassium in plants It is accepted in the scientific community that life was originated in the sea water environment where Na+ concentration is high. For unknown reasons, however, sea water living cells selected K+ over Na+ as the normal cation in the cellular milieu through the course of ti me . The earliest and primary functions of K+ were probably electrical charge balance and osmotic adjustments, where the fulfilling of these functions led to a K+ rich cellular environment, in which further physiological evolution resulted in a large number of physiological functions that became K+ dependent. With time, K+ became essential for the existence of all living organisms on earth. For example, animals require K+ to maintain the osmotic equilibrium within the body, as well as for neuronal signalling, muscle activity, heartbeat, and activation of a large number of enzymes involved in various metabolic processes. This K+ is obtained either directly from plants or indirectly through the animal products in the diet. In plants, K+ is the second most abundant mineral nutrient (after nitrogen), forming up to 10% of the plant’s dry weight (Marschner 2012). Also, K+ is the most abundant cation in plant cells’ cytosol and vacuole under normal growth conditions. This high concentration of “free” K+ in the cells of growing tissues and reproductive organs reflects again the vital role it plays from cellular to the whole plant level. However, although K+ was not considered an essential element earlier in the past century according to the strict criteria of Arnon and Stout (1939), as it is partially substituted by Na+ under some circumstances as aforementioned in the last chapter, nowadays, it is widely accepted that K+ is indeed an essential macronutrient required in relatively high concentrations in plants (Britto and Kronzucker 2008). K+ is characterized by high mobility in the plant at all levels; within individual cells, within tissues, as well as in long-distance transport via the xylem and phloem. It is not metabolized and it forms only weak complexes in which it is readily exchangeable, and thus not strongly competing for binding sites of divalent cations. Amtmann et al. (2004) classified the functions that K+ fulfills in the plant into two classes. The first class are the functions that depend on high and relatively stable concentrations of K+ in cellular compartments or tissues. These include the activation of over 50 key metabolic enzymes including those involved in photosynthesis, oxidative metabolism, and protein synthesis. Stabilization of protein synthesis, neutralization of 29

negative charges on proteins, and maintenance of cytoplasmic pH homeostasis optimal for most enzymatic reactions (pH ~ 7.0), fall under this class as well. The second class of functions include those that depend on K+ movement between different organelles, cells, or tissues. First and foremost, cell and plant growth requires directed movement of K+ where its accumulation in plant vacuoles creates the required osmotic potential for cell expansion. In addition, this class includes functions where K+ movement is the driving force for osmotic changes in stomatal movement, light-driven and seismonastic movements of organs, and phloem transport. Moreover, K+ movement provide a charge balancing counter-flux necessary for the transport of other ions and molecules such as − sugars, amino acids, and NO3 (Marschner 2012 and references therein). Recently, it has been attributed a new function where K+ circulation in the phloem serves as an energy source suitable for temporarily overcoming locally occurring energy limitations as in cases of reduced activity of the H+-ATPase due to downregulation or low ATP levels (Gajdanowicz et al. 2011).

2.2.2. Potassium deficiency symptoms K+ is frequently in short supply in many agricultural systems, due its high demand as a nutrient (Öborn et al. 2005), resulting in K+ deficiency. Plants experiencing mild K+ deficiency rarely show evident and visible symptoms as K+ is readily redistributed within the plant via the phloem from mature to developing tissues. (Mengel et al. 2001, Fageria 2009). However, when plants experience more severe K+ deficiency, they exhibit symptoms consistent with the vital functions of this element mentioned earlier. Morphologically, K+ deficient plants exhibit scorching along the margins of older leaves. They grow slowly, become stunted with poorly-developed root systems, and are more susceptible to abiotic and biotic stress such as drought, cold, salinity, and fungal attacks (Marschner 1995). Both leaves and roots of K+ deficient plants are short-lived. Stems are weak, and seed and fruit are small and shriveled. At the physiological level, symptoms include impaired phloem transport, particularly of sucrose, increased leaf carbohydrate concentrations, a reduction in chlorophyll concentrations, decreased water content, decreased turgor, impaired stomatal regulation and reduced transpiration (Cakmak et al. 1994, Mengel et al. 2001, Hermans et al. 2006, Amtmann et al. 2008). Moreover, a severe decline in net photosynthesis occurs (Peoples and Koch 1979, Bednarz and Oosterhuis 1999, Cakmak and Engels 1999) affecting crop 30

yield. This decline appears to be an indirect consequence of sucrose accumulation in leaves (Hermans et al. 2006) as sucrose export from leaves of K+ deficient plants becomes impaired. This impairment is due to an apparent failure of sucrose loading into the phloem due to unfulfilled K+ requirements (Mengel et al. 2001, Deeken et al. 2002, Hermans et al. 2006). At the cellular and metabolic levels, the decline of photosynthesis in K+ deficient plants leads to increased reactive oxygen species production, resulting in photooxidative damage (Cakmak 2005). Also, pH of the cytosol decreases (Walker et al. 1995) leading to a decrease in protein synthesis and a subsequent decline in growth.

2.2.3. Soil and plant concentrations of potassium K+ is the fourth most abundant mineral in the earth crust, constituting about 2.5% of the lithosphere (Sparks and Huang 1985). It is present in the soil in four different pools: (i) mineral K+; (ii) fixed K+; (iii) exchangeable K+, and (iv) soil solution K+ (Mclaren and Cameron 1996, Syers 1998). This relatively large amount does not however reflect the availability of K+ to plants as the largest fraction, mineral K+, is almost completely unavailable to plants (Pal et al. 1999). Moreover, the release of exchangeable K+ is often slower than the rate of K+ acquisition by plants (Sparks and Huang 1985) and the exchange of K+ between different pools is strongly dependent upon the concentration of other macronutrients in the soil solution (Yanai et al. 1996). For example, as mentioned in the past chapter, the presence of high levels of other + + + monovalent cations such as Na and NH4 interfere with K uptake (Spalding et al. 1999, Qi and Spalding 2004, Rus et al. 2004). Consequently, K+ content in soils is often low ranging between 0.1 to 6 mM, depending on the soil type (Adams 1971, Pretty and Stangel 1985, Johnston 2005). This represents only 0.1%–0.2% of the total soil K+, of which 1%–2% is “exchangeable” K+, 1%–10% is “non-exchangeable” K+ associated with clay lattices, and 90%–98% is present as K+ minerals (Mengel et al. 2001, Rengel and Damon 2008, Fageria 2009). Thus, taking all the above into consideration, supplying K+ fertilizer to soils is of crucial importance in crop production, being done for over 250 years. Nowadays, farmers apply roughly over 30 million tons of K+ fertilizer annually (FAO 2011). In contrast to the low K+ concentration in soils, plants strictly maintain relatively high K+ levels in its cells in the millimolar range. K+ is present in all compartments of plant cells but occurs mainly in two major pools, one in the vacuole and one in the 31

cytosol. Other pools of K+ are also present within the cell, although in much smaller quantities. Cytosolic K+ is strictly controlled at approximately 100 mM for enzyme activation and protein biosynthesis (Maathuis and Sanders 1994, Walker et al. 1996, Leigh et al. 1999, Cuin et al. 2003). Cytosolic K+ concentration may decreases however to as low as 15 mM under saline conditions (Cuin et al. 2003). On the other hand, vacuolar K+ content is less strictly regulated where large fluctuations in vacuolar content may occur mirroring the external supply of K+ (Leigh and Wyn Jones 1984, Malone et al. 1991, Walker et al. 1996, Leigh et al. 1999). For example, under K+ replete conditions, the concentration of vacuolar K+ is typically around 200-250 mM but can reach 500 mM in open stomatal guard cells (MacRobbie 1998). On the other hand, under K+ limiting conditions, the osmotic functions of K+ in vacuoles may be substituted by other cations (Na+, Ca2+ or Mg+) or organic solutes (sugars) and thus decreasing drastically to very low levels (Amtmann et al. 2006), where the lowest limit for vacuolar K+ concentration appears to be 10–20 mM (Leigh and Wyn Jones 1984). Also, vacuolar K+ levels vary depending on the organ of the plant. For example, it was reported that K+ concentrations in the leaf cell vacuoles are approximately 230 mM (Cuin et al. 2003) compared to 120 mM in the root cell vacuoles (Walker et al. 1996) contrary to root and leaf cytosols that are similar (Walker et al. 1996, Cuin et al. 2003). However this heterogeneity is slight between different cell types of the same organ (e.g. leaves), while becomes more pronounced under K+-limiting conditions, where K+ concentrations are maintained in mesophyll cells higher than in epidermal cells of the leaves (Fricke et al. 1994). K+ is also maintained at high concentrations in other metabolically active compartments of the cell such as the nucleus, the stroma of chloroplasts and the matrix of mitochondria. For example, K+ plays an important role in charge balancing in thylakoid membranes (Pottosin and Schöenknecht 1996, Hinnah and Wagner 1998), as well as in enzymatic control of leaf photochemistry in stroma. This may explain the levels of K+ (50 to 100 mM) found in the stroma of chloroplasts (Demmig and Gimmler 1983, Pier and Berkowitz 1987).

2.2.4. Potassium homeostasis and transport in plants In order to ensure normal plant growth and reproduction, K+ must be maintained at the adequate concentrations in all cell organelles and compartments. The concept of homeostasis (from the Greek hómoios meaning “similar” and stásis meaning “standing 32

still”) was introduced in the early 20th century, where it was used to describe the capacity of a living organism to control its internal conditions (temperature, pH, ions, etc.) and maintain them constant and stable (Cannon 1928). In our case, plant ion homeostasis describes the capacity of plants to control the concentration of an ion within a defined space despite the fluctuating concentrations in the environment (Amtmann and leigh 2010). Although the term “ionic” applies to Na+ as well, some authors believe that it is rather incorrect to use the term “Na+ homeostasis” due to the great range of variability in the values reported for cytosolic or vacuolar Na+ concentrations in the literature which, thus, makes the homeostatic concept of maintaining Na+ within narrow limits not applicable, advising to use the term “tissue Na+ content” instead (Kronzucker and Britto 2011). On the other hand, Amtmann and Leigh (2010) see that these contradictions may be reconciled by using “K+/Na+ homeostasis” as an alternative term, given that this ratio seems to be maintained well in some plant tissues. However, as described earlier, this is not the case for K+, and the term “K+ homeostasis” is legitimate, given that it is strictly maintained at more or less stable levels in the cell. K+ homeostasis at the cellular, tissue and whole plant level is accomplished by a battery of channels and transporters. Three gene families encoding K+ transporters have been identified in the model plant Arabidopsis thaliana: 1) KT/HAK/KUP, 2) TRK/HKT and 3) CPA, subdivided in CPA1 y CPA2 families (Mäser et al. 2001). Also, the K+ channels implicated in transport and homeostasis include: 1) shaker-type family of K+ channels; 2) “two-pore” K+ channels; 3) cyclic- nucleotide-gated channels; and 4) glutamate receptors. Both transporters and channels are located in different cellular membranes, have different properties and are differentially expressed among the different plant tissues.

2.2.4.1. Long distance potassium transport 2.2.4.1.1. Potassium influx K+ moves from the soil solution to the root surface by diffusion and enters the root symplast through the cell plasma membrane of root hair epidermal and cortical cells. Also, as in the case of Na+ described in the past chapter, K+ enters the apoplastic route until reaching the Casparian strip. This K+ uptake by roots and its subsequent accumulation by plants is determined by the K+ uptake capacity of the roots, the K+

concentration at the root surface, and the replenishment of rhizosphere K+ (Jungk and Claassen 1997). Differences in K+ uptake, however, do exist not only between plant species but between genotypes of the same species as well (Rengel and Da mo n 2008). The pioneer work of Epstein and his colleagues about 60 years ago revealed that plant roots absorb K+ over a wide range of concentrations and that at least two K+ transport mechanisms exist exhibiting biphasic kinetics in response to increasing external K+ concentrations; system I, which mediates high affinity K+ uptake at external K+ concentrations below 1 mM and system II, which operates at higher external K+ concentrations as a low affinity uptake mechanism (Epstein et al. 1963). Nevertheless, apparently it is not the “affinity” for K+ that differentiates K+ transport mechanisms in the root plasma membrane, but their coupling to pH and voltage gradients as will be exposed next (Britto and Kronzucker 2008). When external K+ concentrations is less than 0.1 mM, a K+-depleted environment, K+ influx across the plasma membrane of root cells occurs against its electrochemical gradient catalyzed by K+/H+ symporters in the plasma membrane energized by the pH and voltage gradients generated at the plasma membrane by the H+- ATPase (Maathuis and Sanders 1993). This was shown in earlier reports on other model systems such as Neurospora crassa where it was demonstrated that its high affinity K+ uptake is mediated by a K+/H+ symporter (Rodríguez-Navarro et al. 1986). In plants, HvHAK1 transporter of barley was demonstrated to be a high affinity K+ transporter by functionally complementing Saccharomyces cerevisiae yeast mutants defective in their endogenous K+ uptake systems TRK1 and TRK2 (Santa-María et al. 1997). This was also revealed to be the case in plasma membrane transporter AtHAK5 of Arabidopsis (Rubio et al. 2000, Qi et al. 2008) as well as in other HAK transporters of phylogenetically distant plant species including rice (Bañuelos et al. 2002 ), tomato (Wang et al. 2001, 2002), Cymodocea nodosa (Garciadeblás et al. 2002), Physcomitrella patens (Garciadeblás et al. 2007), and Thellungiella halophila (Alemán et al. 2009). Subsequent in planta experiments on T-DNA insertion mutants of Arabidopsis confirmed the notion that root high affinity K+ uptake was impaired in hak5 mutants (Gierth et al. 2005, Rubio et al. 2008). In fact, it has been recently established that AtHAK5 is the only system mediating K+ uptake at concentrations below 10 µM (Rubio et al. 2010). Other characteristics of high affinity uptake observed in planta are + + its upregulation by K starvation, its inhibition by NH4 , and its lack of any 34

discrimination between K+ and its analogue Rb+ (Kochian and Lucas 1988, Maathuis and Sanders 1996, Rodríguez-Navarro 2000, Rodríguez-Navarro and Rubio 2006). At soil K+ concentrations above 1 mM, common in well-fertilized agricultural soils, K+ influx to root cells is energized by the voltage gradient alone and facilitated by K+ channels. Sentenac et al. (1992) isolated Arabidopsis Shaker K+ channel AKT1 and provided evidence that it operates at millimolar K+ concentrations and is involved in low affinity transport. It localizes to the plasma membrane (Hosy et al. 2005) and is preferentially localized in the peripheral cell layers of the root mature regions (Lagarde et al. 1996). Since its discovery, other members of the Shaker family of K+ channels have been identified in Arabidopsis sharing similar properties to AKT1, although present in numerous cell types within the plant and performing different functions. These are classified into three functional groups based on their dependency on membrane voltage, (i.e. hyperpolarized or depolarized membrane potentials) (Very and Sentenac 2003). These groups are: (1) inward-rectifying channels AKTl, KAT1, KAT2 and SPIK involved in K+ influx and are activated by membrane hyperpolarization (Anderson et al. 1992, Sentenac et al. 1992, Pilot et al. 2001, Mouline et al. 2002); (2) weakly-inward-rectifying channels AKT2/3 mediate both K+ influx and efflux depending on the local K+ electrochemical gradients (Marten et al. 1999, Deeken et al. 2000, Lacombe et al. 2000, Gajdanowicz et al. 2011); and (3) outward-rectifying channels SKOR and GORK involved in K+ efflux from the cell and are activated by membrane depolarization (Gaymard et al. 1998, Ache et al. 2001, Ivashikina et al. 2001, Hosy et al. 2003). As aforementioned, the main K+ channel among these three groups that is involved in K+ influx, at least in A. thaliana, is AtAKT1. Other K+ channels with homology to AKT1 were identified in several other species as well (Hartje et al. 2000, Buschmann et al. 2000, Golldack et al. 2003, Garciadeblás et al. 2007). The currently established model of action of AtAKT1 is that the regulatory subunit AtKC1, of high homology to K+ channels and strongly expressed in roots, together with the syntaxin SYP121, regulate AtAKT1 function by forming a ternary complex that acts as a functional unit mediating K+ uptake in Arabidopsis roots (Reintanz et al. 2002, Pilot et al. 2003, Honsbein et al. 2009). Interestingly, this channel mediates K+ uptake within a very broad range of external K+ concentrations, including micromolar concentrations (i.e. high affinity range) as reported by Hirsch et al. (1998), where they found that

Arabidopsis T-DNA knock-out mutant akt1 had a reduced plant growth in the presence + + of 2 mM NH4 when external K was below 1 mM. At this moment, the accumulated body of evidence suggests that there is a certain functional overlap between the high and low affinity K+ uptake systems, rather than a strict classification of either high or low affinity K+ uptake system. The current model of the relative contributions of each of these two systems, AtHAK5 and AtAKT1, to K+ uptake from diluted solutions in Arabidopsis has mainly been established based on studies in hak5 and akt1 single and double mutants (Rubio et al. 2010, Alemán et al. 2011). As mentioned earlier, AtHAK5 is the only system mediating K+ uptake at concentrations below 10 µM. Between 10 and 50 µM K+, AtHAK5 and AtAKT1 are the only systems contributing to K+ uptake. Above 50 µM K+, both systems are active, while the contribution of AtHAK5 decreases at concentrations higher than 200 µM, probably being AtAKT1 the only operating system in this concentration. At external concentrations higher than 500 µM, AtHAK5 is not relevant and the contribution of AKT1 to K+ uptake becomes more significant. That being said, the contribution of other systems in roots to K+ uptake cannot be excluded. For example, the plasma membrane symporter AtCHX13 has been shown to be upregulated + + in K starved roots and that it mediates high affinity K uptake with a Km of 196 µM in plant cells (Zhao et al. 2008). Nevertheless, Rubio et al. (2010) apparently failed to + detect any K uptake at the aforementioned Km value in the hak5 akt1 double mutant of Arabidopsis. On the other hand, the disruption of AKT1 channel does not completely abolish low affinity K+ uptake and thus, other systems could contribute to K+ uptake. These ma y include some CNGCs like the plasma membrane AtCNGC10, which partially rescues Arabidopsis akt1 mutant (Kaplan et al. 2007), and AtCNGC3 (Gobert et al. 2006). Other unidentified members of the CNGCs or unknown systems may also contribute to K+ uptake to promote plant growth even in the absence of AtAKT1 or in the presence of 10 mM K+ where AtAKT1 is not essential. These systems in all cases remain to be discovered.

2.2.4.1.2. Potassium accumulation and release from the vacuole After K+ is transported through the cell plasma membrane, it will participate in the different metabolic reactions of cytosol, transported to other metabolically active

compartments (nucleus, mitochondria, chloroplasts and Golgi apparatus), or stored into the vacuole. Plant vacuoles, especially the central lytic vacuole (LV) that occupies over 80% of the cellular volume, play important roles in the plant at the cellular and tissue level (Leigh and Sanders 1997, Martinoia et al. 2012). Vacuoles basically act as reservoirs for minerals and water, where the most important role of the central vacuole is, thus, to increase cell size by water uptake. In addition to growth, vacuoles play important metabolic roles in the cell. They serve as storage organelles for sugars and polysaccharides, organic acids and proteins that can be retrieved from the vacuole and utilized in metabolic pathways. Also, vacuoles accumulate heavy metals, such as cadmium (Vogeli-Lange and Wagner 1990) and other toxic ions like Na+, as described in the previous chapter. Vacuoles have also a role in defense against microbial pathogens and herbivores by accumulating phenolic and alkaloid compounds as well as enzymes that break down fungal cell walls, such as chitinase (Boller and Vogeli 1984). In addition, some plant species vacuoles store water-soluble flavonoid pigments (anthocyanins) that give the characteristic flower color of petunias, and that in leaves, they protect the essential components of the photosynthetic apparatus to prevent photo- oxidation (Chaves et al. 2009). Furthermore, vacuoles contain many acid hydrolase enzymes, such as proteases and ribonucleases, that are used in degradation of both exogenous and endogenous compounds such as proteins. Another major function of plant vacuoles is their contribution to pH and ionic homeostasis. All metabolic reactions in the cytosol are sensitive to changes in pH, which is usually maintained at 7.5. The pH of the vacuole of higher plants is acidic and typically ranges between 5.0 and 6.0. However, as in the case of K+, the pH range is more strictly controlled in the cytosol than in the vacuole, where the vacuolar pH may reach extremely low pH levels like 0.6, as in the case of certain algae (McClintock et al. 1982). The pH homeostasis of the cytosol is partially regulated by pumping excess amounts of protons out of the cytosol into the lumen of the vacuole, that at the same time generate the adequate proton motive force and vacuolar membrane (tonoplast) potential (−10 to −30 mV) necessary for ion transport into and out of the vacuole. This is achieved by vacuolar proton pumps H+-ATPase and H+-pyrophosphatase (Gaxiola et al. 2001). Consequently, inward and outward ion movements are achieved by secondary transport mechanisms and channels (as previously described in section 2.1.4). In addition to pH, metabolic reactions in the cytosol are sensitive to ionic concentrations, 37

and thus, ionic homeostasis in the cytosol is strictly controlled with the help of the vacuole that acts as a valve for cytosolic excesses of ions (Martinoia et al. 2012). Ca2+ cytosolic concentrations is strictly regulated and maintained in nanomolar concentrations, while being in the millimolar range in the vacuole. This steep gradient between the vacuole and cytosol is necessary due to the importance of Ca2+ in signal transduction mechanisms in the cell, where extremely low concentration are needed to induce or inhibit several metabolic and regulatory reactions. In fact, the large number of primary and secondary active Ca2+ transport systems reflects the importance of this strict regulation (Manohar et al. 2011, Pittman 2011). This cytosol-to-vacuole strict regulation is also true for many other important ions such as K+. As mentioned earlier in this chapter, the vacuolar concentrations of K+ varies considerably depending on many reasons, and considering the vacuole as a reservoir, its K+ concentration is not strictly regulated as in the case of cytosolic K+. The transport of K+ across the vacuolar membrane is accomplished by proton-coupled antiporters and ion channels. As previously described in the past chapter, the Na+/H+ exchanger (NHX) class of transporters, originally identified as a group of Na+/H+ antiporters, was also shown to play an important role in K+ delivery into the vacuole. Isoforms NHX1 and NHX2 of Arabidopsis (functionally redundant) are essential for active K+ accumulation at the tonoplast, where it was demonstrated that nhx1 nhx2 double mutants had a reduced ability to create the vacuolar K+ pool, consequently leading to greater K+ retention in the cytosol (Barragán et al. 2012). Indeed, tomato plants overexpressing AtNHX1 accumulated more K+ in the vacuole and retained less K+ in the cytosol than non transformed plants (Leidi et al. 2010). Vacuolar channels, on the other hand, mediate K+ release to the cytosol. Among the K+ vacuolar channels, the Arabidopsis slow vacuolar (SV) channel was the first to be discovered its encoding gene (Peiter et al. 2005); AtTPC1 (two-pore channel 1). This is a ubiquitous voltage dependent nonselective cation channel (NSCC) which has been found in many different plant species and all plant tissues (Hedrich and Marten 2011). Its voltage dependence show a characteristically slow activation (hence called slow vacuolar “SV” channel) at depolarizing tonoplast potentials. Thus, its current is predominantly directed from the vacuole to the cytosol. TPC1 mediates large Ca2+-dependent K+ and Na+ currents (Pottosin and Schöenknecht 2007), and is permeable to some divalent cations like Ca2+ and Mg2+. In addition to its Ca2+ dependence, TPC1 is regulated by several mechanisms 38

including phosphorylation (Bethke and Jones 1997), organic cations and redox potential (Scholz-Starke et al. 2004). However, although it transports several ions, these ion contents were not significantly affected by TPC1 expression, indicating that the SV channel is unlikely to be particularly important in plant nutrition (Maathius 2010). Another channel involved in K+ release from the vacuole is the vacuolar K+ (VK) channel known as AtTPK1 (Gobert et al. 2007). Three other isoforms, TPK2, TPK3 and TPK5, of this channel are located in the vacuolar membrane, however not well characterized like TPK1. Similarly sized TPK families have been found in genomes of other species such as rice, tobacco and Physcomitrella (Dunkel et al. 2008). These VK channels are highly selective for K+, lack voltage dependence and show much lower requirement for cytoplasmic Ca2+ (1 µM) than the SV channel. Knockout mutants of TPK1 gene are sensitive to both high and low K+ concentrations, suggesting that this channel functions as a valve that maintains the desired K+ concentration in the cytosol (Gobert et al. 2007). A third type of channel, although its encoding gene remains unknown, is the fast vacuolar (FV) channel. It has a very low selectivity for K+ or Na+ (Bruggemann et al. 1999) where it shows moderate outward rectification (Tikhonova et al. 1997). It is worth mentioning that it is erroneous to attribute all K+ release activity from the vacuole to channels. For example, it was demonstrated that the OsHAK10 transporter of rice, closely related to AtKUP transporters, localizes to the tonoplast (Bañuelos et al. 2002), supporting the idea that HAK/KUP/KT transporters is implicated in transporting K+ through the vacuole under K+ deficiency. The most probable activity of that transporter is K+ release from the vacuole by a symport mechanism (Rodríguez- Navarro and Rubio 2006). The KUP family members have been previously characterized and it was reported that they transport K+ (Ahn et al. 2004) and, in fact, recent studies demonstrate that five members of the KUP family of Arabidopsis are expressed in the vacuolar tonoplast (Whiteman et al. 2008). This implies a role of these transporters in vacuolar transport. Also, the Arabidopsis CHX17-19 transporters (Chanroj et al. 2011) localize to prevacuolar compartments, which suggests their implication in K+ transport as well. In all cases, a rather complicating factor when analyzing the physiological roles of vacuolar transporters and channels, is the seemingly high level of functional redundancy in vacuolar cation transport, as reported in the case NHX1 and NHX2 transporters (Barragán et al. 2012). In fact, Maathius (2010) reported 39

that the disruption of either the SV or VK channel did not affect plant growth in most conditions. He also mentions that even in plants where both these channels are interrupted, no or little phenotype is observed. As will be reported in subsequent chapters of this thesis, we provide evidence for the possible implication of CHX2 transporter of Physconitrella patens in vacuolar K+ release.

2.2.4 .1.3. Potassium efflux In contrast to K+ influx, the physiology and molecular biology of K+ efflux from plant roots is poorly studied and characterized. Among the characterized channels possibly implicated in K+ efflux in Arabidopsis, is the shaker-type outward rectifying channel AtGORK. As well as being expressed in guard cells, this voltage dependant channel is also expressed in root hairs and root epidermal cells where it probably mediates K+ efflux from these cells into the root medium in response to external stimuli (Ivashikina et al. 2001). It may also perform similar functions to Tok1 channel of Saccharomyces cerevisiae in membrane potential maintenance (Maresova et al. 2006). The similar SKOR channel, expressed in stele cells (discussed later on), together with GORK channel are the only channels identified to date that are implicated in K+ efflux. However, the identity of cortical K+ efflux channels, for example, remains to be + + + + discovered. The sudden exposure of roots to cations such as Na , Rb , Cs and NH4 causes membrane depolarization (i.e. becomes less negative), which in root cells leads to a subsequent efflux of K+ to counteract this change (Nocito et al. 2002). The role of GORK in epidermal root cells is, thus, to rectify the alterations in the plasma membrane electrical potential difference (ΔΨ), brought about by the above mentioned depolarization events, by effluxing K+ out of the cell, counteracting the depolarization of the cell. This is an interesting trait useful in salinity tolerance, that will be discussed later on in this chapter. However, GORK channel is predominantly expressed in leaves, and thus, the principal role attributed to it in the literature is the rapid removal of K+ during stomatal closure (Ache et al. 2000), where gork-1 mutants showed disrupted water relations (Hosy et al. 2003). K+ efflux not only occurs under circumstantial events as described above, it is also an important process in establishing the steady state together with K+ influx. It is well documented that that K+ acquisition is accompanied by a certain degree of K+ efflux. This efflux increases progressively and approaches the rates of influx as the 40

external K+ concentrations rises, consequently resulting in an increased futile K+ cycling at the plasma membrane that may have energetically expensive consequences (Britto and Kronzucker 2006). On one hand, under these high external K+ conditions, the strategy of some organisms like yeast and other non-vascular bryophyte plants like P. patens, is to efflux K+ by pumping it out against its concentration gradient (higher outside) via Na+ (and K+) ENA ATPases (Benito et al. 2002, Benito and Rodriguez- Navarro 2003, Fraile-Escanciano et al. 2009). On the other hand, however, these ATPases are not present in higher plants, and thus, an alternative strategy probably exists. For this purpose, it was proposed in past reviews the implication of cation/proton antiporters in K+ efflux (Pardo et al. 2006). The proposed model of K+ efflux suggested the use of the chemical potential of plasma membrane proton gradient in an electroneutral K+-H+ antiport exchange mechanism (Britto and Kronzucker 2006). These authors proposed as candidates the transporters from CPA2 family (CHX and KEA transporters) due to their similarities with KefB and KefC, that mediate K+ efflux in Escherichia coli. However, the only transporter characterized to date from this family that expresses in roots and localizes to the plasma membrane, AtCHX13, seems to be a high affinity K+ influx transporter (Zhao et al. 2008), while the rest of members are endomembrane transporters and preferentially or exclusively expressed in pollen only. Recently, a study on intact barley roots using 42K+ labeling combined with several pharmacological blocking agents, provided evidence that K+ efflux at low K+ (0.1 mM) and high-K+ (1 mM) growth conditions represent two distinct phenomena, as in the case of high affinity and low affinity K+ uptake (Coskun et al. 2010). They suggest that the thermodynamics of K+ transport between influx and efflux are probably + reversed, i.e. at micromolar [K ]ext concentrations influx is active and efflux is passive + process, while at millimolar [K ]ext influx is passive and efflux is active process. They also report that channel-mediated K+ efflux is likely to be the acting mechanism under low external K+. However, under high-K+ conditions it is inoperative together with any other possibly existing K+ pumps, suggesting instead that K+ efflux under these high-K+ conditions occurs from the apoplast instead of the plasma membrane, in way similar to apoplastic bypass flow phenomenon of Na+ in rice. In all cases, K+ efflux studies are still in their beginning, and any conclusions drawn or inferences made on other plant species is a risky task. Nevertheless, this field of study is critical for future investigations for the improvement of the K+ use efficiency by enhancing the ability of 41

plants to maximize K+ uptake either by increasing influx, decreasing efflux, or both (Szczerba et al. 2009), which consequently will lead to lowering the worldwide potash fertilizer expenses made in agriculture.

2.2.4.1.4. Potassium loading to the xylem K+ is transported across the root cells symplastically. As in the case of Na+, it can also be transported apoplastically until reaching the Casparian strip where it enters the symplastic route to reach the stellar parenchyma cells in order to be loaded into the xylem. Recent studies reported that the increased suberization of the root endodermis in the esb1 (enhanced suberin 1) mutant of A. thaliana had little effect on shoot K+ concentration (Baxter et al. 2009), implying that the symplastic pathway, most probably through plasmodesmata, delivers the majority of K+ to xylem parenchyma cells. Early reports on the presence K+ channels in the xylem parenchyma (Wegner et al. 1994) as well as patch-clamp experiments that revealed the presence of KORCs (K+ outward rectifying channels) in stellar root tissues (Roberts and Tester 1995), drew the attention towards investigating the degree of implication of these channels, especially KORCs, in the K+ xylem loading process. Subsequent evidence from A. thaliana mutants demonstrated that the KORC AtSKOR (Shaker K+ outward rectifying channel), present in the root pericycle and stelar parenchyma, contributes to about 50% of the K+ translocated towards the shoot (Gaymard et al. 1998, Lacombe et al. 2000, de Boer and Volkov 2003, Johansson et al. 2006). The xylem K+ concentration ranges from about 2 to 25 mM, depending upon various factors (Marschner et al. 1997). That fact, together with the voltage across the symplast/xylem boundary being about -80 mV (de Boer and Volkov 2003), allows KORCs to load the xylem with K+ concentrations up to about 4 mM. However, loading the xylem with K+ concentrations higher than 4 mM through KORCs requires a substantial depolarization of the stellar parenchyma cells. Thus, other alternative options for loading higher K+ concentrations into the xylem may be achieved by active transport systems against the K+ concentration gradient in the xylem (de Boer 1999).

2.2.4.1.5. Potassium unloading from the xylem, phloem loading/unloading and recirculation K+ is then delivered via the xylem by transpirational water flows until reaching the leaves. K+ is then unloaded from the xylem by AtAKT2/3 channel. AKT2 and AKT3 are differentially initiated transcripts from a single gene and are predominantly expressed in phloem and xylem parenchyma (Marten et al. 1999, Lacombe et al. 2000). AKT2/3 is a weakly-rectifying K+ channel which represents a special case since it was shown to operate in two gating modes. Depending on its phosphorylation state, the AKT2/3 channel is shifted from a voltage-independent mode into a voltage-dependent, hyperpolarization-activated inward rectifier mode (Michard et al. 2005). Thus, this channel mediate both K+ uptake and release depending on the local K+ electrochemical gradients. This curious characteristic makes it involved in phloem loading and/or unloading as well (Marten et al. 1999, Lacombe et al. 2000, Deeken et al. 2002). Transport of K+ in the phloem is important for K+ nutrition of specific tissues (e.g. the root tip) under normal conditions and becomes particularly important under K+-deficient conditions where leaf K+ stores can be mobilized to supply roots with essential K+. It appears that K+ phloem transport plays a central role in the fine-tuning of K+ fluxes within tissues and the whole plant (Amtmann et al. 2004).

2.2.4.2. Potassium homeostasis under salinity stress As described in the first chapter, Na+ competes with K+ on its metabolic functions inside the plant causing toxicity. However, it is likely that the cytosolic K+/Na+ ratio and not the absolute quantity of Na+ per se determines cell metabolic competence and, ultimately, the ability of a plant to survive in saline environments (Shabala and Cuin 2008). It is, thus, widely accepted that plant salinity tolerance may be achieved not only by cytosolic Na+ exclusion mechanisms, described in the first chapter, but also by an efficient cytosolic K+ retention. This notion is based on several reports documenting a strong positive correlation between shoot K+ concentration and genotype’s salinity tolerance in a wide range of plant species (Cuin et al. 2003, 2010, Chen et al. 2005, 2007b, Colmer et al. 2006). Under salinity, root plasma membrane is strongly depolarized as a consequence of a massive influx of positively charged Na+ ions. This makes K+ uptake through KIR

channels thermodynamically impossible, where plants should rely exclusively on K+ uptake via high affinity transport systems, which is much less efficient. This depolarization not only makes K+ uptake more problematic but also causes a massive K+ efflux through GORK channels. These two factors result in a massive depletion in the cytosolic K+ pool. The production of ROS exacerbates the problem by inducing more K+ leakage through GORK and NSCC channels. To overcome these effects, the loss of cytosolic K+ is rapidly reversed through repletion by K+ from the vacuole (Shabala et al. 2006). However, this postpones but does not abolish the problem, as this results in the loss of turgor and immediate growth penalties. Furthermore, if salinity stress is too long, the vacuolar K+ pool is also depleted and the cell collapses. Another temporary strategy for salinity tolerance in plants is to prevent depolarization-induced K+ leakage by restoring (rectifying) the depolarized membrane potential via more active H+ pumping. This strategy also comes at a high metabolic ATP cost and, thus, will result in yield penalties. While Arabidopsis plants show a progressive decline in leaf K+ content with increasing salinity, its salt-tolerant relative Thellungiella halophila was shown to be capable of increasing mesophyll K+ contents under saline conditions. Both KOR and NSCC in T. halophila root plasma membranes display a higher degree of K+/Na+ selectivity (Volkov et al. 2004). Consequently, Na+ influx in T. halophila is reduced as compared to A. thaliana. This leads to the paradigm that the extent of NaCl-induced K+ efflux depends on both the magnitude of salt-induced membrane depolarization (H+ pump activity vs. Na+ entrance through NSCC) and on the activity of all K+-release channels (e.g., KOR and NSCC). It, thus, appears evident that although K+ homeostasis and transport in the plant is mediated by a huge number of transporters and channels, however, under saline conditions only two of these play the major role in maintaining the optimal cytosolic K+ homeostasis and controlling salinity-induced K+ leakage: 1) the depolarization-activated outward rectifying K+ channels (KOR), and 2) and nonselective cation (NSCC) channels. Therefore, together with sequestering and excluding Na+ from the cell, a reasonable strategy to tolerate salinity is to retain K+ in the cell, where screening for salinity resistant plants would be made according to this high K+/Na+ ratio heritable trait (Chen et al. 2005, 2007b, 2007c). Future studies on salt tolerance by targeting K+ homeostasis, functional expression and control of GORK genes are expected to be fruitful in the light of the above mentioned studies. 44

2.2.5. Poorly studied cation transporter families: a new source of information on potassium homeostasis? In order to the elucidate the mechanisms of K+ efflux and vacuolar K+ transport in plants, poorly studied transporter families should be thoroughly investigated and characterized to unravel novel genes and transporters implicated in these mechanisms. Several authors speculated in early reviews that the CPA2 monovalent transporter family might be implicated in K+ efflux (Britto and Kronzucker 2006). This possibility might turn to be correct as in the case of previous speculations on the implication of CHXs in vacuolar K+ transport (Maathuis 2010). The CPA2 family is considered to be the largest and the most diverse of all CPA families. Members of CPA2 family are prevalently found in bacteria, fungi and plants; however, they are rare in animals (Brett et al. 2005). They are predicted to have 8-14 transmembrane helices with a Pfam domain for Na+(K+)/H+ exchange. In general, they catalyze electrogenic transport either through carrier-mediated mode or channel-mediated mode (Saier et al. 2000). The CPA2 family includes the KEA (K+ Efflux Antiporter) and CHX (Cation H+ Exchanger) transporter families. To date, the only KEA transporter characterized in plants is KEA2 of Arabidopsis which is expressed in chloroplasts playing a possible role in K+ and pH homeostasis in that organelle (Aranda-Sicilia et al. 2012). CHX family members on the other hand have been slightly better characterized than KEA transporters and recently, indeed, several CHX transporters (AtCHX17-19) have been demonstrated to localize to prevacuolar compartments, although their in planta roles remain to be characterized (Chanroj et al. 2011). With the exception of AtCHX13 (Zhao et al. 2008) and AtCHX21 (Hall et al. 2006) that localize to plasma membrane, most CHX transporters appear to reside in internal membranes such as prevacuolar, endoplasmic reticulum, Golgi or other nonspecific endomembranes (Padmanaban et al. 2007, Chanroj et al. 2011, Lu et al. 2011). The characterized CHX transporters to date, thus, carry out diverse functions in different plant tissues (roots, leaves, pollen) in the abovementioned organelles ranging from K+ high affinity uptake to internal organelle and cellular K+ homeostasis. In addition to their actual roles in vacuolar K+ transport and speculated implication in K+ efflux, the CHX family is an attractive source of transporters to study for further reasons. Although the CHX family is the most numerous cation transporter family in Arabidopsis (28 members), it is considered as one of the least studied 45

transporter families (Sze et al. 2004). The members of Arabidopsis CHX family, for example, are either preferentially or exclusively expressed in pollen. In light of that, and knowing that this family exists in both early non vascular plants (i.e. bryophytes, among others) that have no pollen, several questions rise on the functions, importance and evolution of CHX transporters in early land plants into higher flowering plants. The answers to these questions ma y shed the light on the important plant changes that happened after plants conquered land regarding K+ transport homeostasis and transport, together with the accompanied changes that occurred in this transporter family. Bearing all these details in mind, we decided to study the CHX family in an early land plant; the bryophyte moss Physcomitrella patens. As will be discussed in the next chapter, the phylogenetic position of P. patens in land plant evolution, among many other characteristics of this model plant, makes it an ideal candidate for our study of the CHX transporter family.

2.3. THE MODEL PLANT Physcomitrella patens 2.3.1. Biological and evolutionary significance of bryophytes The dominance of vascular higher plants in the landscape vegetation throughout much of the world today is the result of a long history of plants conquering land, where the ancestor to land plants was probably born about 490 millions of years ago in the Ordovician period of the Paleozoic era (Sanderson 2003). Millions of years later, a diversity of plants adapted to the terrestrial environment and developed the ability to absorb water and nutrients that will be transported and distributed throughout their aerial shoots, occupied at least some portions of the land masses. Soon thereafter, plants were freed from the need of water for sexual reproduction, by transporting their sperm cells in pollen grains carried by wind or insects to the female sex organs, and seeds protected the newly formed embryo. Angiosperms, with their elaborate flowers and seeds packed with a reserve-filled endosperm, are the last major product of land plant evolution. Interestingly, bryophytes, which evolved during a pivotal moment in the history of life on earth and have persisted for hundreds of millions years (Renzaglia et al. 2007), are considered the closest modern relatives of the ancestors to the earliest terrestrial plants. In fact, they hold the key part in the evolutionary history of land plants; they mark the transition to land and the origin of vascular plants, and hence, link the seed and vascular plants to their algal ancestors. The term “bryophyte” originates from the Greek language and refers to plants that swell upon hydration. Currently, the term bryophyte is a general name given for plants characterized by a life cycle featuring alternating haploid and diploid generations with a dominant gametophyte phase. All bryophytes share several traits, some of which have been retained by all other land plants (e.g. an embryo which gives land plants their name “embryophytes”), and others that are unique (e.g. an unbranched sporophyte, with a single spore producing tissue, or sporangium). Bryophytes can be divided into three major lineages that differ from one another mainly in the architecture of the gametophyte and sporophyte: 1) liverworts (Marchantiophyta), 2) mosses (Bryophyta in the strict sense), and 3) hornworts (Anthocerotophyta). The basis of this classification and differences between the three lineages is beyond the scope of this thesis and can be reviewed elsewhere (Goffinet and Shaw 2009). These three bryophyte lineages together with the fourth lineage, tracheophytes (vascular plants), form the living land plants existing today; all four lineages belong and form the subkingdom Embryophyta and, 47

consequently, all considered embryophytes. Indeed, bryophytes share with other embryophytes many morphological features including multicellular sex organs, a cuticle and the retention of the zygote, which undergoes mitotic divisions within the confines of the archegonium. The microfossil record suggests that bryophytes, and in particular liverworts, were important parts of the earliest plant communities on land, and in fact, the oldest bryophyte fossil is a simple thalloid liverwort (van Aller Hernick et al. 2008). However, fossils offer little valuable chronological information, since most preserved specimens date from more recent periods than the estimated time of divergence. Thus, several attempts to reconstruct the evolutionary origin of embryophytes have been made in the past based on comparing data from morphology, sperm ultrastructure, as well as ribosomal, chloroplastic, or nuclear DNA (Garbary et al. 1993, Mishler et al. 1994, Garbary and Renzaglia 1998, Nishiyama et al. 2003). Nevertheless, the latest effort, combining an extensive set of DNA sequences from all three genomes, sampled from many representatives of the major lineages, provided the most robust hypothesis to date (Qiu et al. 2006): hornworts share a common ancestor with vascular plants, whereas liverworts are a sister lineage to all other extant embryophytes and mosses bridge the gap between liverworts and hornworts (Fig. 1). Although some recent studies report that the dating times of divergences, based on calibrated molecular phylogenies, suggest that mosses arose about 380 millions of years ago (Newton et al. 2007), however, other reports claim an earlier divergence date probably about 450 millions of years ago (Rensing et al. 2008). In all cases, this phylogenetic position of bryophytes in land plant evolution provides great information to reconstruct the origin of the critical morphological, anatomical and physiological innovations of plants to conquer land including all the genes and functions been lost or acquired during the evolution of seed plants. In that way, bryophytes are a primer for understanding the selection forces that shaped the evolution of plants following the transition to land.

Fig. 1. Phylogenetic relationships among extant lineages of green plants (after Mishler and Oliver 2009).

2.3.2. The moss Physcomitrella patens: a model plant for the new millennium In the recent years, Physcomitrella patens (Hedw.) Bruch and Schimp. (sometimes called Aphanorrhegma patens) emerged as a novel plant model system for studying the so called “evo-devo” (evolutionary and developmental) biology of plants. P. patens belongs to the moss lineage of bryophytes (Table 1), an like most plants, it shows an alternation of generations: a haploid phase that produces gametes (the gametophyte generation) and a diploid phase that produces haploid spores by meiosis (the sporophyte generation). However, in contrast to phanerogam embryophytes (seed plants), the gametophyte is the dominant phase in P. patens (Cove 2005).

Table 1. Scientific classification of Physcomitrella patens. Taxon Na me kingdom Plantae Subkingdom Embryophyta Division Bryophyta Class Bryopsida Order Funariales Family Funariaceae Genus Physcomitrella Species Physcomitrella patens Scientific name Physcomitrella patens (Hedw.) Bruch and Schimp.

The life cycle of P. patens (Fig. 2) can be completed in culture in about 8 weeks. Spores germinate to produce the protonemal stage of the gametophyte. Protonema are filaments of cells that extend by the serial division of their apical cells that first develop in chloronemal filaments made up of cells densely packed with large chloroplasts that develop and transition to caulonemal filaments containing fewer, less developed chloroplasts. Thus, the characteristics of the two protonemal filament types, chloronemal and caulonemal, suggest that they apparently have assimilatory and adventitious roles, respectively. Caulonemal filaments divide to produce side branches, which most develop into chloronemal filaments, while others develop into caulonemal filaments or buds. Bud production involves a transition from two-dimensional filament growth to three-dimensional leafy shoot development. These shoots produce gametangia and gametes, and thus, termed “gametophores”. These gametophores (0.5-5 mm) have a stem-like structure that lacks true vascular system, and all along the axis of the stem, it has photosynthetically active leaves that are only one cell layer thick. At the base of the gametophore there are rhizoids, root-like structures that help to keep the gametophore upright (anchoring). Counter-intuitively, the rhizoids do not carry out the same function of higher plant roots (i.e. water and nutrient uptake), given that P. patens is poikilohydric; their water content is directly regulated by the ambient humidity and it is not necessary for them to develop a root system to draw water from the soil. This alternative strategy enables them to grow on very hard surfaces such as rocks and tree 50

trunks that are inhospitable to most vascular plants. P. patens is monoecious; it develops both male (antheridia) and female (archegonia) sex organs on the same gametophores, where self-fertilization is usual. The fertilized zygote develops into the diploid sporophyte which comprises a short seta (stalk that bears the capsule in which spores are produced) which, when mature, contains the spores (approximately 4000 per capsule). Although, there are several described ecotypes of P. patens, nevertheless, the Gransden ecotype (collected in a field near Gransden Wood in Huntingdonshire, England) is the most widespread in laboratories all over the world.

Fig. 2. Life cycle of Physcomitrella patens showing alternation of generations.

P. patens possesses several traits that make it an ideal model plant. As mentioned earlier, the dominant phase in the life cycle of this moss is the haploid gametophytic phase (protonema and gametophores). This phase can easily be propagated and maintained under aseptic lab conditions. Moreover, the moss shows high capacity for tissue regeneration, and thus, growth tests can be started easily using small protonemal inocula as a starter for larger cultures. The relatively easy attainment of the haploid phase allows the immediate identification of mutant phenotypes without the exhaustive work required to obtain haploid plants as in the case of other model

plants like Arabidopsis. The simple protonema structure gives great accessibility to direct observation under different microscopy techniques, allowing a detailed analysis at an individual cell level. Furthermore, protoplast extraction protocols developed to date, yield high percentage of intact and viable protoplasts ready to be used for genetic transformation. These transformation procedures depend mainly on using polyethylene- glycol (PEG)–mediated DNA uptake (cloned in plasmids containing a cassette coding for antibiotic resistance by the isolated transformed protoplasts). Currently, this procedure yields relatively high percentage of transgenics that could be selected as antibiotic-resistant regenerants. This high frequency of transformation in P. patens is primarily thanks to its high frequency of successful homologous recombination; genetic recombination in which nucleotide sequences are exchanged between two similar or identical molecules of DNA, which in this case is between the P. patens specific genome sequences and homologous sequences contained in a transforming plasmid (Schaefer and Zryd 1997), leading to partial or complete knock-out of the desired gene. In fact, current transformation protocols of P. patens permits multiple gene knock-out in a sole reaction, as long as different transforming plasmids with distinct cassettes coding for different antibiotic resistances are available for each gene to be knocked out. Also, several knockout mutants are increasingly being used in biotechnology, especially to produce complex biopharmaceuticals (Decker and Reski 2007). Another major breakthrough that established a solid ground for P. patens as a model plant is the relatively recent publication of its complete genome (500 Mbp organized into 27 chromosomes) sequence (Rensing et al. 2008) together with the availability of advanced bioinformatic tools and databases (www.phytozome.net). This is of great use knowing that P. patens possess 66% homologue genes of the approximately 26000 genes present in the model plant A. thaliana (Nishiyama et al. 2003), which opens important lines of investigation especially in the comparative genomics field. It is also worth mentioning that different P. patens ecotypes, mutants, and transgenics are freely available to the scientific community by the International Moss Stock Center (www.moss-stock- center.org/). A rather interesting trait of P. patens is its high salinity tolerance. Several studies documented Physcomitrella’s tolerance to salt and indicated that not only when exposed to salt concentrations up to 350 mM was able to recover normally (Frank et al. 2005), but was also able to tolerate NaCl concentrations up to 600 mM when the plants had 52

been slowly adapted to increasing salt concentrations (Benito and Rodriguez- Navarro 2003). This high degree of tolerance has been attributed to the presence of a Na+-pump ATPase PpEna1, which is usually not found in flowering plants and therefore may have been lost during the evolution of land plants (Benito and Rodriguez- Navarro 2003, Fraile-Escanciano et al. 2009). In addition, P. patens has all the K+ and Na+ gene families present in A. thaliana. This encouraged scientists to consider P. patens as a good source of genes involved in salt tolerance, and encouraged the unraveling of other implicated mechanisms in its salinity tolerance (Wang et al. 2008).

P. patens is also a very attractive model plant to study K+ homeostasis in general, and under salinity conditions in particular. After conquering land, plants faced a substantially new oligotrophic habitat where K+ is far less available compared to their original sea water habitat. Inevitably new land plants, thus, had to broaden the spectrum of their K+ utilization and internal cellular homeostasis. This was achieved by evolving their K+ transport systems, mainly in terms of multiplying already existing transporters or by developing unique transport systems for K+ accumulation and distribution in different organelles with different functions and affinities, especially during the process of vascularization of plants (Gomez-Porras et al. 2012). Once again, the phylogenetic position of P. patens in land plant evolution opens the possibility to study key changes in K+ transporter families during the course of evolution and thus, as will be described in the next chapters of this thesis, we have studied two members of CHX K+ transporter family of Physcomitrella patens, a family that has suffered major changes of diversification during the course of evolution of early plants into flowering plants.

3. MATERIALS AND METHODS

3. MATERIALS AND METHODS 3.1. Biological materials 3.1.1. Bacterial strains used and culture conditions The different Escherichia coli strains used during the accomplishment of this work are described in table 2.

Table 2. Bacterial strains used. Bacteria Genotype Reference F- endA1- hsdR17(rK- mK+) supe44 DH5α thi-1 lambda- recA1 gyrA96 realA1 (Hanahan 1983) 80dlacZAM15 thi rha lacZ nagA recA Sr::Tn10 (Schleyer and Bakker TKW4205 ∆kdpABC5 trkA405 kup1 1993)

The E.coli strain DH5α was routinely used for plasmids multiplication (Sambrook et al 1989) (Table 2). The strain was routinely grown in Lysogeny Broth (LB) medium (Table 3), supplemented with 50 mM KCl, at 37 ºC. For the complementation drop-tests of strain TKW4205, solid minimal medium (MM) (Table 3) supplemented with trace elements was used as described in Rhoads (1976). MM medium pH was either adjusted to 5.5 by adding 10 mM MES, or to pH 7.5 by adding 10 mM HEPES.

Table 3. Composition of culture media for bacteria. LB MM

Yeast extract 0.5% MgSO4 0.4 mM

Bacto-Tryptone 1% FeSO4, 6 µΜ NaCl 1% Citric acid 1 mM Biotin 1 mg l-1 Glycerol 0.2% Asparagine 8 mM

CaCl2 20 mM

3.1.2. Yeast strains used and culture conditions The Saccharomyces cerevisiae strains used in this work are described in table 4.

Table 4. Yeast strains used. Yeast Genotype Reference (Haro and Rodríguez- W∆6 Mat a ade2 ura3 trp1 trk1∆::LEU2 trk2∆HIS3 Navarro 2003)

B3.1 ena1∆::HIS3::ena4 nha1∆::LEU2 (Bañuelos et al. 1998)

MATa ade2-1 can1-100 his3-11,15 leu2-3,112 trp1-1 ura3-1 (Maresova and LMB01 mall0 ena1Δ::HIS3::ena4Δ nha1Δ::LEU2 Sychrova 2005) kha1Δ::kanMX

The yeast strain W∆6 defective in the main influx systems of K+ (TRK1 and TRK2) was used for the functional characterization of possible high affinity K+ or Na+ influx transport. This mutant displays a phenotype of non-growth in the presence of low concentrations of K+. The strain B3.1, defective in the systems of Na+ efflux ENA1-4 and NHA1, was used for the functional characterization of possible Na+ or K+ efflux transport, because it displays a phenotype of non-growth in the presence of high concentrations of Na+ or K+. Also, the strain LMB01 defective in the systems of Na+ and K+ efflux ENA1-4, NHA1 as well as Golgi complex K+ antiporter KHA1 was used. 58

This strain display defective growth at alkaline pH with very low concentrations of K+ and was used for the functional characterization of internal pH and K+ homeostasis. All yeast strains were cultured in either rich medium Yeast-extract Peptone Dextrose (YPD) (Sherman 1991), Synthetic Defined minimal medium (SD) (Sherman 1991), or Arginine Phosphate (AP) medium (Rodríguez-Navarro and Ramos 1984) (Table 5). In order to select and maintain a particular plasmids or knock-out strain, the SD and AP media were supplemented with the corresponding amino acids, vitamins and trace elements (Table 6). AP and SD media were prepared using Milli-Q water. Growth of all strains was carried out at 28 ºC, and in a shaking incubator at 250–300 r.p.m. in the case of liquid media. Yeast strains and clones were stored at –80°C in YPD with 15% glycerol for future use. Depending on the experimental conditions, AP media were supplemented with the indicated K+ concentrations and with 10 mM Mes-Ca2+ for pH 6.0, 10 mM Taps- Ca2+ for pH 7.5, and 10 mM tartaric acid-Ca2+ for pH 4.5. The growth of yeast cells was monitored by recording the absorbance at 540 nm in a Microbiology Reader Bioscreen C workstation (Growth Curves Oy, Finland), which enables many samples to be run at the same time.

Table 5. Composition of different culture media used for Saccharomyces cerevisiae. SD YPD AP Yeast Bacto-agar 2% 1% (w/v) H3PO4 8 mM extract

Glucose 2% Peptone 2% (w/v) MgSO4 2 mM Yeast nitrogen base

w/o amino acids or 0.7% Glucose 2% (w/v) CaCl2 0.2 mM ammonium Bacto- (NH4)2SO4 0.5% 1.5% Arginine 10 mM agar Trace 1 ml elements Vitamins 1 ml Glucose 2% (w/v)

Table 6. Trace elements and vitamins used in culture media of Saccharomyces cerevisiae. Trace Elements Vitamins

BO3H3 0.8 mM Biotin 0.8 µΜ

CuSO4 5 H2O 0.016 mM Niacin 0.32 µΜ KI 0.06 mM Pyridoxine 0.19 µΜ

FeCl3.6 H2O 0.07 mM Thiamine 0.12 µΜ

MnSO4.H2O 0.26 mM Calcium Pantothenate 0.08 µΜ

Na2MoO4 0.08 mM

ZnSO4.7 H2O 0.14 mM

3.1.3. Physcomitrella patens ecotype and culture conditions The ecotype of Physcomitrella patens subsp patens used in this work was the Gransden ecotype, provided by the laboratory of Dr. Andrew Cuming, University of Leeds. Physcomitrella was maintained in the laboratory in its different phases life cycle; protonema, gametophore and sporangiophore using different media and culture conditions as will be described. The manipulation of Physcomitrella was always carried out under aseptic conditions in a laminar air flow chamber.

3.1.3.1. Growth and propagation conditions of protonema and gametophore Protonema and gametophore cultures were grown in growth chambers at 25 ºC under 200 µmol m-2 s-1 of irradiation with permanent illumination. The culture media used are described in table 7. In the case of solid media, 5 g l-1 agar was added to the described media. The BCD medium was routinely used for the culture and the propagation of Physcomitrella gametophores, while BCDNH4 medium was used for the growth of the protonema phase, since ammonium tartrate favors its predominant formation and maintenance, especially the cloronema state (Ashton and Cove 1977). Liquid KFM medium (K+ Free Medium, also named “POM”) is an oligotrophic medium free from Na+ and K+ that allows the control of these cations concentration for the convenience of experiments (Garciadeblás et al. 2007). This medium was used for obtaining Physcomitrella K+-starved plants (Table 7).

Table 7. Media used for growing Physcomitrella patens.

KFM (POM) BCD / BCDNH4

CaCl2 (1M) 1 ml

KNO3 10 mM

Ca(NO3) 4H2O 1.5 mM

MgSO4 7H2O 1 mM 1 mM

FeSO4 7H2O 45 µM 45 µM

CuSO4·5H2O 0.055 mg/l

ZnSO4·7H2O 0.055 mg/l

H3BO3 0.614 mg/l

MnCl2·4H2O 0.389 mg/l 1 ml l-1 1 ml l-1 CoCl2·6H2O 0.055 mg/l KI 0.028 mg/l

Na2MoO4·2H2O 0.025 mg/l

H3PO4 1.8 mM 1.8 mM

NH4 tartrate (4.6 mM ) pH 5.8 6.5

Different concentrations of media components are shown. BCDNH4 medium is equal to BCD medium supplemented with to 4.6 mM ammonium tartrate. The concentration of K+ is 0.17 mM in KFM and 11.8 mM in BCD.

KFM medium was prepared by dissolving the components indicated above in Milli-Q water. For time-saving purposes, the BCD medium was prepared from the three stock solutions indicated in table 8, in addition to the trace element solution indicated earlier in table 6 (1 ml per L of medium). These solutions were always kept at 4 ºC. After the desired volume of BCD medium was prepared from the stock solutions and sterilized by autoclaving, it was left to cool down to avoid the precipitation of the required concentration of CaCl2 (1 M) that will be finally added.

Table 8. Stock solutions for BCD medium. Solution B (for 1 L) Solution C (for 1 L) Solution D (for 1 L)

MgSO4 7H20 KH2PO4 KNO3 ………….……25 g ….……………….25 g ………………101 g

pH adjusted to 6.5 with 4M FeSO4.7H20 KOH .………………...1.25 g

Control plants were grown in KFM medium with 4 mM KCl. K+-starved plants were obtained by transferring plants to KFM medium supplemented with no KCl and grown there for 7-10 d. KFM pH was adjusted to 5.8 as a standard condition. Depending on the experimental conditions, the pH values of media were adjusted with HCl and

NaOH, or KOH in the case of BCD/BCDNH4, and with Ca(OH)2 in the case of KFM. Media with pH 9.0 were buffered with 10 mM TAPS, while buffered with 5 mM tartaric acid when pH is adjusted to 4.5. For the culture and propagation of Physcomitrella protonema and gametophore, the following procedures have been followed: 1) Propagation of protonema in solid media:

The propagation of protonema was carried out in BCDNH4 solid medium plates, on which, previously sterilized cellophane discs (cellulose film 325P, A.A Packaging limited, U.K) were placed. The use of these cellophane discs facilitated the later harvesting of protonema or their transference from one plate to another. The protonema harvesting was carried out with the aid of a sterile tweezers, and the plant material was transferred to a 50 ml sterile Falcon tube. Afterwards, protonema was crushed in 5-10 ml of sterile water, depending on the amount of plant material, with the help of a Polytron PT2100 homogenizer (Kinematica AG, Lucerne, Switzerland). The obtained crushed suspension of plant material was then poured into three BCDNH4 plates. Also, protonema can be obtained from the crushing of gametophore material by the same process described above, although an additional step of crushing with the homogenizer is required, one week after the first one, to obtain a homogeneous protonema culture.

After one week of culture in BCDNH4 plates, newly grown protonema can be observed. 62

2) Propagation of gametophore in solid media: The propagation of gametophores was realized by fragmenting the old gametophore material with a sterile scalpel every 2-3 weeks. The fragmented material was then passed to BCD plates without cellophane to favor a fast growth, and letting new material grow for about 2-3 weeks. 3) Propagation of protonema in liquid media: To propagate protonema in liquid media, 1 L or 500 ml crystal bottles mounted with air bubbling equipment were used. When large amount of plant material is needed, 8 L bioreactors with agitation and ventilation caps were used. In all cases, the cultures were grown for several weeks. As in the case of solid media, the culture of protonema in liquid media was also carried out in BCDNH4. In order to duplicate the mass more quickly, protonema was crushed with the homogenizer directly in the bottle approximately every 4 d. If the protonema culture is left to grow for over a week and a half, protonema begins to differentiate to gametophores. 4) Propagation of gametophores in liquid media: The propagation of gametophores in liquid media was carried out in bottles or biorreactores starting from gametophores previously grown in BCD plates. The material was then fragmented with either a sterile scalpel or homogenizer, ensuring that the resulting fragments were approximately between 1 and 3 cm. The inocula with which we started all experiments were almost exclusively protonemata.

3.1.3.2. Culture conditions for obtaining sporangiophores. Fragmented gametophore material, with either homogenizer or scalpel, was inoculated in sterile jars containing previously hydrated and sterilized Jiffy-7 soil (Jiffy Products International). Physcomitrella sporangiophores were obtained by growing gametophores cultures under 8 h of light and 16 h of darkness at 15 ºC. One month later, a dense turf of gametophores was formed, and after the second month of growth, sporangiophores started to form. Once the capsules of the sporangiophores matured (brown color), they were collected and kept in an eppendorf tube and stored in dry conditions.

3.1.3.3. Conservation of Physcomitrella patens The conservation of the obtained mutants of Physcomitrella (described later on in this chapter) was done by following two different strategies. First, by conserving the sporangiophore capsules, thus, allowing their conservation for several years. Second, by conserving gametophore material in BCDNH4 plates at 7ºC subjected to a cycle of 22 h of darkness and 2 h of light. The latter conditions allowed the cultures to stay viable for several months.

3.1.4. Plasmids The plasmids used in this work are described in table 9.

Table 9. The plasmids used and their characteristics. Plasmid Characteristics Reference Bacterial multicopy Plasmid for PCR product pCR2.1-TOPO Invitrogen cloning. Amp R km R

Bacterial multicopy plasmid of inducible (Guzmán et al. pBAD24 expression. pBAD promoter, araC regulator 1995) dependant on arabinose concentration; Amp R

Multicopy plasmid for expression in yeast. (Brunelli and pYPGE15 PGK promoter, CYC terminator, URA marker, Pall 1993) Amp R

Plasmid of expression in plants for obtaining (Rubio-Somoza pMF6-GFP gene fusions with GFP. 35S promoter, adh1 et al. 2006) intron, mGFP5 gene, 3´NOS terminator, Amp R

Plasmid used for the construction of PpCHX1 Dr.Hasebe, Dr. gene disruption. AmpR for E.coli and HygR for Nishiyama and pTN186 Physcomitrella. Hygromicin cassette under the Dr. Hiwatashi, CamV35S promoter and terminator. Japan

Plasmid used for the construction of PpCHX2 Dr.Hasebe, Dr. gene disruption. A mp R for E.coli and ZeoR for Nishiyama and P35S-Z e o Physcomitrella. Zeocin cassette under the Dr. Hiwatashi, CamV35S promoter and terminator. Japan

3.2. DNA manipulation techniques 3.2.1. Bacterial plasmidic DNA purification Nucleic acid manipulation was carried out following standard protocols (Sambrook et al. 1989). Two methods were implemented to extract plasmidic DNA from bacterial cultures in small scale. The first method, “alkaline lysis and boiling” method was implemented when the plasmidic DNA was going to be used for analysis of clones (Sambrook and Russell 2001). When the plasmidic DNA was aimed for sequencing, the DNA extraction kit “QIAprep spin miniprep kit” (QIAGEN) was used. 65

On the other hand, when plasmidic DNA is needed in large scale, “Maxiprep alkaline lysis” method was used as described by Sambrook (1989). In all cases, a single isolated colony of E.coli was inoculated in liquid LB supplemented with the convenient selection antibiotic and incubated overnight at 37 ºC under continuous shaking of 200 r.p.m. On the following day, cells were harvested by centrifugation at 13000 r.p.m. during 1 min.

3.2.2. Plant genomic DNA purification For the extraction of Physcomitrella plant DNA, the DNA plant extraction Kit “DNeasy Mini Plant Kit” was used. Nevertheless, when large scale DNA purification from gametophore material was needed, an extraction protocol based on CTAB was used as follows (Schlink and Reski 2002): Protocol for genomic DNA extraction by CTAB method: First, 1-5 mg of protonema material was picked from solid media plates and placed in 1.5 ml eppendorf tube. Liquid nitrogen was carefully poured over the material then crushed and homogenized with the aid of a plastic rod fixed to a drilling machine. Immediately afterwards, 300 µl of extraction buffer (Tris-Cl 0.1 M pH 8.0, NaCl 1.42 -1 M, CTAB 2%, Na2EDTA 20 mM, PVP-40 2%) supplemented with 1 mg ml ascorbic acid and 7 µl of β - mercaptoethanol preheated to 65 ºC was added to each 10 ml of crushed plant suspension. To this mixture, 1 µl of RNAse A (10 mg ml-1) was added and incubated during 15 min at 65 ºC. Next, chloroform: isoamyl alcohol (24:1) was added and the tube centrifuged for 10 min at 13000 r.p.m. The supernatant was then transferred to a new eppendorf tube and the genomic DNA was then precipitated with isopropanol, centrifuged for 10 min at 13000 r.p.m, and then the precipitate was washed with ethanol 70% for 5 min by centrifuging at 13000 r.p.m. Finally, the precipitate was dried in a speed vacuum centrifuge and re-suspended in 50 µl of sterile distilled water.

3.2.3. DNA Quantification The extracted DNA concentration was measured using NanoDrop spectophotometer (ND-1000 Spectrophotometer). Alternatively, the DNA concentration was also estimated by comparing the intensities of the DNA bands, separated by gel electrophoresis in TBE agarose gel at 0.8% (w/v) dyed with ethidium bromide, with the

pattern of bands of λ HindIII marker (λ Phage digested with HindIII restriction enzyme).

3.2.4. DNA amplification by PCR PCR experiments were carried out in PCR Thermal Cycler “GeneAmp 2400” (Perkin-Elmer). The amplification enzyme routinely used was “Taq DNA polymerase” (GE Healthcare). When an enzyme with greater fidelity of amplification was required, “Expand High Fidelity PCR System” (Molecular Roche Biochemicals) or PfuII Ultra polymerase (Stratagene) was used. The PCR reactions were carried out following the manufacturer’s recommendations and the primers used in the reactions were synthesized by Sigma-Aldrich. The length of PCR fragments was then analyzed by electrophoresis separation in agarose gel.

3.2.5. Cloning and purification of PCR fragments When cloning or sequencing was required, PCR fragments were cloned into the pCR2.1-TOPO vector of kit “TOPO TA cloning kit” (Invitrogen). DNA fragments from an excised agarose gel band were extracted using the PCR product purification kit “Gene Clean Turbo Kit” (Q-biogene) following the manufacturer’s instructions. Then, the extracted DNA from the agarose gel bands was cloned into the pCR2.1-TOPO vector of kit “TOPO TA cloning kit”. In all the cases, the PCR amplified products were sequenced to verify fidelity of amplification and absence of mutations.

3.2.6. DNA Digestion and ligation of DNA fragments The restriction enzymes used were obtained from Molecular Roche Biochemicals, using the reaction conditions recommended by the manufacturer. In some cases, dephosphorylation of the restricted DNA 5´ ends was carried out, to avoid the digested ends’ re-ligation, using alkaline phospatase purified from shrimp (Molecular Roche Biochemicals), following the manufacturer’s instructions for the required reaction conditions. For DNA fragments ligation, T4 DNA ligase was used (Molecular Roche Biochemicals).

3.3. RNA manipulation techniques 3.3.1. Purification of total RNA of plants For the extraction of Physcomitrella total RNA, the kit “RNeasy Plant Mini kit” (Qiagen GmbH, Hilden, Germany) was used. Nevertheless, when large scale RNA purification was needed, an extraction protocol based on CTAB was used (Schlink and Reski 2002) as follows: The content of approximately two plates of Physcomitrella protonema was placed in a previously sterilized mortar and crushed with liquid nitrogen. 500 µl of extraction buffer (same buffer described earlier in the protocol of DNA extraction) and 500 µl phenol: chloroform (24:1) is added and then passed to an eppendorf tube. The mixture was vortexed until observing the appearance of a green pistachio color. The samples were then centrifuged during 2 min at 13000 r.p.m. Afterwards, 50 µl of sodium acetate 3 M pH 5.5 was added to the obtained precipitate and left in ice for 5 min. Next, the samples were centrifuged for 2 min at 13000 r.p.m., then 500 µl ethanol was added to supernatant, and then the eppendorf tubes were quickly submerged in liquid nitrogen for 5 sec, passed to ice, and centrifuged 5 min at 13000 r.p.m. Then, 30 mg NaCl was added to the obtained precipitate and left overnight at 4 ºC. On the following day, the samples were centrifuged at 13000 r.p.m. at 4 ºC for 10 min, the supernatant was discarded and 40 µl of cold NaCl 2.5 M was added to the precipitate. Finally, the precipitate was washed with ethanol 80% (v/v), dried for 15 min, and resuspended in 30 µl sterile distilled water.

3.3.2. RNA quantification The obtained RNA concentrations were measured using NanoDrop spectrophotometer (ND-1000 Spectrophotometer). In order to verify the quality of the RNA, a small sample was run in 1% TAE gel in which the gel bed and chamber were previously treated with 0.1% SDS. A mixture of blue loading dye containing formamide was added to the samples before running the gels.

3.3.3. cDNA synthesis by RT-PCR and study of the genic expression by quantitative real time PCR (qRT-PCR) To analyze the expression of the PpCHX genes, the plants were grown for one week in different conditions of pH, Na+ and K+ (table 18, in results chapter 4). The PCR amplification of cDNAs was performed with a double-stranded cDNA Synthesis System Kit (GE Healthcare, http://www.gehealthcare.com) from RNA extracted from plants grown under the conditions described in table 18. Quantitative Realtime PCRs (qRT- PCR) were performed using an ABI Prism 7000 Sequence Detection System (Perkin- Elmer Applied Biosystems) and SYBR Green PCR Master Mix (Perkin-Elmer Applied Biosystems). Amplifications were carried out according to standard procedures as described in Garciadeblás et al. (2003). The results were expressed as the transcript level ratios between the studied PpCHX and actin genes in the same preparation. PCR primers (Table 10) were designed to amplify the following fragments (numbering according to database): PpACT5 (from nucleotide 1281 to 1422, sequence 109052 in JGI P. patens genome), PpCHX1 (from nucleotide 1903 to 2051) and PpCHX2 (from nucleotide 2102 to 2260). Each pair of primers was used in PCR experiments with all the purified cDNAs to check that they amplified only the fragment for which they were designed.

Table 10. Primers used in the quantitative RT-PCR. Sequence of primers are 5´to 3´. Transcript Forward Primer Reverse Primer PpCHX1 GCTCTGGCATATGGTTTCCGTA ACCTCCGTCTTGCCAATCTCAG PpCHX2 ACCAAGATCAGGAACGCAAGTTG CTGCTATTGCGGCTTGGACCACC Actin5 GTACGTGGCGATCGACTTC GGCAGCTCGTAGCTCTTCTC

3.3.4. Cloning of PpCHX1 and PpCHX2 cDNAs, and the in vitro construction of PpCHX2.1 Primers used to amplify the different genes and/or cDNAs are summarized in table 11.

Table 11. Primers used to amplify the different cDNAs. Primers Sequence 5´-3´ Observations

PpCHX1-ATG1 TATAAGTTCTACAAAGCCTC Cloning of PpCHX1 PpCHX1-ATG2 CAAGATGGCGGACGCTGTGG cDNA in pCR2.1- PpCHX1-ATG3 GGCGTGCAAAACTATGTCGG TOPO plasmid PpCHX1-STOP TTAGCCAGCCAAGACCTTCGCA PpCHX1-1R AAAGCTCCCTCTGGTGCAAT Sequencing of PpCHX1 PpCHX1-2R GCAACGAAAGGCAGAGTGAT 5’ end PpCHX1-3R AATGGACATAGCTTGCTTTC CGGATCCTAAAAAATGGCGGACGCT PpCHX1-ATGBamHI2 Cloning of PpCHX1 GTGGC cDNA in pYPGE15 GGGGTACCTCACACGCGGTGGTCAG PpCHX1-StopKpnI plasmid GAGAA PpCHX1-639 TTGCATGGGTTCTGTTGTGT Primers used to confirm PpCHX1-1050 TCTGGGTAAAATTCTCGGAAC fidelity of amplification PpCHX1-1050- R GTTCCGAGAATTTTACCCAGA of the PpCHX1 cDNA PpCHX1-1107 CAAAGCTTTAACTCTCGGCATC cloned in the different PpCHX1-1146 ATTGGTGGAGCTTATTGTTCTG plasmids used TACTAGTAGGAGGAGATAAGATGGC SpeIATG1-PpCHX1 Cloning of PpCHX1 GGACGCTGTG cDNA in pBAD24 GGGGTACCTCACACGCGGTGGTCAG PpCHX1-StopKpnI plasmid GAGAA PpCHX2.2 AGTGTTGCAAGATGGCGACCGGA Cloning of PpCHX2 PpCHX2-stop {2B in fig. cDNA in pCR2.1- GTTTTCAAGATGGGTCTAGGATC S4} TOPO plasmid PpCHX2-1R GAAGACAACAAACGGGCCGAATT Sequencing of PpCHX2 PpCHX2-2R AACGGGCCGAATTTCACATCACC 5’ end PpCHX2-3R TATTAAGGAGGACAAAGGAGACC Intron removal for CHX2-in t -2 {1A-B in fig. TTCTTGAACAGATACCATATCGCCAC construction of S4} C PpCHX2.1 version CHX2-in t -2R {2A-B in TATGGTATCTGTTCAAGAAGCGGATC

fig. S4} AC Cloning of PpCHX2 GTGGATCCTACACAATGGCGACCGG PpCHX2-ATG-B a m H I cDNA in pYPGE15 AAATGC plasmid

GAGGTACCAGGATCTAGACATGAGG PpCHX2-Stop-KpnI CGCCGA PpCHX2-622 TGCCTCCTCGCTCTCGCTGT Primers used to confirm fidelity of amplification PpCHX2-1259 TGACTACTCCCCTTGTGATG of the PpCHX2 cDNAs cloned in the different plasmids PpCHX2-1715 TCCACAAAACCCAAAGACTC PpCHX2-1969 GTGATCCGCTTCTTGAACAG Cloning of PpCHX2 TACTAGTAGGAGGAGATAAGATGGC SpeIATG-P p CH X 2 cDNA in pBAD24 GACCGGAAAT plasmid GAGGTACCAGGATCTAGACATGAGG PpCHX2-Stop-KpnI CGCCGA Verification of GCGAGAAAGCGAAGACGGTG existence of PpCHX3 PpCHX3-2 gene in P. patens genome GATGCTGCACAAGTGATTTCG PpCHX3-2R

Verification of TCAAGGCGAAATAGCAGAGAC existence of PpCHX4 PpCHX4-2 gene in P. patens genome GTGCTGAGAATAATTGGTGGG PpCHX4-2R

The in vitro construction of the PpCHX2.1 cDNA was carried out by a 2-step PCR method using the PpCHX2 cDNA as a template, as detailed in figure 3. The fragment removed extended from nucleotide positions 1988 to 2075 of the PpCHX2 cDNA. The primers used to produce the desired fragment removal are described in table 11. The resulting PCR fragments were first cloned into the PCR2.1-TOPO vector using the TOPO TA Cloning Kit (Invitrogen). For expression in yeast and E. coli, the verified PpCHX2.1, PpCHX1, and PpCHX2 cDNA sequences (cloned in PCR2.1- TOPO) were used as will be described next.

Fig. 3. Two step PCR strategy for in vitro construction of PpCHX2.1.

3.4. Methods of cellular transformation 3.4.1. Transformation of bacteria and yeast For the transformation of Escherichia coli, competent cells were prepared by the “18 ºC” method in the protocol described by Sambrook et al. (1989). E. coli DH5α competent cells were refreshed (zigzag) on a solid LB medium plate starting from a previously frozen (-80 ºC) culture and incubated O/N at 37 ºC. On the following day, a pre-inoculum was made, where 4 ml of liquid SOB (Table 12) were inoculated with the fresh cells grown on the LB plate and left shaking for the whole morning. At the afternoon, approximately 50 µl of the pre-inoculum was added to 250 ml flasks containing fresh SOB medium and left for 2 d at 18 ºC shaking at 200-250 r.p.m. After

2 d, the flasks were removed from incubation and placed in ice. Then, the culture O.D550 was measured, and incubated again if necessary, until reaching 0.6-0.7 (if the culture exceeded that value, the process should be re-started all over again from the pre- inoculum stage). Under aseptic conditions of the laminar air flow chamber, DH5α cells, sterile TB solution (Table 11), beaker flasks, plastic bottles, and pipettes were kept cold on ice. The cell culture was evenly distributed between two sterile plastic bottles and 72

centrifuged for 10 min at 6000 r.p.m. and 4 ºC. The supernatant was then discarded and the pellet of each bottle resuspended in 40 ml TB. The resuspended cells of the two bottles were finally joined in one bottle and left for 10 min in ice. The suspension was then centrifuged for 10 min at 6000 r.p.m. at 4 ºC and the supernatant discarded. Next, the pellet was resuspended in 20 ml TB with added 7% DMSO and left in ice for 10 min. Finally, the suspension was divided in aliquots of 400 and 600 µl, quickly frozen in liquid N2 and stored at -80 ºC for future use. In the case of preparation of E. coli strain TKW4205 competent cells, 25 ml of liquid LB medium with added 50 mM K+ was inoculated with fresh cells (previously grown O/N on solid LB with 50 mM K+ solid medium plates) and incubated at 37 ºC until reaching O.D550 of 0.4-0.5. The culture was then divided into two ice-cold falcon tubes and centrifuged for 5 min at 6000 r.p.m. The supernatant was discarded, the pellet resuspended in 12.5 ml of cold CaCl2 100 mM and left in ice for 30-60 min in cold chamber (4 ºC). Next, the suspension was centrifuged for 5 min at 6000 r.p.m., the supernatant discarded and the pellet resuspended in 2.5 ml CaCl2 100 mM. The cells were kept at 4 ºC in the cold chamber until being used in the same day for transformation.

Table 12. Composition of SOB and TB. SOB TB Triptone 2% 10 g Pipes (10 mM) 1.2 g

Yeast extract 2.5 g CaCl2 (15 mM) 0.88 g NaCl (10 mM) 0.28 g KCl (250 mM) 7.44 g

KCl (2.5 mM) 0.092 g MnCl2 (55 mM) 3.56 g

MgCl2 (10 mM) 1 g H2O (Final volume) 400 ml

MgSO4 (10 mM) 1.2 g pH 6.7 adjusted with

H2O (Final volume) 500 ml KOH (2 M)

For the transformation procedure of E.Coli competent cells, a 200 µl aliquot was used. The cells were incubated for 30 min in ice with the suitable concentration of DNA (10 µl ligation DNA, 2 µl PCR DNA or 1 µl purified DNA). Then, the cells were subjected to a heat shock of 42 ºC for 30 sec for DH5α (90 sec in the case of TKW4205

strain) and immediately transferred back again to ice for 2 min. Next, 800 µl liquid LB were added to the suspension and incubated for 45-60 min at 37 ºC with continuous shaking at 200 r.p.m. Passed that time, the cells were centrifuged for 5 min at 6000 r.p.m. and the supernatant discarded afterwards. The pellet was then resuspended in 100 µl LB and uniformly distributed into LB solid medium plates (containing adequate concentration of the desired antibiotic for plasmid selection) with the aid of glass beads. Finally, the plates were shortly left in the laminar air flow chamber to dry and then incubated O/N at 37 ºC for the selection of transformed colonies the next day. For yeast transformation, the procedure followed was cellular permeabilization with PEG4000, described by Elble (1992). A flask containing YPD liquid medium (with added K+ when necessary, depending on the strain needs) was inoculated with the desired yeast strain to be transformed and left to grow O/N under continuous shaking at 28 ºC. Next day, 1 ml of the culture is centrifuged for 2 min at 6000 r.p.m. and the supernatant discarded afterwards (this step was repeated twice). Approximately 1 µg plasmidic DNA was added to the pelleted cells together with 500 µl of PEG 35% (w/v)- LiAc 0.1 M solution and the mix was carefully resuspended and left O/N at room temperature. Next day, the suspension is centrifuged for 2 min at 6000 r.p.m. and the supernatant discarded afterwards. Then, the pellet was washed with 1 ml sterile water by centrifuging again for 2 min at 6000 r.p.m. After discarding the supernatant, the pellet was resuspended in 100 µl of sterile water and uniformly distributed into AP solid medium plates (containing adequate concentration of the desired amino acids depending on the strain auxotrophy) with the aid of glass beads. Finally, the plates were shortly left in the laminar air flow chamber to dry and then incubated for 4 d at 28 ºC for the selection of transformed colonies.

3.4.2. Transformation of protoplasts of Physcomitrella For the transformation of Physcomitrella we used a protocol developed by the laboratory of the Dr. Cuming, University of Leeds, based on Grimsley et al. (1977). This protocol involves the transformation of protoplasts, where they were obtained from protonema by either one of the following three procedures: (1) from gametophore culture of Physcomitrella previously crushed and grown for 7 - 10 d in bioreactors with

BCDNH4; (2) from protonema grown on cellophane discs in solid BCDNH4 plates for 5 d; or (3) from protonema grown in bottles of liquid BCDNH4 for 5 d.

Transformation of Physcomitrella protoplasts according to the protocol of Dr. Cuming laboratory: 1) Obtaining and transformation of protoplasts First, we prepared driselase/mannitol solution, where 100 mg of driselase was dissolved in 10 ml of 8% mannitol (nonsterile). The driselase is dissolved directly in a centrifuge tube (10 ml) and mixed from time to time by inverting the tube during 15 min. Then, the solution is centrifuged at 2500 r.p.m. for 5 min and the supernatant is sterilized by filtration through 0.22 or 0.45 µm Millipore filters. Afterwards, the content of approximately two plates of protonema grown in solid BCDNH4 medium was collected and added to the 10 ml solution of driselase/mannitol and left incubating between 45 and 90 min at room temperature, mixing the suspension from time to t ime until all the filaments of protonema became almost invisible. The suspension of protoplasts was then filtered through a 100 µm sieve, and the filtered protoplasts transferred to 10 ml sterile crystal tubes. These tubes were centrifuged at 700 r.p.m. for 4 min with brakes in off-mode. The precipitated protoplasts were then very smoothly resuspended in a volume of approximately 0.1-0.2 ml, slowly spinning the tube between the fingers. Next, the protoplasts were washed twice with 10 ml of 8% mannitol and counted under the microscope on a Thoma counting chamber. Finally, the protoplasts were re-suspended in a volume of 8% mannitol so that the number of protoplasts was 3.6 x 106 protoplasts ml-1. Then, 150 µl of protoplasts were transferred using sterile Gilson pipette yellow tips cut in their ends to 2 ml eppendorf tube containing magnesium-mannitol-MES solution (MMM) (Table 13) and mixed smoothly. Next, 15 µg DNA was quickly added to the protoplast suspension and mixed with the yellow tip making very smooth circular motions. All the suspension was then transferred to a 10 ml tube containing 300 µl PEGcms (Table 13) using a sterile glass Pasteur pipette. The tubes were transferred to a water bath at 45 ºC for 5 min and left afterwards for 10 min at room temperature. 2) Regeneration of protoplasts: The suspension was then diluted successively with 8% mannitol in 6 consecutive steps; 300 µl was added once, 600 µl twice, 1 ml twice and finally completing to a final 75

volume of 8 ml. In each dilution the protoplasts were mixed smoothly and slowly spinning the tube between the fingers. After the successive dilutions, the protoplasts were left for several hours to settle at the bottom of the tube. Finally, the supernatant was discarded and 2 ml of liquid protoplast regeneration medium (PRM-L medium) (Table 14) was added. The protoplasts were left all night in darkness at 25 ºC. The following morning, the protoplasts were transferred to light for several hours, discarding the supernatant and re-suspending the protoplasts in 10 ml of protoplast regeneration medium-top layer (PRM-T medium) (Table 14) previously preheated to 45 ºC. Finally, this mixture was distributed in plates of protoplast regeneration medium-bottom layer (PRM-B medium) (Table 14) containing cellophane discs, where the regeneration of the protoplasts would take place later on.

Table 13. Composition of MMM and PEGcms solutions. MMM (for 100 ml) PEGcms (for 10 ml)

MgCl2·6H2O 6.1 g Ca(NO3)2·4H2O 0.2360 g Mannitol 8 g HEPES 0.0476 g MES 0.2 g Mannitol 0.7280 g PEG6000 4 g pH adjusted to 5.6 with KOH (1 pH adjusted to 7.5 with KOH (1 M), and left at M). Sterilized by filtration 0.22 or room temperature all night. Sterilized by filtration 0.45 µm Millipore filters. 0.22 or 0.45 µm Millipore filters.

Table 14. Composition of different protoplast regeneration media used in Physcomitrella transformation. PRM-B (500ml) PRM-T (100ml) PRM-L (5ml) Stock B (100x) 5 ml 1 ml 200 µl Stock C (100x) 5 ml 1 ml 200 µl Stock D (100x) 5 ml 1 ml 200 µl Ammonium 5 ml 1 ml 200 µl Tartrate 0.5M (100x) Trace elements 0.5 ml 0.1 ml 20 µl (1000x) Mannitol 30 g 6 g 1.2 g Agar (plant 0.55% 0.4g (0.4%) Agar, Duchefa)

CaCl2 (1 M) 200 µl Solution sterilized in Solution sterilized in Final volume autoclave. Once autoclave. Once taken to 20ml. sterilized, 5 ml of 1 M sterilized 5 ml Sterilized by

CaCl2 is added. CaCl2 1 M is added. filtration through For each DNA The regeneration of 0.22 or 0.45 µm transformed, 3 plates of protoplasts requires Millipore filters 20-25 ml were prepared a higher calcium covered with cellophane concentration (10 paper discs. mM ) than normal growth (1 mM)

3) Selection of transformants: Passed 5 d of incubation, the cellophane discs with the regenerated protoplasts were transferred to BCDNH4 plates, containing the suitable antibiotic and left for 2 weeks. Later on, successive transfers of protoplasts to BCDNH4 medium plates for 2 weeks followed by another transfer for 2 weeks to BCDNH4 medium containing the suitable antibiotic were performed after the appearance of resistant colonies of Physcomitrella to the antibiotic. A small sample of these colonies was taken to extract 77

genomic DNA for PCR analysis to verify the correct integration of the gene disruption construction. The following table details the different media used throughout the transformation protocol of protoplasts.

3.5. Construction of Physcomitrella knock-out mutants by genetic replacement (homologous recombination) A series of constructs has been designed to disrupt the different genes of interest that are detailed next. All of these constructs have two homologous fragments of DNA to the 5’ and 3’ regions flanking the gene of interest to knocked out, and a cassette of resistance to antibiotic that will replace the gene.

3.5.1. Design of the construction for the disruption of PpCHX1 and PpCHX2 genes The pTN186 vector (Table 9, and http://moss.nibb.ac.jp/) was used to construct the PpCHX1:Hyg knockout fragment (Fig. 23A). In this fragment, the hygromycin resistance cassette, which contained the aph4 gene under the control of the promoter and terminator of the 35S gene, was flanked by two fragments of the non-codifying 5´ and 3´ regions of PpCHX1 gene. These fragments were amplified by PCR using several pairs of primers (Table 15) that included the restriction enzymes sequences to direct the cloning into the corresponding sites of the pTN186 vector polylinker. The 5´ fragment extended from nucleotide positions -975 to 80 and was inserted between the KpnI and HindIII sites. The 3´ fragment extended from position 3034 to 4103 and was inserted between the restriction sites XbaI and SacI of the pTN186 vector polylinker. The ∆Ppchx1 knockout lines were generated by transforming P. patens protoplasts with 25 µg of the linear fragment obtained by digesting the pTN186:Ppchx1 plasmid with the KpnI and SacI restriction enzymes. Stable antibiotic-resistant clones were selected after two rounds of incubation in BCDAT medium supplemented with 30 µg ml-1 of hygromycin (Invitrogen, the USA). The p35S-Zeo vector (Table 9, and http://moss.nibb.ac.jp/) was used to construct the PpCHX2:Zeo knockout fragment (Fig. 23B). In this fragment, the zeocin resistance cassette, which contained the ble gene under the control of the promoter and terminator of the 35S gene, was flanked by two fragments of the non-codifying 5´ and 3´ regions of 78

the PpCHX2 gene. These fragments were amplified by PCR using primers (Table 15) that included the restriction enzymes sequences to direct the cloning into the corresponding sites of the vector polylinker. The 5´ fragment extended from nucleotide positions -1066 to 314 of the ATG and was inserted between the XhoI and EcoRI sites. The 3´ fragment extended from positions 2081 to 3114 and was inserted between the restriction sites SacII and NotI. The ∆Ppchx2 knockout lines were generated by transforming P. patens protoplasts with 25 µg of the linear fragment obtained by digesting the p35S-Zeo:Ppchx1 plasmid with the XhoI and NotI restriction enzymes. Stable antibiotic-resistant clones were selected after two rounds of incubation in BCDAT medium supplemented with 50 µg ml-1 of zeocin (Invitrogen, the USA). Double Mutants ∆Pphak1 ∆Ppchx2 were constructed, implementing the same strategy described above, by disrupting the PpCHX1 gene of the ∆Pphak1 mutant previously obtained in our laboratory (Garciadeblás et al. 2007).

Table 15. Primers used for the generation of different constructions for gene disruption. Primers Sequence 5´-3´ Observations

PpCHX1-5’-KpnI GGTACCTCTGTCCCGCCTCTCATATATAAC Cloning of the 5´ fragment in pTN186 for PpCHX1 gene PpCHX1-5’-HindIII AAGCTTAGCAAAATGAACGGGCACATCTCC knock-out

PpCHX1-3’-XbaI-2 TATTCTAGATATTTTGGTTGGCCGATCGAG Cloning of the 3´ fragment in pTN186 for PpCHX1 gene PpCHX1-3’-SacI-2 TTGAGCTCACAGATCCATTCCATCAAAACA knock-out

Cloning of the 5´ PpCHX2-5’EcorI2 TGAATTCTATCCCCTTGCCACACGCC fragment in p35S- Zeo for P p CH X 2 gene knock-out PpCHX2-5’XhoI2 ACTCGAGAATTTCATCACCTGCCTAC

Cloning of the 3´ PpCHX2-2-3’(1680) AACTGACAAACGAGCCGCAATGATC fragment in p35S- Zeo for P p CH X 2 PpCHX2-3’ NotI-2 GCGGCCGCGACCAATCCACTTCACTAAA gene knock-out

3.5.2. Screening and analysis of putative Physcomitrella knockout clones by PCR The screening of putative Physcomitrella knockout clones was performed by PCR on genomic DNA purified from transformed plants. Four independent PCR reactions were performed to confirm that both 5’ and 3’ insertion sites were sequentially correct. For both sites, one primer corresponded to a genomic sequence fragment outside the knockout construction and was used in both PCR reactions (Table 16); of the other two primers, one was specific for the marker and the other for the wild-type gene (Fig. 23). Clones in which the sequences of the knockout insertions were correct and the fragments of the wild-type gene could not be amplified were selected. Four ∆Ppchx1- (1-4 ), seven ∆Ppchx2-(1-7), and six ∆Pphak1∆Ppchx2-(1-6 ) lines were isolated and

studied; however, repeated experiments were performed only with ∆Ppchx1-1, ∆Ppchx2-1, and ∆Pphak1∆Ppchx2-1 plants. The total DNA of the possible mutants was obtained following the CTAB method protocol (see section 3.2.2) or by “Green PCR” method (Schween et al. 2002) that allows processing a high number of samples, as follows: “Green PCR” method In order to verify the integration of our designed construction described earlier in the genome of Physcomitrella, a fast and simple method of PCR was used. A small sample of fresh Physcomitrella colonies of approximately 1 week was placed in an eppendorf tube containing 30 µl of Taq polymerase 10x reaction buffer. The sample was then crushed with a sterile wood tooth pick for 2 min until the buffer color become green. Next, the sample was incubated for 10 min at 68 ºC and then centrifuged for 5 min at 4 ºC. For PCR analysis, 5 µl of the supernatant was used in a final volume of 50 µl. The obtained PCR product of was verified by electrophoresis in agarose gels (Schween et al. 2002, modified by T. Nishiyama, http://moss.nibb.ac.jp/).

Table 16. Primers used for the verification of Physcomitrella knockouts. Primers Sequence 5´-3´ Observations Anneals to 35S promoter of PM35S-1R {2D in p35S-Zeo and pTN186 GCCCTTTGGTCTTCTGAG fig. S2-A&B} cassettes of resistance to antibiotics Anneals to 35S terminator 35SPA-1 {1C in fig. of p35S-Zeo and pTN186 AGGGCCATGAATAGGTCT S2-A&B} cassettes of resistance to antibiotics PpCHX1-ATG3 {1A 5´ end verification of GGCGTGCAAAACTATGTCGG & 1D in fig. S2- A} PpCHX2 knock-out PpCHX1-STOP {2B 3´ end verification of TTAGCCAGCCAAGACCTTCGCA & 2C in fig. S2- A} PpCHX2 knock-out PpCHX1-ATG3 {1B GGCGTGCAAAACTATGTCGG Amplification of knocked- in fig. S2- A} out fragment of PpCHX2 PpCHX1-1R {2A in AAAGCTCCCTCTGGTGCAAT gene fig. S2- A} PpCHX2-5’-G1 {1A 5´ end verification of CAGCGAACGAACAAACCAAC & 1D in fig. S2- B} PpCHX1 knock-out PpCHX2-3’-GR {2B 3´ end verification of CAAACTGCCAAAACATATGA & 2C in fig. S2- B} PpCHX1 knock-out PpCHX2-4R {2A in CAATGACCCGAGGCTGTTTC Amplification of knocked- fig. S2- B} out fragment of PpCHX1 PpCHX2- (1680) {1B AACTGACAAACGAGCCGCAATGATC gene in fig. S2- B}

3.6. Design of gene fusions with GFP to determine the subcellular localization of the transporters Localization of the PpCHX1-GFP and PpCHX2-GFP fusion proteins in P. patens was performed by transient expression in protoplasts. The PpCHX1-GFP and PpCHX2-GFP genes were cloned under the control of the PpACT5 promoter containing a large intron in the 5’ untranslated region (5’UTR). The expression plasmid, pRHACT5, was constructed from the PCR2.1-Topo vector. For this purpose, the 5’UTR-PpAct5 fragment was amplified by PCR using the primers published by Weiss et al. (2006): number 13 as the forward primer and number 14 as the reverse primer; the

latter was modified to contain a BamHI cloning site at the 3’ end of the primer. Then, a BamHI-HindIII fragment from the pMF6 vector containing the GFP:nos-3’ terminator (Rubio-Somoza et al. 2006) was cloned downstream of the PpACT5 promoter. Finally, the PpCHX1 and PpCHX2 cDNAs were cloned in frame into the BamHI site at the 5’ of the GFP gene. Experiments of co-localizations were performed with organelles marker used in others plants (Nelson et al. 2007), http://www.bio.utk.edu/cellbiol/markers/). The tonoplast marker was achieved by fusing the coding region of the YFP to the C- terminus of γ-TIP, an aquaporin of the vacuolar membrane (Saito et al. 2002). The Golgi localization was based on the fusion of the YFP with the cytoplasmic tail and transmembrane domain (first 49 aa) of GmMan1, soybean α-1,2-mannosidase I (Saint- Jore-Dupas et al. 2006). The resulting constructs were used for transient expression in P. patens protoplasts. After transformation, the protoplasts were kept in the dark for 24 h in BCDNH4 medium (Table 7) supplemented with 6% mannitol and 5% glucose, followed by cultivation in the same medium for 3–4 d under normal growth conditions. Z series images of protoplasts were obtained on a Leica TCS SP8 confocal microscope (LeicaMicrosystems, Wetzlar, Germany). The GFP signal and chlorophyll autofluorescence were collected simultaneously under the laser excitation lines of 488 nm and 633 nm, respectively. To rule out any crosstalk between GFP and YFP, the YFP signal was collected sequentially under the laser excitation line of 514 nm. Images were processed using the LAS AF Lite 3.1.0 (Leica Microsystems, Wetzlar, Germany). In table 17, the primers used for the different GFP constructs carried out are detailed.

Table 17. Primers used for the fusion of PpCHX proteins with GFP. Primers Sequence 5´-3´ Observations

PpCHX1-ATGBamHI2 CGGATCCTAAAAAATGGCGGACGCTGTGGC Cloning of the PpCHX1 in phase TTGGATCCGCACTCTGTGGTCAGGAGAAGAC PpCHX1-NoStop-B a m H I with GFP gene AA

PpCHX2-ATG-B a m H I GTGGATCCTACACAATGGCGACCGGAAATGC Cloning of the PpCHX2 in phase PpCHX2-NoStop-B a m H I TTGGATCCGTACATGAGGCGCTGACTTC with GFP gene

3.7. Functional complementation “drop tests” of yeast and bacteria mutants and yeast growth curves To analyze the function of transporter proteins codified by the identified genes of Physcomitrella, a functional reconstruction approach of Na+ or K+ transport in yeast or bacteria mutants defective in their Na+ or K+ transport was carried out. Complementation tests in yeast strains defective in Na+ and/or K+ transport were carried out on solid media plates of YPD or Arginine phosphate (Table 5) supplemented with the indicated concentrations of Na+ and/or K+. Cellular suspensions were prepared from fresh cultures of the transformed yeast strains. Then, 3-5 serial 1:10 dilutions were made from these suspensions. Of each dilution, a 5 µl drop was applied in the surface of the plates and left to dry in the laminar air flow chamber. The plates were incubated during 3 or 4 d at 28 ºC. The complementation tests of transformed bacteria mutant TKW4205 were done in minimal medium MM (Table 3) supplemented with different concentrations of K+ or Rb+ in addition to different concentrations of arabinose with the purpose of regulating the level of expression of the transporters. The preparation of bacterial suspensions was realized as described above for yeast. The time course yeast growth curves assays were monitored by recording the absorbance at 540 nm in a Microbiology Reader Bioscreen C workstation (Growth Curves Oy, Finland), which enables many samples of different mutants under different conditions to be run at the same time.

3.8. Cation contents and cation uptake experiments in yeast To carry out Rb+, K+, or Na+ influx tests in yeast, cultures of different transformed yeast clones were grown in 100 ml of AP medium supplemented with the convenient concentrations of K+ indicated for each test, and left over night in agitation at 28 ºC. On the following day, the cells were harvested by centrifugation and washed two times with H2Om q (treated in 185 Milli-Q-Extra Millipore, Molsheim, France). Finally, the cells were re-suspended in arginine phosphate medium supplemented with the convenient concentrations of the adequate cation indicated for each test. Samples of 5 or 10 ml were taken alternatively at different times and filtered through 0.8 µm Millipore AAWP filters (Molsheim, France) with the help of a vacuum pump where the

cells were thoroughly washed with 20 mM MgCl2 solution for 5 min. The extraction of the cellular Rb+, K+, or Na+ was realized with a solution composed of 0.2 M HCl and 10 mM MgCl2 for 8 h. Finally, the cations concentrations of the samples were measured by an atomic emission spectrophotometer (Model 2380, Perkin-Elmer Norwalk, CT, the USA).

3.9. Cation fluxes tests in Physcomitrella plants In order to have sufficient amount of Physcomitrella culture for different experiments, 3-5 g of fresh weight was grown in BCDNH4 medium for 2-3 weeks. Under these conditions, Physcomitrella displayed contents of 1800-2500 nmoles mg-1 for K+ and 15-60 nmoles mg-1 for Na+ in plant dry weight. For the cation influx tests in millimolar range, 3-5 g samples of fresh weight were taken from the different mutants and lines of Physcomitrella of approximately and were transferred to 300 ml of liquid KFM medium containing the convenient cation concentrations and pHs indicated in each case. At different times, samples of the plant suspension were taken with the help of tweezers and placed into a washing solution containing MgCl2 1 mM and left for 1 min. Passed that time, the samples were filtered through a 0.8 µm Millipore filter. For the quantification of internal cation contents in Physcomitrella, the samples were dried in a stove to 68 ºC and then weighed. The internal cations were extracted by adding to the plant samples an extraction solution containing 0.1M HCl and 10 mM MgCl2. The determination of cation content was realized in a spectrophotometer of atomic emission PerkinElmer AAnalyst 200/400. Alternatively the quantification of the internal cation contents of Physcomitrella was realized weighing the washed samples (fresh weight) and extracting directly with the extraction solution described above. In order to refer the collected data to dry weight, we multiplied it by a factor of 17.25, a value previously verified in different tests representing the difference existing between dry weight and fresh weight. On the other hand, cation uptake tests at micromolar concentrations of K+, Na+ or Rb+ were carried out by determining the depletion of these cations in the medium containing Physcomitrella cultures previously starved for K+ for 2 weeks in KFM medium. The K+ starved culture of Physcomitrella was transferred to 30 ml of KFM and the desired amount of the corresponding cation (usually 100 µΜ RbCl, NaCl or KCl)

was added to the medium to start the experiment. Throughout the experiment, 200 µl samples were taken from the medium and diluted in 1 ml of H2Om q for afterward measurement of the cations by atomic emission spectrophotometer. At the end of the experiment, the plant material is collected and dried in a stove at 68 ºC for 24 h to determine the dry weight used in the experiment. Throughout the period of plants culture previous to experiments, several samples of plant material were taken and inoculated in LB medium in order to detect any possible yeast or bacterial contaminations that could alter the results.

3.10. Software and bioinformatic Tools In order to design the different cloning strategies, described earlier, together with the analysis of restriction enzyme sites, sequences and any other routine manipulation of DNA and protein sequences in electronic format, DNA Strider 1.2 program was used (C.Marck, Dept. Biologie Cellulaire ET Moléculaire, CEA, France). For primer analysis and design, Oligo 4 software was used throughout this study (Molecular Biology Insights, Inc., CO, USA). The analysis of PCR amplified sequences was carried out by using Sequencher 4.0 program (Gene Codes Corporation, Ann Arbor, MI, USA). Multiple alignments of sequences were done using the ClustalW 2 tool of the European Bioinformatics Institute (http://www.ebi.ac.uk/Tools/msa/clustalw2/). The Phylogenetic trees were constructed and visualized by Treeview 1.6.6 software (R.D.M. Page, University of Glasgow, UK). The DNA and protein CHX sequences of P. patens were blasted against the sequences available at National Center for Biotechnology Information (NCBI) databases (http://blast.ncbi.nlm.nih.gov) using the BLAST tool (Altschul et al., 1990). The search for existing genes in the Physcomitrella genome was done through online pages of JGI Joint Genome Institute (http://genome.jgi- psf.org/Phypa1_1/Phypa1_1.home.html) and Phytozome databases (http://www.phytozome.net). Finally, graphs and statistical analysis were carried out using Microsoft Office Excel 2007 software (Microsoft, USA).

4. RESULTS

4. RESULTS 4.1. The CHX transporters of P. patens At the beginning of the research of this thesis work, shortly after the publication of the complete genome sequence of Physcomitrella patens (Rensing et al. 2008) by the Doe Joint Genome Institute (JGI), we carried out a BLASTP search in the corresponding database website (http://genome.jgi- psf.org/Phypa1_1/Phypa1_1.home.html, accessed in November, 2009) using the Arabidopsis thaliana AtCHX23 protein (AT1G05580.2) as a query sequence to identify the number of CHX genes in Physcomitrella patens genome. That search resulted in the identification of two CHX genes; PpCHX1 (estExt_fgenesh1_pm.C_1230004, also known as PHYPADRAFT_107047 in NCBI database) and PpCHX2 (estExt_gwp_gw1.C_1620028, also known as PHYPADRAFT_191426 in NCBI database). Previous searches carried out by our group in the aforementioned database failed to identify any homologous sequence to the AKT K+ inward rectifying channel gene, despite the certainty of the existence of at least two AKT genes (PpAKT1-2) in P. patens, before its genome even being sequenced and published (Garciadeblás et al. 2007). Knowing this beforehand, these search results of P. patens having only two CHX genes were not considered definitive and were taken with care. Later on, our skepticism was confirmed to be correct after the publication of a newly revised version of the genome sequence in the Phytozome database (http://www.phytozome.net, accessed in 2011). Indeed, the new BLASTP search carried out in that database identified four CHX genes (PpCHX1-4) in the genome of P. patens, instead of just two; PpCHX1 (Pp1s123_30V6.1), PpCHX2 (Pp1s162_22V6.1), PpCHX3 (Pp1s270_2V6.1), and PpCHX4 (Pp1s375_26V6.1). Nevertheless, and surprisingly enough until this moment, the sequences of PpCHX3 and PpCHX4 are not identified when the same BLASTP query is carried out in NCBI databases (http://www.ncbi.nlm.nih.gov, accesed in February 2013). A phylogenetic study of these transporters has been recently published (Chanroj et al. 2012), and this report, together with the Phytozome database sequences of PpCHX genes, were used as the starting point of our study. The structure of introns in the PpCHX1-4 genes is a rather confusing issue, as in the case of PpCHX2 gene which will be discussed later on in this chapter. Nevertheless, the genomic sequences revealed that PpCHX1 and PpCHX2 have 3 and 2 introns

respectively, while PpCHX3 and PpCHX4 are similar to each other both having 7 introns (Fig. 4).

Fig. 4. Exon-intron structure of the four P. patens CHX genes.

The genes sequence analysis revealed different CDS (coding DNA sequence) lengths for PpCHX1-4; 2544, 2484, 2667, and 2856 nucleotides, respectively. PpCHX1- 4 cDNA sequences could encode proteins sequences presenting hydrophobic N-terminal halves of 424, 429, 422, and 427 residues, respectively, and hydrophilic C tail of 420, 398, 467, and 525 residues, respectively. Also, the four PpCHX1-4 protein sequences have 12, relatively coinciding, transmembrane helices as predicted by the TMHMM2 algorithm (http://www.cbs.dtu.dk/services/TMHMM-2.0) and shown in figure 5. None of the 4 CHX protein sequences contained a predicted organelle targeting sequence according to SignalP 4.1 program (http://www.cbs.dtu.dk/services/SignalP).

Fig. 5. Pp C HX1 -4 translated amino acid sequences showing the different lengths of sequences, where the vertical bars represent the relatively coinciding transmembrane helices.

This relatively long length of the C tail is a mysterious characteristic of CHX transporters, since they do not seem to contain any conserved motifs (Chanroj et al. 2012). However, many conserved residues do exist in the C tail and can be identified when aligning PpCHX proteins with other CHX proteins of flowering plants such as Arabidopsis thaliana (Fig. 6), suggesting that these regions might carry out regulatory functions.

MI PpCHX1 ----MADAVACKTMSATSNGVWQGDVPVHFALPLLIVQIVLVLAITRALAFVLKPLKQPR 56 PpCHX2 MATGNATVTTCKTMPASSNGVWQGDIPIHFALPLLIIQICIVLTITRVLAALFKPLKQPR 60 PpCHX3 -----MASENVTDVSVTSSGVVAGDNPLHHTLPLLIIQVMVIITVSRCVAVLLRPLKQPR 55 PpCHX4 -----MTSNQTSDVSVTSRGVLAGDDPLHHALLLLIIQVIIIICLTRFLALLLRPLKQPR 55 AtCHX18 MAT-NSTKACPAPMKATSNGVFQGDNPIDFALPLAILQIVIVIVLTRVLAYLLRPLRQPR 59 AtCHX20 ------MPFNITSVKTSSNGVWQGDNPLNFAFPLLIVQTALIIAVSRFLAVLFKPLRQPK 54 : .:* ** ** *:..:: * *:* ::: ::* :* :::**:**: MII MIII PpCHX1 VVAEIIGGILLGPSAFGRNKDYLHTIFPHESVIILEVFADMGLLFFLFMVGLELDMTQIR 116 PpCHX2 VIAEVVGGILLGPSALGKNTAYIANIFPKQSVIILEVFAQMGLIFFLFMVGLELDIRQIR 120 PpCHX3 VVAEILGGILLGPTAFGHIPGFTKNIFPESSLPVLETVAEVGLIFFLFLVGLELDTKQIR 115 PpCHX4 VVAEILGGIILGPTGFGSIPGITDTIFPESSLTVLDTAANVGLIFFLFLVGLELNIKTIV 115 AtCHX18 VIAEVIGGIMLGPSLLGRSKAFLDAVFPKKSLTVLETLANLGLLFFLFLAGLEIDTKALR 119 AtCHX20 VIAEIVGGILLGPSALGRNMAYMDRIFPKWSMPILESVASIGLLFFLFLVGLELDLSSIR 114 *:**::***:***: :* :**. *: :*: *.:**:****:.***:: : MIV MV PpCHX1 KTGKQAMSIAAAGITLPFVAGVGVSFVLHLTIAPEGA---FGPFLVFMGVAMSITAFPVL 173 PpCHX2 RTGFQALVISAAGIAVPFSTGVGVSFVLLNTIAGDVK---FGPFVVFMGVAMSITAFPVL 177 PpCHX3 RSGVMTLWLSAAGIILPFGLGAVAAFIIFKLQNSLH-HPNFGAFVLFLGVALSVTAFPVL 174 PpCHX4 KSGVISLWMAAAGIIFPFGLGSIVACLISKLLPQMDSHHSFGVFTMFLGVALSITAFPVL 175 AtCHX18 RTGKKALGIALAGITLPFALGIGSSFVLKATISKGVN---STAFLVFMGVALSITAFPVL 176 AtCHX20 RSGKRAFGIAVAGITLPFIAGVGVAFVIRNTLYTAADKPGYAEFLVFMGVALSITAFPVL 174 ::* :: :: *** .** * : :: * :*:***:*:****** MVI PpCHX1 ARILAERKLLTTEVGQLAMSAAAVNDVVAWVLLALAVALSGSG------RSPAIVAWVL 226 PpCHX2 ARILAERKLLTTEVGQLAMSVAAVDDVVAWCLLALAVALTGTN------TKPSVVAWVL 230 PpCHX3 ARILTERKLLHTDIGQMAMAAAAINDVVAWVLLALAVAITSSG------SDPLVALWVL 227 PpCHX4 ARILAERKLLNTEIGQMAMSAAAINDIVAWILLALALALTNSG------STPVVAIWVL 228 AtCHX18 ARILAELKLLTTEIGRLAMSAAAVNDVAAWILLALAIALSGSN------TSPLVSLWVF 229 AtCHX20 ARILAELKLLTTQIGETAMAAAAFNDVAAWILLALAVALAGNGGEGGGEKKSPLVSLWVL 234 ****:* *** *::*. **:.**.:*:.** *****:*::... * : **: 91

MVII MVIII PpCHX1 LCGIAFCLAIFLVVQPCMQWVAHR-SPDNEPVKEYIVALTLLCVLVAGFCTDAIGVHSIF 285 PpCHX2 LTGIAFIITMFVVVQPVMRWVATR-SADNEPVKEILVCLTFAGVLIAAFTTDLIGIHAIF 289 PpCHX3 LLGTVFIVFMLLVISPITYALAHH----SEPATESVIAMTLMLVLGAAFITDLIGIHVIF 283 PpCHX4 LLGVAFSAFMFVVVSPVMYALANY----RELPPEPVVAMTLVIVLGSAFFSDLIGIHVIF 284 AtCHX18 LSGCAFVIGASFIIPPIFRWISRR-CHEGEPIEETYICATLAVVLVCGFITDAIGIHSMF 288 AtCHX20 LSGAGFVVFMLVVIRPGMKWVAKRGSPENDVVRESYVCLTLAGVMVSGFATDLIGIHSIF 294 * * * .:: * :: : * :. *: *: ..* :* **:* :* MIX PpCHX1 GAFLFGLVIPKEGPFA----AALVEKLEDFVSILLLPLYFASSGLKTNIGAIHSAQSFGL 341 PpCHX2 GAFLFGLIVPKDGPFA----VALVEKIEDFISILMLPLYFASSGLKTNIGAIKTGQSFGL 345 PpCHX3 GAFICGLIIPKDGPFT----AILIEKVEDYVSVLFLPLYFAASGLKTHLSAINNGTAVLI 339 PpCHX4 GAFICGLIVPKDGSFAGTLTSTMIEKVEDYVSVLFLPLYFAISGLKTHLSEVNNGTAVLI 344 AtCHX18 GAFVVGVLIPKEGPFA----GALVEKVEDLVSGLFLPLYFVASGLKTNVATIQGAQSWGL 344 AtCHX20 GAFVFGLTIPKDGEFG----QRLIERIEDFVSGLLLPLYFATSGLKTDVAKIRGAESWGM 350 ***: *: :**:* * ::*::** :* *:*****. *****.:. :. . : : MX MXI PpCHX1 LVLVISVACLGKILGTFAAAKACRVDARKALTLGILMNTKGLVELIVLNIGLDRGVLNSE 401 PpCHX2 LVLVIAVACFGKMCGVFLAATASKVNPRKALTLGVLMNTKGLVELIVLNIGKDRGVLNEE 405 PpCHX3 LFLVIASACVGKVLGTFIVAKVWGVGSRKAIALGFLMNTKGLVELIILNIGLSKGVLNEE 399 PpCHX4 LFMVIATACIGKVMGTLIVAKIWGVENSKALALGFLMNTKGLVELIVLNIGLSKRVINQE 404 AtCHX18 LVLVTATACFGKILGTLGVSLAFKIPMREAITLGFLMNTKGLVELIVLNIGKDRKVLNDQ 404 AtCHX20 LGLVVVTACAGKIVGTFVVAVMVKVPAREALTLGFLMNTKGLVELIVLNIGKEKKVLNDE 410 * :* ** **: *.: .: : :*::**.***********:**** .: *:*.: MXII PpCHX1 TFAIMVLMALFTTFMTTPLVMAIYKPARNPTP---YTRRTLEMEDSKDDLRILSCVHGMK 458 PpCHX2 TFAIMVLMALVTTFMTTPLVMALYKPARNPIP---YNRRKLAMEDSKDDLRILSCVHGMK 462 PpCHX3 LFAIMVIMTITTTFITTPVVMWLYKPACDIPP---YKRRTINSGDDRDELRMLFCLLGSW 456 PpCHX4 LFAIMVVMALVTTFITTPVVMWLYTPARDIPP---YKRRSIGSDDDKDELRMLLCPVGEW 461 AtCHX18 TFAIMVLMALFTTFITTPVVMAVYKPARRAKKEGEYKHRAVERENTNTQLRILTCFHGAG 464 AtCHX20 TFAILVLMALFTTFITTPTVMAIYKPARGTHR--KLKDLSASQDSTKEELRILACLHGPA 468 ***:*:*:: ***:*** ** :*.** . . . :**:* * *

PpCHX1 NVAAMINLTEATRGMRKR-TLRLYILHLMELSERTSAIMIVQRARRNGRP-FFNQSKHSD 516 PpCHX2 NVPAMINLTEGTRGIRKR-ALRLYILHLMELSERTSAIMIVQRARKDGRP-FFNQRKSAE 520 PpCHX3 NINSMMKMAEITGGEDYK-NFRAYVLHLVEYSERLSTIQMSKFSKRESEDGAGNE--GNT 513 PpCHX4 NIPGMVNIIEITRGKHHK-SLRAYVLHLIECSERLSSIRMSTFSRRNSRDNFMNEEHGNT 520 AtCHX18 SIPSMINLLEASRGIEKGEGLCVYALHLRELSERSSAILMVHKVRKNGMP-FWNRRGVNA 523 AtCHX20 NVSSLISLVESIR-TTKILRLKLFVMHLMELTERSSSIIMVQRARKNGLP-FVHRYRHGE 526 .: .::.: * : : :** * :** *:* : :::. :.

PpCHX1 NKDQIVAAFETYEQLSKVTVRPMTAISGFDDMHEDICATAADKRTALIMLPFHK------570 PpCHX2 SRDQIVAAFETYGHLSKVTVRPMTAISNFEDMHEDICATATDKRAAMIILPFHK------574 PpCHX3 ELDAIEVAFQKSSQLTKVKVKTEVAISAFHNMHIDVCNAAYSNRVNFLLLPFHR------567 PpCHX4 EVEMVEVAFQSYGKLGRVQVKTAVAVSAFRNMHVDVCNIACSNRVNFLLLPLHM------574 AtCHX18 DADQVVVAFQAFQQLSRVNVRPMTAISSMSDIHEDICTTAVRKKAAIVILPFHK------577 AtCHX20 RHSNVIGGFEAYRQLGRVAVRPITAVSPLPTMHEDICHMADTKRVTMIILPFHKRWNADH 586 . : .*: :* :* *:. .*:* : :* *:* * ::. :::**:*

PpCHX1 ---SPRLDG-----HFDST-PGFRTVNQKVLKHAPCSVAILIDRGVG----GSAQVPSSN 617 PpCHX2 ---TQRLDG-----QFDTTAPGFRLVNQKVLQHAPCSVAILIDRGVG----GSAQVAPNN 622 PpCHX3 ---RRRFDK-----NFETVASGLKEVNMKVFQDPPCSVGLLVDNGFG----DHAATTPG- 614 PpCHX4 ---RRSHDG-----NFGTMFPELKKLNMKILRDAPCSVGLLVDNGLG----GPTATPPTN 622 AtCHX18 ---HQQLDG-----SLETTRGDYRWVNRRVLLQAPCSVGIFVDRGLG----GSSQVSAQD 625 AtCHX20 GHSHHHQDGGGDGNVPENVGHGWRLVNQRVLKNAPCSVAVLVDRGLGSIEAQTLSLDGSN 646 * . : :* ::: ..****.:::*.*.*

PpCHX1 VDHNVVVYFFGGPDDREALAYGFRMAEHPGVKLHVIRFLSHSVVMDDGHGGLASVGSEVS 677 PpCHX2 VDHKVVVYFFGGQDDREALAYGLRMAEHPGIQLHVIRFLNSNIDTTIEIDGQHSESLGLS 682 PpCHX3 SCQHILVLFFGGPDDRESLMLGRRMLKNDGVKLTVIQFVVQDPKRNHLCSIRRVSSRTLK 674 PpCHX4 YSQHIYVLFFGGPDDREALMLARRMLQHGGIKLTVIQIVIQGLVHHHLGRIRRISSRMLR 682 AtCHX18 VSYSVVVLFFGGPDDREALAYGLRMAEHPGIVLTVFRFVVS------PERVG 671 AtCHX20 VVERVCVIFFGGPDDRESIELGGRMAEHPAVKVTVIRFLVRETLRS------TAVTL 697 : * **** ****:: . ** :: .: : *::::

PpCHX1 EIGKTEVSDTRFQFAMHGLD------QNRQRELDEE 707 PpCHX2 KLHHSSL-DKGYHIATGALD------QDQERKLDEA 711 PpCHX3 SHLQTESAAKRGWSALKHVARDVVDFFAAPWVKTKECGRAKQND------LSPESSGKMD 728 PpCHX4 SHLPKKPPAWRRWEAWNWLVCEVVNFFKAPWVKPKKCVKPNQNSEAEKTCLTTDSTGDTN 742 AtCHX18 EIVNVEVSNNNNENQS------VKNLKSDEE 696 AtCHX20 RPAPSKGKEKNYAFLTTNVD------PEKEKELDEG 727 . : .

PpCHX1 ALGHVRRRQAS------EDGRVTYVEMQVSEPLEEVVRLSSSR------EHDIIL 750 PpCHX2 ALDGIRKGEKGKVDADEVHSKVSWEECRVADPFEAVVQAAIAG------DHNIIL 760 92

PpCHX3 VTDATETQPDGETHVLVDDILLQKERDMNVQALAPILAAATAARKEFR--NRPMNNASLE 786 PpCHX4 VADAIMRNSNGEIHVVMKDILSESERIKDMQALAPILATATKANQEEVGDTGDRNGESLQ 802 AtCHX18 IMSEIRKISSVDESVKFVEKQIENAAVDVRSAIEEVRRS------NLFL 739 AtCHX20 ALEDFKSKWKE------MVEYKEKEPNNIIEEILSIGQSK------DFDLIV 767 . : : :

PpCHX1 VGRSRRPTPFLERFRRKHAE-----YAELGPIGDALMAP--QVRASVLVFQQHDHVLADP 803 PpCHX2 VGRSRRPTAFVGSMVHRHPE-----YTELGPLGEALMAP--EVRASVLVFQQYD-----P 808 PpCHX3 TGQGPPTTAVEELQSEVSCRNLTLRVVETQNLKKSVLDTVKSAEPNGLIITGLHLHEHST 846 PpCHX4 SNRSLHTTN-PGLQSEIACRNLTLRVIETQKLEESILSVVNSAESNGLIITGMHLHENSP 861 AtCHX18 VG--RMPGGEIALAIRENSE-----CPELGPVGSLLISPESSTKASVLVIQQYNGTGIAP 792 AtCHX20 VGRGRIPSAEVAALAERQAE-----HPELGPIGDVLASSINHIIPSILVVQQHN-KAHVE 821 . . . . * : . : .. *:. .

PpCHX1 LPNTSETEAVKELQTFPSSKELVDRKGDVQKIDLSSPDHRV------844 PpCHX2 LLDLDAAASAKSAKSAPHV------827 PpCHX3 IKQSGDFIPVEDYGLGPLGNYLVS-KHLHMHTSLLVVKQHISA------888 PpCHX4 VLQSYAFPMIEDHGLGLVGNFLVSNKHPHMEASLLVVKHYGPSQDTVLTSSIPADSSANL 921 AtCHX18 DLGAAETEVLTSTDKDSD------810 AtCHX20 DITVSKIVSESSLSINGDTNV------842 .

PpCHX1 ------PpCHX2 ------PpCHX3 ------PpCHX4 LSVANVEVGSSTFTGPSDINDSGSSQNKLG 951 AtCHX18 ------AtCHX20 ------Fig. 6. CHX complete sequence alignment. Sequence alignment was performed using clustalW2 (http://www.ebi.ac.uk/Tools/msa/clustalw2/). Highlighted in grey and marked with M indicates transmembrane region for PpCHX1-4 which were calculated from the TMHMM tool (CBS prediction server, http://www.cbs.dtu.dk/services/TMHMM/) and for the AtCHX18 and AtCHX20 as indicated by Chanroj et al. 2011. In red, residues conserved in all CHX from Arabidopsis including the Aspartic in the MVI for the cation binding site. Highlighted in black, the sequence from the putative intron of PpCHX2.1.

Additional searches on Phytozome database also revealed that the similarity between PpCHX1 and PpCHX2 is relatively high (76.7%), while much lower when comparing PpCHX1 with either PpCHX3 (47.9%) or PpCHX4 (51.2%). PpCHX3 and PpCHX4, on the other hand, are much more similar to each other (68.9%). A subsequent BLASTP search in the NCBI databases (http://www.ncbi.nlm.nih.gov) using PpCHX1 protein as a query sequence revealed that, among the sequenced plant species in the database, the proteins with highest identity to PpCHX1 (58-60%) were the three CHX proteins of the non-vascular club moss Selaginella moellendorffii (SmoCHX1-3), followed by a 54% identity of the higher-plant black cottonwood Populus trichocarpa PtrCHX17 (POPTR_0001s10170.1). This was also the case for BLASTP searches using PpCHX2-4 as a query. Regarding the percentage of identity to CHX proteins of Arabidopsis thaliana, the highest identity was found to occur with several “clade IV”

CHX proteins (Sze et al. 2004), where AtCHX18 (AT5G41610.2) and AtCHX19 (AT3G17630.1) proteins showed 53% identity, while AtCHX17 (AT4G23700.1) and AtCHX20 (AT3G53720.1) showed 52% and 51% of identity, respectively. This similarity was indeed confirmed by the clustering of PpCHX1-4 together with clade IV CHX proteins of A. thaliana by phylogenetic analysis of their sequences together with the 28 full-length CHX proteins sequences from A. thaliana (Fig. 7).

Fig. 7. Phylogenetic tree of CHX transporters of Physcomitrella patens and Arabidopsis thaliana.

It is also worth noting that in contrast to the relatively constant number of the CPA2 family KEA genes among early non-vascular and flowering plants (e.g. 7 PpKEA genes in P. patens and 6 AtKEA in A. thaliana), CHX gene orthologs had seemed to multiply during the evolution of early plants into flowering plants (e.g. 4 PpCHX genes in P. patens and 28 AtCHX in A. thaliana) as shown in figure 8.

Fig. 8. Phylogenetic tree of CPA2 family showing both CHX and KEA transporters in A) Physcomitrella patens and, B) Arabidopsis thaliana.

4.2. PpCHX1 and PpCHX2 gene expression and cDNAs cloning To investigate whether the four genes were functional in normal conditions, we performed a search in the EST database of the NCBI website (http://www.ncbi.nlm.nih.go/, accessed in June, 2012) using the sequences of the four CHX genes of P. patens. The search retrieved 27 ESTs for PpCHX1 and 54 for

PpCHX2, but none for either PpCHX3 or PpCHX4. As mentioned earlier, the sequences of PpCHX3 and PpCHX4 genes are inexistent in NCBI databases until this very moment. This perplexing fact motivated us to investigate if PpCHX3 and PpCHX4 genes really do exist in the genome of P. patens plant material manipulated in our laboratory, and thus, we first carried out a PCR using genomic DNA as a template extracted from plants grown in normal conditions. DNA fragments were amplified and detected for both genes by PCR, confirming their existence in the genome (Fig. 9). Next, we investigated the existence of transcripts of these two genes by RT-PCR, finding that in normal conditions of pH (from 5 to 7), or K+ and Na+ concentrations (from 0.1 to 10 mM) the expression of the PpCHX3 or PpCHX4 genes was not detectable. In view of these results we concentrated our study in the PpCHX1 and PpCHX2 genes.

Fig. 9. PCR confirmation for existence of PpCHX3 and PpCHX3 genes in the P. patens genome. Figure shows a UV irradiated agarose electrophoresis gel showing wells A and B with bands of PCR amplified sequences of PpCHX3 and PpCHX4 genes, respectively, run in parallel to gene ladder marker (100 bp). Wells C and D show primer controls (Table 11).

To accurately characterize PpCHX1 and PpCHX2 expression, we monitored both genes’ expression using a quantitative real time PCR (qRT-PCR) method. The qRT-PCR study of the transcript expressions of PpCHX1 and PpCHX2 revealed that both genes were expressed almost to the level of the ACT5 gene of P. patens. In standard growth conditions, PpCHX1 showed a four-fold higher expression than PpCHX2. The expression of PpCHX1 was constant in most growth conditions except for a three-times reduction at pH 4.5. In contrast, the expression of PpCHX2 was lower in standard conditions and slightly increased when the plants were stressed (Table 18). These results suggest that both PpCHX1 and PpCHX2 are housekeeping genes whose expressions are required in all growing conditions. Although the transcript levels showed variability, the detected variations seem too low to have any biological relevance.

Table 18. Real-time PCR determination of transcript expression of CHX1 and CHX2 genes of P. patens at different pH values in the presence and absence of Na+ or K+. Conditions PpCHX1 PpCHX2 pH 5.8 0.84 0.19 pH 5.8 - K+ 1.00 0.59 pH 5.8 + 100 mM Na+ 0.94 0.73 pH 4.0 0.34 0.62 pH 9.0 0.98 0.42

Reported values are the ratio between the transporter transcript abundance and actin transcript abundance. Data represent results of two independent experiments with two replicates each.

Next, we went on with cloning PpCHX1 and PpCHX2 cDNAs from plants growing in standard conditions. As described in materials and methods chapter, we sequenced the whole cloned cDNA (including both the 5’ and 3’ ends) to verify the correct cDNAs sequence of both genes. The first observation we noticed is that the open reading frame (ORF) of PpCHX1 cDNA we cloned did not correspond with the published PpCHX1 CDS sequence (estExt_fgenesh1_pm.C_1230004) in the early JGI database (http://genome.jgi-psf.org/Phypa1_1/Phypa1_1.home.html, accessed in November, 2009); we detected in our cloned cDNA that the real start codon was 27 bp upstream the annotated one and that the real Stop codon was present in an exon 97

incorrectly excluded by the database algorithm. These two annotation errors were corrected later on in the new phytozome database sequence of the gene transcript (Pp1s123_30V6.1) (http://www.phytozome.net, accessed on 27/09/2012). When cloning the PpCHX2 cDNA, we also noticed that the ORF of PpCHX2 cDNA sequence cloned did not match the published CDS sequence either (estExt_gwp_gw1.C_1620028, JGI database); the annotated database CDS sequence lacked 87 bp, from 1989 to 2076, that were excluded by the database algorithm.

4.3. Alternative PpCHX2 proteins The amino acid alignment of the abovementioned conceptual translation of the PpCHX2 gene (estExt_gwp_gw1.C_1620028 in JGI database also known as PHYPADRAFT_191426 in NCBI database) revealed that the translated polypeptide lacked 29 residues in the C tail with reference to other conceptual translations. The missing residues, from 656 to 694, seemed to correspond to a putative intron that might be incorrectly eliminated by some conceptual translations. Consistent with this notion, we found that the corresponding mRNA fragment was present in all ESTs in the database. Furthermore, by RT-PCR, we could not detect the presence of PpCHX2 mRNAs in which this putative intron had been processed, neither in normal conditions of pH (from 5 to 7) and K+/Na+ concentrations (from 0.1 to 10 mM) nor under high Na+ or K+ limited conditions. However, the alternative splicing of this intron could not be absolutely excluded because some CHX transporters showed a gap coinciding with this 29-residue fragment that interrupts a region where CHX transporters keep high sequence homology (Fig. 10). Therefore an alternative splicing in PpCHX2 at the point described above was kept in mind throughout this study.

Fig. 10. Alignment of alternative conceptual translations of the PpCHX2 gene with several mRNA translations. Sequence alignment was performed using Clustal Omega (http://www.ebi.ac.uk/Tools/msa/clustalo/). Sequences used in the alignment are PpCHX2.1 conceptual translated ORF PHYPADRAFT_191426, PpCHX2 translated mRNA, translated ESTs FG570725.1 of Brassica napus, EX095781.1 of Brassica rapa, Y911015.1 of Helianthus annus, FN025549.1 of Petunia x hibrida and FS069311.1 of Solanum melongena.

We, thus, cloned the PpCHX2 cDNA that corresponds to the actual CDS sequence in Phytozome database (Pp1s162_22V6.1) and, for the reasons discussed above, we also constructed in vitro via a 2 step PCR as described in materials and methods, the shorter PpCHX2.1 clone, deleting the 87 bases of the putative intron in the PpCHX2 gene (Fig. 3 in Materials and Methods chapter).

4.4. Subcellular localization of PpCHX1-GFP, PpCHX2-GFP and PpCHX2.1-GFP protein fusions in yeast cells and Physcomitrella patens protoplasts To localize subcellularly the proteins of our study, we constructed expression vectors with the three CHXs tagged at the carboxyl end to green fluorescent protein (GFP) under the expression control of P35S promoter in the case constructs to be expressed in yeast. For P. patens constructs, GFP was under the expression control of the PpACT5 gene promoter, which shows an expression level similar to those of PpCHX1 and PpCHX2 genes (Table 18), as described in materials and methods chapter. First, we expressed the three constructs in the yeast Saccharomyces cerevisiae mutant strain W∆6 (Haro and Rodríguez-Navarro 2003), defective in its main influx systems of K+ TRK1 and TRK2. All three constructs failed to show a clear GFP signal of localization confined to a specific membrane of the cell. The GFP signal appears,

instead, to be present in all the cell; plasma membrane, cytoplasm and other internal membranes (Fig. 11).

Fig. 11. Subcellular localization of A) PpCHX1-GFP, B) PpCHX2-GFP, and C) PpCHX2.1-GFP fusion proteins in Saccharomyces cerevisiae yeast mutant strain W∆6.

Afterwards, we expressed the corresponding constructs in P. patens protoplasts. In these experiments, PpCHX1-GFP showed a punctuate pattern that was compatible with the Golgi network (Fig. 12A). Although the details of the images varied depending on the observed protoplast, the pattern showed in figure 12A was observed in all protoplasts showing the GFP signal. W e confirmed the Golgi network localization afterwards as the GFP signal colocalized with the GmMan1-YFP fluorescent marker of the Golgi complex (Fig. 12B-C). The GFP signal of PpCHX2-GFP showed a double localization, to the cell periphery, enclosing the chloroplasts, and to internal round structures (Fig. 12D and 12G). The peripheral signal was characteristic of the plasma membrane while the internal GFP signal appeared to correspond to the vacuolar membrane. To confirm the latter localization we co-expressed either PpCHX2-GFP or PpCHX2.1-GFP and the γ- TIP-YFP marker of tonoplast. These co-expressions showed that the GFP internal round structures and YFP signals co-localized, demonstrating the tonoplast localization of PpCHX2 (Fig. 12D-F) and PpCHX2.1 (Fig. 12G-I). Unfortunately, we could not co- localize PpCHX2 with a positive control of plasma membrane. Therefore, the temporal or circumstantial localization of PpCHX2 to the plasma membrane is currently only a possibility, which does not have physiological support (see below).

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Fig. 12. Localization of PpCHX1–GFP and PpCHX2–GFP fusion proteins in protoplasts of P. patens. Images of the green fluorescence of PpCHX1–GFP (A), PpCHX2–GFP (D) and PpCHX2.1– GFP (G). Images of the yellow fluorescence of the GmMan1–YFP marker of the Golgi complex (B) and the γ-TIP–YFP marker of the tonoplast (E, H). Merged images of the GFP, YFP marker fluorescence and chloroplast fluorescence are in red (C, F, I). Images show the maximum projection of 11 consecutive sections (A–C), 25 consecutive sections (D–F) and nine consecutive sections (G– I). Frequency of the green fluorescence patterns: (A) pattern appeared in all protoplasts that showed fluorescence with small differences; (D) pattern, most fluorescent protoplast showed peripheral and internal labelings, some protoplasts showed only internal labeling and very few only peripheral labeling; (G) pattern appeared in all protoplasts that showed fluorescence, with small differences.

4.5. Growth rescue of Escherichia coli mutants Before carrying out the functional characterization of the CHX transporters in planta, we needed to have an idea of their mechanism of function by expressing them in simple heterologous systems such as Escherichia coli bacterium and Saccharomyces cerevisiae yeast. E. coli is by far the most widely used expression host for the testing and production of recombinant proteins. Its short generation time, low cost and ease of

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use, as well as its extensive characterization make it an ideal candidate (Sahdev et al. 2008). Another extremely helpful characteristic of E. coli, and bacteria in general, is that it has no subcellular membranes, and thus, any protein (transporter) expressed in it will be inserted in its plasma membrane regardless of it being natively expressed in endomembranes, which consequently facilitates the characterization of such protein (Uozumi 2001). Knowing beforehand that CHX proteins characterized to date in Arabidopsis thaliana are implicated in K+ transport, we chose the TKW4205 mutant of E. coli defective in its K+ transport systems Kdp, TrkA, and Kup (Schleyer and Bakker 1993) which requires a high K+ concentration to grow at pH 5.5. This defect can be highly reduced by heterologous K+ transporters in an expression level-dependent manner (Senn et al. 2001). To test whether the PpCHX proteins transported K+, we cloned the PpCHX1, PpCHX2, and PpCHX2.1 cDNAs into pBAD24 plasmid (Guzman et al. 1995) under the control of the arabinose-responsive PBAD promoter. PpCHX1 partially rescued the growth of TKW4205 at 10 μM arabinose (not shown in Figure 13) but when increasing the arabinose concentration to 100 μM the growth of TKW4205 was excellent at 5 mM ( Fig. 13). As shown with other K+ transporters (Senn et al. 2001), K+ became toxic to E. coli when the expression level of PpCHX1 was increased by increasing the arabinose concentration up to 13 mM (Figure 13 shows the growth inhibition at 13 mM arabinose, 5 mM and higher concentrations of K+). However, when K+ was decreased to 2 mM, the PpCHX1 clone grew fairly well at 13 mM arabinose but not at 100 μM arabinose. PpCHX2 and PpCHX2.1 partially rescued the growth of TKW4205 at 100 μM arabinose (Fig. 13). When increasing the arabinose concentration to 13 mM, the growth of PpCHX2 and PpCHX2.1 was excellent at 5 mM K+, slightly positive at 2 mM K+ (Fig. 13), and was inhibited at 50 mM K+.

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Fig. 13. Growth complementation tests of E. Coli mutant strain TKW4205 at low K+ pH 5.5 by empty vector pBAD24 or PpCHX1, PpCHX2 and PpCHX2.1 constructs. A colony of fresh transformed cells was picked and grown for approximately 3 hours in LB media supplemented 50 mM KCl. Cultures were brought to a uniform cell density and 3-fold serial dilutions were placed on pH 5.5 MM media at 100 μM or 13 mM arabinose and varying concentrations of KCl of 2, 5, 10, 20 or 50 mM. Photos were taken after 4 d of growth at 37 ºC.

When carrying the same experiments on solid medium at pH 7.5, the empty vector pBAD24 clone started growing at very low K+ concentrations (lower than 5 mM). This prevented us to notice the suppression of the defective growth of the mutant in PpCHX1, PpCHX2 or PpCHX2.1 clones compared to empty vector. However, we noticed that only PpCHX1 caused toxicity at 100 µM arabinose concentration, while at 13 mM arabinose concentration, all CHX cDNAs caused toxicity (Fig. 14).

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Fig. 14. Toxicity of E. Coli mutant strain TKW4205 with empty vector pBAD24 or PpCHX1, PpCHX2 and PpCHX2.1 constructs. A colony of fresh transformed cells was picked and grown for approximately 3 hours in LB media supplemented 50 mM KCl. Cultures were brought to a uniform cell density and 3-fold serial dilutions were placed on pH 7.5 MM media at 100 μM or 13 mM arabinose and varying concentrations of KCl of 2, 5, 10, 20 or 50 mM. Photos were taken after 4 d of growth at 37 ºC.

In some kinetic studies it is necessary to measure zero-trans influxes, which cannot be performed with K+. In these studies, Rb+ might substitute for K+ if it is transported. Therefore, for future experiments, it was convenient to know whether PpCHX1 and PpCHX2 transported Rb+. This possibility can be tested in growth experiments because Rb+ can substitute for cellular K+ in a large proportion without any toxic effects. Therefore, we repeated the growth experiments with TKW4205 described above, but using Rb+ in this case instead of K+. The results showed that both PpCHX1 and PpCHX2 rescued the growth of TKW4205 at Rb+ concentrations that were only slightly higher than those found for K+; 10 mM Rb+ instead of 5 mM K+, at 100 μM arabinose for PpCHX1 and 13 mM arabinose for PpCHX2 (Fig. 15). These results demonstrated that both PpCHX1 and PpCHX2 are K+ transporters that show a notable capacity to transport Rb+.

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Fig. 15. Growth complementation tests of E. Coli mutant strain TKW4205 at low Rb+ pH5.5 with empty vector pBAD24 or PpCHX1, PpCHX2 and PpCHX2.1 constructs. A colony of fresh transformed cells was picked and grown for approximately 3 hours in LB media supplemented with 50 mM KCl. Cultures were brought to a uniform cell density and 2-fold serial dilutions were placed on pH 5.5 media at 100 μM or 13 mM arabinose and varying concentrations of RbCl of 10, 20 or 50 mM. Photos were taken after 4 d of growth at 37 ºC.

4.6. PpCHX1 and PpCHX2 complement the kha1 mutation in yeast Yeast, in particular Saccharomyces cerevisiae, share many characteristics of E.coli bacterium that makes it a great model organism for biological and genetic studies including the heterologous expression of many proteins; completely sequenced genome , short life cycle, and easy genetic transformation and manipulation in the laboratory. S. cerevisiae, however, has an additional advantage being an eukaryote and, thus, phylogenetically less distant to plants than the prokaryote E. coli. Several yeast mutants have been extensively used to functionally characterize plant transporters (Dreyer et al. 1999). For example, the heterologous expression of plant transporters in the S. cerevisiae yeast mutant W∆6 (Mat a ade2 ura3 trp1 trk1∆::LEU2 trk2∆HIS3), defective in the TRK1 and TRK2 K+ uptake systems (Haro 105

and Rodríguez-Navarro 2003), is routinely performed to test K+ and Na+ influxes by the expressed transporters. However, these mutants can usually be complemented only with plasma membrane transporters and not by transporters of internal membranes. As previously described in this chapter, the subcellular localization of our three cloned CHX transporters using GFP in yeast cells did not show a clear localization to a specific cellular membrane, being instead present in various locations including the plasma membrane (Fig. 11). This result, together with PpCHX2p being located to the plasma membrane in P. patens protoplasts (Fig. 12), left the window open to a possibility, although minimal, that any of these transporters might promote K+ or Na+ influx in the W∆6 mutant. First we tested whether PpCHX1, PpCHX2 or PpCHX2.1 suppressed the defect of this mutant by promoting high affinity K+ uptake, and we found that none of these transporters succeeded in suppressing the defective growth of the mutant at low K+ concentrations when carrying out yeast drop-test experiments (Fig. 16). Also, their implication in high affinity Na+ transport in the W∆6 mutant turned to be null, as K+- starved yeast clones of PpCHX1, PpCHX2 and PpCHX2.1 failed to deplete micromolar concentrations of Na+ from liquid AP media during the course of time.

Fig. 16. Tests of functional expression of PpCHX1, PpCHX2 and PpCHX2.1 in yeast mutant W∆6. Cells transformed with empty pYPGE15 vector or PpCHX1, PpCHX2 and PpCHX2.1 cDNA constructs were grown overnight in YPD growth medium supplemented with 50 mM KCl. In the following day, cultures were brought to a uniform cell density of 0.3 and 3-fold serial dilutions were placed on (SD + Adenine and Tryptophan) media at pH 6.5 and varying concentrations of KCl of 100 μM, 1 mM, or 5 mM. Photos were taken after 4 d of growth at 28 ºC.

As previously reported, PpCHX1, PpCHX2 and PpCHX2.1 showed the ability to transport Rb+. Thus, using Rb+ as an analogue of K+, we wanted to test these

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transporters ability to mediate low affinity K+ transport in the millimolar range in the W∆6 mutant. Again, none of the three transporters promoted a faster millimolar Rb+ influx in the clones compared to the empty vector pYPGE15, being the low affinity Rb+ influx rate almost identical in all the clones (Fig. 17). This was also the case for Na+ low affinity influx experiments.

Fig. 17. Time course of Rb+ influx measuring its internal content (nmol/mg) in S. cerevisiae strain W∆6 mutant transformed with empty vector pYPGE15 (triangles), PpCHX1 (rombuses), PpCHX2 (squares), and PpCHX2.1 (circles). Cells were prepared from cultures grown overnight in liquid YPD media supplemented with 50 mM KCl. Next day, cells were starved for K+ for 4 hours, and influx tests were started by the addition of 10 mM RbCl, taking samples during the course of one hour.

Another possibility needed to be tested was if these transporters are involved in Na+ and/or K+ effluxes. In this sense, we used the S. cerevisiae yeast mutant B3.1 (ena1∆::HIS3::ena4 nha1∆::LEU2) defective in its Na+ and K+ efflux systems ENA1-4 ATPases and NHA1 efflux antiporter (Bañuelos et al. 1998). Expressing our cDNAs in this mutant resulted again in no complementation using the drop-test experiments; the transporters failed to improve the defective growth of B3.1 under either high Na+ (Fig. 18 A) or high K+ conditions (Fig. 18B), concluding that none of these transporters are involved in Na+ or K+ efflux in yeast.

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Fig. 18. Tests of functional expression of PpCHX1, PpCHX2 and PpCHX2.1 in yeast mutant B3.1. Cells transformed with empty pYPGE15 vector or PpCHX1, PpCHX2 and PpCHX2.1 constructs were grown overnight in growth medium YPD. In the following day, cultures were brought to a uniform cell density of 0.3 and 3-fold serial dilutions were placed on media at pH 6.5 and varying concentrations of A) KCl: 100 mM, 400 mM and 800 mM; or B) 1 mM KCl with either 10 mM or 30 mM NaCl . Photos were taken after 4 d of growth at 28 ºC.

Negative results in this type of experiments may be explained because either the protein is not targeted to the yeast plasma membrane or the transporter is not active in yeast cells. The latter occurs frequently with HAK transporters (Rubio et al. 2000, Garciadeblás et al. 2007). Yeast mutants in the KHA1 gene, on the other hand, have also been used for the functional expression of several endomembrane transporters. KHA1 is a K+ antiporter that localizes to Golgi like structures in S. cerevisiae (Maresova and Sychrova 2005, Ariño et al. 2010). Yeast kha1 single mutants do not show a clearly defective phenotype. However, additional mutations in ENA1-4 and NHA1 Na+ and K+ efflux

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systems, mutant strain LMB01, produce a defective growth at acidic (4.5) or alkaline (7.5) pHs with low K+ concentrations in the medium (Maresova and Sychrova 2005). This defect becomes even more pronounced when an additional mutation in TRK1, TRK2 and TOK1 K+ influx systems is added to the LMB01 genotype (mutant strain LMM04) (Maresova and Sychrova 2005). These defective growths at acidic or alkaline pHs with low K+ was reported to be improved by several plant CHX transporters (Maresova and Sychrova 2006, Zhao et al. 2008, Chanroj et al. 2011). To test whether PpCHX1, PpCHX2 or PpCHX2.1 suppress the defect of the kha1 mutation, we transformed the kha1 ena1-4 nha1 mutant (LMB01) with the PpCHX1, PpCHX2, and PpCHX2.1 cDNAs. At pH 7.5 the three transporters improved the growth at low K+, 50 and 100 μM, and the effect was still observable at 1 mM K+ (Fig. 19A). It is worth highlighting that the defect suppressed by the PpCHX clones was exclusively due to the kha1 mutation. The ena1-4 nha1 KHA1 (B3.1) strain grew normally at pH 7.5 and 50 μM K+, and the PpCHX clones had no effect on the slow growth of this strain in the medium without added K+ (Fig. 19B).

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Fig. 19. Tests of functional expression of PpCHX1, PpCHX2 and PpCHX2.1 in yeast mutants LMB01 and B3.1. (A) LMB01 or (B) B3.1 cells transformed with empty pYPGE15 vector or PpCHX1, PpCHX2 and PpCHX2.1 constructs were grown overnight in growth medium (SD + Adenine and Tryptophan) supplemented with 50 mM KCl. In the following day, cultures were brought to a uniform cell density of 0.3 and 3-fold serial dilutions were placed on media at pH 7.5 and varying concentrations of KCl of 50 μM, 100 μM or 1 mM. Photos were taken after 4 d of growth at 28 ºC.

The described successful complementation of strain LMB01 at pH 7.5 occurred only at this pH. As expected, the only defect of the LMB01 mutant at pH 6.5 was a barely detectable impairment of growth at 50 μM K+, which was not significantly modified by the PpCHX clones (Fig. 20A and B).

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Fig. 20. Functional expression of PpCHX1, PpCHX2 and PpCHX2.1 in yeast mutants LMB01 and B3.1. (A) LMB01 or (B) B3.1 cells transformed with empty pYPGE15 vector or PpCHX1, PpCHX2 and PpCHX2.1 constructs were grown overnight in growth medium (SD + Adenine and Tryptophan) supplemented with 50 mM KCl. In the following day, cultures were brought to a uniform cell density of 0.3 and 3-fold serial dilutions were placed on media at pH 6.5 and varying concentrations of KCl of 50 μM, 100 μM or 1 mM. Photos were taken after 4 d of growth at 28 ºC.

The abovementioned results point to the fact that the mechanism of action of CHX transporters of P. patens is more of a pH and/or K+ homeostasis regulation in internal compartments, probably including Golgi apparatus, rather than its direct implication in K+ and Na+ influx or efflux through the plasma membrane of the yeast cell. Another aspect that we observed in our drop-test experiments using the LMB01 strain, is the slight impairment of growth in PpCHX1 clone at pH 7.5 in the presence of low concentrations of Na+ (10 mM) compared to the empty vector pYPGE15, PpCHX2 and PpCHX2.1 clones (Fig. 21).

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Fig. 21. Growth of yeast mutants LMB01 and B3.1 in the presence of 1 mM KCl and 10 mM NaCl at A) pH6.5 and B) pH 7.5. Cells transformed with empty pYPGE15 vector or PpCHX1, Pp C HX 2 and PpCHX2.1 constructs were grown overnight in growth medium YPD. In the following day, cultures were brought to a uniform cell density of 0.3 and 3-fold serial dilutions were placed on media at pH 6.5 and 7.5 and 1 mM KCl with 10 mM NaCl. Photos were taken after 4 d of growth at 28 ºC

This growth impairment of PpCHX1 in the presence of Na+, although very minimal, indicates that PpCHX1 transporter does not seem discriminate between K+ and Na+, in contrast to PpCHX2 and PpCHX2.1. Consequently PpCHX1 seems to deliver Na+ to internal compartments resulting in toxicity and an impaired growth of the clone. This slight growth impairment of PpCHX1 was later confirmed after carrying out several growth curve experiments comparing its growth to the growth of pYPGE15 and PpCHX2 clones. In these experiments, the growth impairment was also observed under both pH 6.5 and 7.5 (Fig. 22).

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Fig. 22. Gro wt h curves of the yeast mutant LMB01 transformed with empty vector pYPGE15 (solid squares), PpCHX1 (open circles) and PpCHX2 (open triangles) cDNAs. Cells were grown in SD medium (+ adenine and tryptophan) at pH 6.5 or 7.5 supplemented with either 1 mM KCl or 1 mM KCl and 10 mM NaCl at A) pH 6.5 and B) pH 7.5. The results were obtained from three independent measurements using the Bioscreen C device.

4.7. Functional analyses of ∆Ppchx1 and ∆Ppchx2 plants To elucidate the in planta roles of PpCHX1 and PpCHX2, we disrupted the PpCHX1 and PpCHX2 genes individually, using the disruption fragments shown in figure 23 (described in section 3.5.1 of Materials and Methods chapter). We isolated four ∆Ppchx1 and seven ∆Ppchx2 lines, in which the hygromycin and zeocin resistance cassette substituted for the coding region of PpCHX1 or PpCHX2, respectively. These mutant lines were named ∆Ppchx1-(1-4) and ∆Ppchx2-(1-7). Also, a ∆Pphak1 ∆Ppchx2 double mutant was constructed by disrupting the PpCHX1 gene of the ∆Pphak1 mutant previously obtained in our laboratory (Garciadeblás et al. 2007), using the same strategy shown in figure 23B. In all these knockout mutant lines, the absence of the corresponding transcripts was verified by RT-PCR.

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Fig. 23. Targeted disruption of the PpCHX1 and PpCHX2 genes by double homologous recombination. The disruption fragment that was transformed into P. patens is shown in parallel with (A) PpCHX1 gene and (B) PpCHX2 gene (see section 3.5.1 of Materials and Methods chapter).

During the process of obtaining and multiplying of all these lines, the growth and morphological characteristics of the mutant plants were absolutely normal when grown in standard conditions. With the aim of identifying the growth defects of the ∆Ppchx1 and ∆Ppchx2 plants, we applied a battery of growth tests including: high K+ or Na+ concentrations, K+ starvation, high and low pH values, high Ca2+ concentrations, and the combination of some of them. In all cases, we found no differences between wild-type and mutant plants in terms of morphology and growth (Fig. 24). Also, it is worth noting that regarding the expression of PpCHX1 and PpCHX2, we found that in ∆Ppchx1 or ∆Ppchx2 plants, the disruption of one of the two genes did not significantly affect the expression level of the other. In the case of PpCHX3 and PpCHX4 genes, their transcripts were approximately 500 and 2000 times less abundant, respectively, than PpCHX1 transcripts in wild type plants, and the disruption of either PpCHX1 or PpCHX2 did not change these low expression levels of PpCHX3 and PpCHX4.

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Fig. 24. Normal morphology and growth of the single mutants ∆Ppchx1 and ∆Ppchx2, as well as the double mutant ∆Pphak1 ∆Ppchx2 compared to wild-type Physcomitrella patens plants either under A) normal conditions on solid medium, or B) K+ starvation for 2 weeks in liquid medium.

The obvious next experiment was to study K+ and Rb+ fluxes, because a mild defect in these fluxes might not affect the growth of the mutant plants. However, this approach applied only to ∆Ppchx2 plants because the location of PpCHX1 to the Golgi apparatus made it unlikely that ∆Ppchx1 plants were defective in K+ and Rb+ fluxes. In its plasma membrane location, PpCHX2 might mediate either K+ influx or K+ efflux. In the former case, the absence of PpCHX2 should result in a defective K+ uptake. To test this possibility, we followed how ∆Ppchx2 plants depleted micromolar concentrations of K+ and Na+, 160 and 120 µM respectively, in the medium compared

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to wild-type plants. This experiment revealed an almost identical pattern of high affinity K+ uptake followed by Na+ uptake in both wild-type and ∆Ppchx2 plants (Fig. 25).

Fig. 25. Time course of the decrease of external K+ and Na+ concentrations of P. patens plants. Symbols: wild-type plants, closed squares (K+ uptake) or open squares (Na+ uptake); ∆Ppchx2 plants, closed triangles (K+ uptake) or open triangles (Na+ uptake).

Nevertheless, as reported in previous studies, PpHAK1 transporter mediates “active” high affinity K+ influx in P. patens (Garciadeblás et al. 2007). Thus, the presence of this transporter might conceal any defect produced by the disruption of PpCHX2. Therefore, we constructed the ∆Pphak1 ∆Ppchx2 double mutant, which was tested in parallel with the ∆Ppchx2 single mutant. First, we tested K+ or Rb+ uptake in three different preparations of plants: normal, K+-starved, and Rb+ loaded, performing the tests at neutral, low, and high pH values. In these tests, ∆Ppchx2 plants did not show any appreciable defect. Then we used conditions of high Ca2+, in which ∆Pphak1 plants showed more clearly its defective Rb+ uptake, but again the addition of the ∆Ppchx2 mutation did not reveal any detectable effect due to the lack of the PpCHX2 transporter (Fig. 26).

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Fig. 26. Time course of the decrease of external Rb+ concentration of K+-starved P. patens plants. Symbols: wild-type plants, open circles; ∆Ppchx2 plants, closed circles; ∆Pphak1 plants, closed squares; and ∆Pphak1 ∆Ppchx2 plants, closed triangles.

Another possibility tested was whether PpCHX2 promotes low affinity Rb+ uptake. Thus, we followed the kinetic of influx at a wide range of Rb+ concentrations, 3, 5 and 10 mM, measuring the internal Rb+ concentrations in both wild-type and ∆Ppchx2 plants over the course of several hours (Fig. 27). No significant differences were observed between both plants in low affinity Rb+ uptake.

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Fig. 27. Time course of Rb+ influx measuring internal content ( n m o l / m g ) in P. patens plants. Symbols: wild-type plants, open circles; ∆Ppchx2 plants, closed circles. Plants grown in normal conditions for a week and influx tests were started by the addition of 10 mM RbCl, taking samples during the course of 5 hours.

The tests described above were aimed to detect defective K+ or Rb+ influxes, but the PpCHX2 transporter might mediate K+ efflux. To test this possibility, we grew wild- type and ∆Ppchx2 plants in normal BCDNH4 medium for one week and then the plants were thoroughly rinsed and transferred to KFM media for another week. When we measured the decrease of K+ contents in wild-type and ∆Ppchx2 plants through the course of several days in KFM media, we found no significant differences between both of them. Next, to investigate other possible functions of PpCHX2, we tested for alterations in K+/Rb+ exchanges. For this purpose, we substituted 10 mM Rb+ for the K+ content of the culture medium. First we grew the plants in normal BCDNH4 medium for one week, then the plants were rinsed and transferred to KFM medium with 10 mM Rb+ for a second week. During the course of the second week, we followed the external K+ concentration and the internal Rb+/K+. When plants were grown at 10 mM Rb+, the re- uptake of the K+ lost by the plants is inhibited, and consequently a defective K+ efflux 118

should result in a slower increase of external K+ of the medium in the case if ∆Ppchx2 was defective in efflux. Once again, however, we did not find differences that reveal a defective K+ efflux. In contrast, when we recorded the Rb+/K+ ratio we found that the ratio was slightly higher in ∆Ppchx2 plants. In general, the variability was lower in experiments carried out with the same batch of plants –e.g. a time course experiment (Fig. 28)– than in experiments with independent batches of plants (Fig. 29). Although the differences were sma l l , the statistical analyses of independent five-days exchange experiments (Fig. 29) revealed that the Rb+/K+ ratio in ∆Ppchx2 plants was significantly higher than in wild type plants (2.24 ± 0.46 versus 1.75 ± 0.21; n = 7; t-test, p = 0.024).

Fig. 28. Rb+/K+ exchange in wild-type and ∆Ppchx2 plants. Plants grown at 10 mM K+ for one week then transferred to K+-free medium with 10 mM Rb+. Internal contents of Rb+ and K+ were measured and the Rb+/K+ ratio calculated. Data points represent a time course experiment of the Rb+/K+ content; symbols: wild-type plants, open circles; ∆Ppchx2 plants, closed circles.

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Fig. 29. Means of Rb+/K+ ratios in seven independent experiments measured at the fifth day for wild-type plants (closed bars) and ∆Ppchx2 plants (open bars). Standard errors are shown and means are significantly different (P = 0.024) according to t-test.

In the described experiments, the K+ loss to the external medium was determined with high precision and no differences were found between ∆Ppchx2 and wild type plants and; on the other hand, we found that Rb+ uptake was not affected in ∆Ppchx2 plants. Thus, the defect underlying the higher Rb+/K+ ratio in ∆Ppchx2 plants seems to be a higher retention of Rb+ in the steady state–influx versus efflux–that determines the Rb+ content. This may result from a slower transfer of Rb+ from either the vacuole to the cytosol, resulting in a higher vacuolar Rb+ content, or from the cytosol to the external medium, resulting in a higher cytosolic Rb+ content. Unfortunately, it is not currently possible to distinguish between these two possibilities by measuring Rb+ contents.

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5. Discussion

5. DISCUSSION 5.1. Analysis in silico and cloning of CHX transporters of Physcomitrella patens This study investigates the function of CHX transporters in P. patens and is the first of its kind to investigate members of the CPA2 family in early non-vascular plants. This family, comprised of CHX and KEA transporters, is the less studied family of cation-proton antiporters in plants (Sze et al. 2004, Aranda-Sicilia et al. 2012). In the case of CHX transporters, scarce information is known, as most functional studies on CHX transporters are relatively recent. These are so far limited to Arabidopsis thaliana (Chanroj et al. 2012) as well as some superficial studies on Oryza sativa rice, (Sze et al. 2004, Senadheera et al. 2009). It is suggested from comparative genomics studies that CHX transporters have evolved from fresh water green algae charophytes rather than from marine green algae chlorophytes (Chanroj et al. 2012). This proposal is supported by the fact that CHX homologs are absent in some chlorophytes, like Chlamydomonas reinhardtii or Volvox carteli, but present in the charophyte Spirogyra pratensis (one CHX homolog). Whether the ancestral CHX gene was horizontally transferred or not to charopytes from the bacterial GerN or cyanobacterial NhaS4 highly similar homolog genes, probably through endosymbiosis, is a mystery and cannot be asserted for sure. However, comparative genomics studies between fresh water green algae (charophytes), early non-vascular land plants (bryophytes) and flowering plants suggest that CHX genes have suffered several events of duplication along the course of evolution from fresh water green algae into higher plants. The first event of duplication of the ancestral plant CHX gene probably occurred in early non-vascular plants (e.g. 3 CHXs in Selaginella moellendorffii and 4 in Physcomitrella patens). Interestingly, subsequent duplications of CHX genes in flowering plants raise many questions on their specialized functional importance to flowering plants. As mentioned in the past chapter, in contrast to the relatively constant number of the CPA2 family KEA genes among early non-vascular and flowering plants, CHX gene orthologs have continued to multiply during the evolution of flowering plants. For example, dicotyledonous plants have roughly double the number of CHX genes in monocotyledonous plants (e.g. 15 CHX genes in Zea mays and 28 in A. thaliana). Nitrogen fixating plants (legumes) have also around double the

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number of CHX genes than non-nitrogen fixating plants (e.g. Glycine max have 46 predicted CHX genes). The abovementioned apparent duplication events of the CHX genes, in contrast to their closely related CPA2 gene family members KEA, might suggest that CHX genes have played crucial roles for adaptation during the evolution from marine algae to freshwater algae, and then to land plants, as well as many specific cell-type functions in flowering plants. This specific cell-type functions are evident, for example, in A. thaliana, where its 28 CHX transporters are expressed either specifically or preferentially in pollen grains (Sze et al. 2004). In fact when aligning CHX members of A. thaliana together with the 18 members of rice and drawing a phylogenetic tree, it was found that all rice CHX members grouped only to clades I, IV and V, with no rice CHX transporter orthologues grouping to clades II or III (Sze et al. 2004). It is worth noting that, 10 out of 15 AtCHX members of clades II and III are specifically expressed in pollen, while the 5 remaining are preferentially expressed in pollen. This suggests that the AtCHX members of clades II and III play a specific and important role in A. thaliana pollen, one not present in pollen of monocot plants like rice. This possible role, and the significance of this great number of specialized pollen transporters, however, remains to be clarified knowing that only clade IV AtCHX21 and AtCHX23 transporters seem to have a significant effect on the pollen tube guidance process to the ovule (Lu et al. 2011), while none from clades II and III have been discovered with this function until this moment. Besides the important functions of A. thaliana CHX transporters in pollen, further results on other CHX transporters lead to the establishment of an emerging model suggesting that these transporters play important roles in the cellular homeostasis of K+. For example, AtCHX20 transporter was found to be expressed and functional in guard cells (Cellier et al. 2004) while AtCHX17 and AtCHX13 are functional in roots (Padmanaban et al. 2007, Zhao et al. 2008). Regarding the subcellular localization of CHXs, although AtCHX13 (Zhao et al. 2008) and AtCHX21 (Hall et al. 2006) are apparently functional in the plasma membrane, most CHX transporters appears to reside in internal membranes such as prevacuolar, endoplasmic reticulum, Golgi or other nonspecific endomembranes (Padmanaban et al. 2007, Chanroj et al. 2011, Lu et al. 2011).

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Concerning the phylogenetic relationship between P. patens and A. thaliana CHX transporters, PpCHX1-4 transporters group to the clade IV (Fig. 7) of the Arabidopsis thaliana CHX phylogenetic tree (Sze et al. 2004, Chanroj et al. 2012). As expected, none of the four P. patens CHX transporters grouped to clades II and III, the two clades comprising AtCHXs specific to A. thaliana pollen, as was the case of rice mentioned earlier. Nevertheless, clade IV has two AtCHX members that are “specifically” expressed in pollen, AtCHX15 and AtCHX23 (Sze et al. 2004). These, however, are phylogenetically located at a further distance from other AtCHXs in this cluster, as well as PpCHX1-4. It might be speculated that the presence of AtCHX15 and AtCHX23 in clade IV suggests that both are conserved CHXs that might have served as a basis for duplication of other members during the evolution of flowering plants (e.g. me mbers of clades II and III). It is also worth noting that the remaining AtCHX members in clade IV are also expressed in pollen. However, in this case they are not specific to pollen and are expressed in other structures including roots and leaves (AtCHX16, 17, 18, 19, 20, and 21). In general, the relatively close phylogenetic distance between PpCHX1-4 and AtCHX15-23 in clade IV may lead us to suspect that these AtCHXs have been relatively conserved during evolution, given their higher similarity to the more “primitive” CHXs of P. patens. Another fact worth noting is that PpCHX3 and PpCHX4 are phylogenetically more distant to PpCHX1 and PpCHX2, suggesting that they might play different roles in P. patens. Given that we did not detect any expression of PpCHX3 and PpCHX4 genes in gametophytes, it might be suspected that these could be expressed in sporophytes, and thus, play a possible role there. The PpCHX3 and PpCHX4 in subclade IV are phylogentically close to AtCHX20, a K+ transporter expressed in guard cells (Padmanaban et al. 2007), which makes the possibility that PpCHX3 and PpCHX4 play a role in P. patens guard cells a reasonable hypothesis for further investigation. It is well documented, however, that P. patens gametophyte phase cells (protonema and gametophores) lack guard cells which are, nevertheless, present in sporophytes (Knight et al. 2009). In all cases, this hypothesis is rather difficult to test. On one hand, the current RNA and DNA extraction protocols in P. patens are adapted for gametophyte tissue (especially protonema), and when applied to sporophyte tissues (sporangiophores), the DNA yield is very low and the RNA yield is almost non-existent. Thus, assessing gene expression of PpCHX3 and PpCHX4 is a complicated task. If this 125

shortcoming is overcome soon, future works on PpCHX3 and PpCHX4 are mandatory. On the other hand, the sporophyte phase is too minute to work with and no current protocols are developed to test any possible defective K+ uptake or any other abnormal transport in P. patens mutants. Although pollen and guard cells do not exist in P. patens protonema, the basic cellular functions mediated by the CHX transporters, that are primary altered in these mutants (e.g. membrane trafficking events that affect protein and cargo sorting; Chanroj et al. 2012), are probably very similar in Arabidopsis and P. patens. Also, our results on the localization of PpCHX1 to the Golgi complex and PpCHX2 to plasma membrane and tonoplast seem consistent with the Arabidopsis model mentioned earlier. The double localization of a transporter is not uncommon and was previously reported in the HAK transporter TRH1 showing a double localization in the plasma membrane and tonoplast (Rigas et al. 2012). Interestingly, PpCHX2.1, the shorter version of PpCHX2, did not localize to the plasma membrane. When analyzing the PpCHX2 protein sequence with various algorithms, we detected the existence of a low complexity region (LCR) in the residues that we excluded (HSESLGLSKLHHSSL) to construct the PpCHX2.1 version using the SEG algorithm program (ftp://ftp.ncbi.nih.gov/pub/seg/seg/) (Wootton and Federhen 1996). These low complexity sequences are sequences with overrepresentation of a few residues that strongly indicate disorder in the protein. Such disordered regions have been shown to be involved in a variety of functions, including DNA recognition, modulation of specificity/affinity of protein binding, molecular threading, activation by cleavage, and control of protein lifetimes (Dunker et al.1997). The shorter PpCHX2.1 version not localizing to the plasma membrane like PpCHX2, thus, might suggest that the missing residues with reference to PpCHX2 have localization determinants. Currently it cannot be predicted whether this finding reflects a physiological process, e.g. alternative splicing, or if it is a fortuitous experimental response. Alternative splicing in transporters is well documented in animal cells, where it is found that it can change not only the cellular localization of the transporter, but its transport properties as well (Lazaridis et al. 2000). In plants, although less studied, is not uncommon either (Takahashi et al. 2007, Mao et al. 2008, Cotsaftis et al. 2012). In any case, the missing residues in PpCHX2.1 did not affect the functional expression of the protein in either E. coli or yeast mutants. 126

5.2. Functional analysis of PpCHXs in heterlogous systems of expression 5.2.1. Escherichia coli To deepen our knowledge regarding the transport functions of PpCHX1 and PpCHX2, we started by their functional study in simple heterologous systems of expression; K+ uptake deficient mutants of Escherichia coli bacterium and Saccharomyces cerevisiae yeast. K+ uptake deficient E. coli TKW4205 mutant has been successfully used for expressing and studying the function of different plant transporters such as HAK (Senn, 2001), NHAD (Barrero Gil et al. 2007), SOS1 (Garciadeblás et al. 2007), and recently CHX transporters (Chanroj et al. 2011). Overall, our study revealed that the functional expression of PpCHX1 and PpCHX2 in the E. coli TKW4205 mutant (Fig. 13) was similar to previous findings with AtCHX17, AtCHX20 (Chanroj et al. 2011), and AtCHX23 (Chanroj et al. 2011, Lu et al. 2011). All these transporters improved substantially the capacity of TKW4205 to grow at low K+, which demonstrates that all these proteins mediate K+ uptake. According to these growth experiments PpCHX1 and PpCHX2 seem to play identical functions in E. coli, however, they show different specific activities. This is suggested by the different concentrations of arabinose that are required to obtain similar responses with the two cDNAs, which implies different expression levels of the + arabinose-responsive PBAD promoter . For example, for rapid growth on 5 mM K , the PpCHX1 clone required 100 μM arabinose, while PpCHX2 clone required 13 mM arabinose, which represents more than tenfold different transcript expression levels (Guzman et al. 1995). On the other hand, a concentration of 13 mM arabinose was toxic for the PpCHX1 clone at 5 mM and higher K+ concentrations, but suitable for PpCHX2 clone. Also, PpCHX1 clone growth at 2 mM K+ occurred at 13 mM but not at 100 μM arabinose. This suggests that 100 μM arabinose concentration was not sufficient to induce the expression of the adequate number of PpCHX1 transporters required to fulfill the K+ necessities of PpCHX1 clone to grow at 2 mM K+. By the same reasoning, it is highly likely that the slower growth of PpCHX2 and PpCHX2.1 clones at 2 mM K+ compared to PpCHX1 (Fig. 13) can be explained because of the low the number of transporters, i.e. the low Vm a x of the system. Overall, the slower growths, thus, can be

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explained by the slower rates of K+ uptake, which depends on the number of + + transporters, K concentration, and K influx kinetics, i.e. Km and specific activity of the system. These observations are important regarding the mechanism of transport discussed below because it strongly suggests that the growth at low K+ is limited by the kinetics and not by the thermodynamics of the system. To understand the physiological function of CHX transporters, the study of their functional mechanism is of crucial importance. For this issue, the mechanism underlying the CHX mediated K+ uptake in E. coli constitutes the most basic information. The first question to answer is whether or not they mediate “active” K+ uptake. In our experiments (Fig. 13) this uptake could be mediated by either a “passive” K+ uniporter or by an “active” K+-H+ symporter. The possibility of an electroneutral K+/H+ antiporter is improbable as it would mediate K+ efflux (where no growth would be observed). To address this question, the simplest way to distinguish between the uniport and symport mechanisms is to calculate whether the value of the membrane potential can explain the internal/external ratio of K+ concentrations in growing cells of E. coli. To calculate this ratio, the external K+ concentration is 2 mM, because the PpCHX1 clone was found to grow at this concentration. In a medium without Na+, the internal concentration of K+ in growing cells of E. coli is 211 mM (Schultz and Solomon 1961). This concentration might be reduced at K+ limited growth rates, but not very much, considering that the K+ content of chemostat cultures of Enterobacter + + aerogenes are not greatly reduced even in the presence of NH 4 and Na (Tempest and Strange 1966). According to these data, thus, an internal K+ concentration lower than 150 mM is absolutely unlikely. Consequently, this corresponds to a minimal internal/external ratio of 75:

To get this ratio by “passive” K+ uptake, it would require a membrane potential of -112 mV. However, E. coli cannot attain that membrane potential at pH 5.5. The membrane potential in E. coli has been extensively studied (Padan et al. 1976, Zilberstein et al. 1979, Felle et al. 1980, Bakker and Mangerich 1981, Kashket 1982, Setty et al. 1983). According to these studies a likely value of the membrane potential of 128

E. coli at pH 5.5 is between -90 and -95 mV. The most negative value is given in the study of Felle et al. (1980) by microelectrode recording, reporting a measured value of - 80 mV that is corrected to -100 mV after calculating the effect of the current leakage through the membrane seal. Accordingly, given that a membrane potential value of -112 mV cannot be attained at pH 5.5 in E. coli cells, and consequently neither the value of 75 internal/external K+ ratio, it can be concluded by these simple calculations that it is highly unlikely that either PpCHX1 or PpCHX2 mediate a “passive” K+ uptake. Also, to interpret these calculations, it is worth noting that a positive growth is an unequivocal proof of fulfilling an energetic requirement, while a negative growth might be the consequence of an influx that is too slow to support a detectable growth rate. We discussed above that the growth of the PpCHX1 and PpCHX2 clones of TKW4205 at low K+ depended on the arabinose concentration, which points out that growth is limited by the kinetics (i.e. amount of transporters) rather than by the thermodynamics (i.e. energetic of transport) of the system. These abovementioned thermochemical calculations do not prove but strongly suggest that PpCHX1 and PpCHX2 mediate an “active” K+ uptake. Therefore, assuming this conclusion, the most likely mechanism of action is a K+-H+ symport in E. coli (Rodriguez-Navarro et al. 1986). Driven by this mechanism of transport, one K+ ion can move against its electrochemical gradient with the concomitant movement of one proton, which may move up or down its electrochemical gradient. This same conclusion can be applied to AtCHX20 and AtCHX17 based on similar growth tests of E. coli transformants reported previously by Chanroj et al. (2011). These coincidences between Arabidopsis and P. patens CHXs give strong support to the notion that a K+-H+ symport mechanism might apply to many, if not all, CHXs. This mechanism is of crucial important to develop a comprehensive functional model of these transporters that explains the results obtained with the yeast mutants. In contrast, it is worth noting that if the mechanism of the CHX transporters were K+ uniport they would be equivalent to K+ channels (without voltage gating), through which K+ moves but not H+. Using a different approach, the same issue has been previously discussed by Chanroj et al. (2011) for the Arabidopsis transporters AtCHX17 and AtCHX20. This approach was not followed in present study because of its methodological uncertainties. First, the authors tried to extract mechanistic conclusions from differences in transport rates at pH 7.2 and 6.2, which is a difficult 129

task to achieve. Instead, these differences can be explained on kinetic basis in most cases. For example, the toxicity of PpCHX1 and PpCHX2 at pH 7.5 (Fig. 14) cautions about the use of neutral pHs for K+ uptake experiments in the TKW4205 strain. The observed toxicity at pH 7.5 can be explained by a high K+ influx which, together with the intrinsic influx of the mutant cells and the influx mediated by the CHX transporter, cannot be controlled by the bacterial cells and consequently causing toxicity in them. This is similar to the toxic effect discussed earlier produced at pH 5.5 and 13 mM arabinose, at 50 mM K+ for PpCHX2 and 5 mM K+ and higher concentrations for PpCHX1 (Fig. 13). At pH 5.5 the interpretation of the results is simpler because at this pH the intrinsic K+ influx in the bacterial cells is negligible; for that reason the mutant does not grow without the CHX transporters. In contrast, at pH 7.5, the intrinsic K+ influx is much higher, sufficient to support a rapid growth at 5 mM K+ and lower concentrations (not shown, but see the excellent growth at 5 mM K+ in Fig. 14), and exceeding the influx that can be mediated by the expression of the CHX transporters at any K+ concentration. The second uncertainty in Chanroj et al. (2011) approach is that the kinetic analysis of Rb+ influx performed is unlikely to provide reliable mechanistic information. Specifically, these analyses are performed using the initial rates of Rb+ uptake, which means that they are performed when the internal concentration of Rb+ is very low and the chemical Rb+ gradient is huge. The rate tests under these conditions, including those performed with uncouplers, will reveal kinetic responses of the transporters but very little of their thermodynamical dependence. In summary, although K+-H+ symport is currently the most likely functional mechanism of CHX transporters as proposed above, a definitive demonstration is still pending. Most likely this demonstration will require the use of membrane vesicles obtained from the plant membranes where the transporters are expressed. Unfortunately, both the preparation these vesicles and the tests for K+-H+ symport in plant endomembrane vesicles present many technical difficulties, which makes the use of indirect approaches inevitable in the current studies on the functional mechanism of CHX transporters.

5.2.2. Saccharomyces cerevisiae So far, much of the knowledge generated on the function of different plant transporters has been acquired by heterologous expression in Saccharomyces cerevisiae 130

yeast mutants defective in K+ and/or Na+ transport systems (Hasegawa et al. 2000, Serrano and Rodriguez -Navarro, 2001). Our aforementioned obtained results using the E. Coli mutant defective in K+ uptake, indicate that the CHX cDNAs used in the present study are implicated in K+ transport, and thus, further elucidation on their transport characteristics was required before proceeding with their study in planta. To achieve this, we used distinct defective S. cerevisiae mutants in their K+ and/or Na+ transport systems. We used the trk1 trk2 mutant (W∆6) defective in its K+ uptake systems (Haro and Rodríguez-Navarro 2003) to test any possible K+ and Na+ influxes promoted by our cloned CHX cDNAs. Also, we used the ena1-4 nha1 mutant (B3.1) defective in its K+ and/or Na+ efflux systems to test if they promote K+ and/or Na+ efflux under high K+ or Na+ conditions (Bañuelos et al. 1998). Finally, any possible roles in regulating internal K+ and pH homeostasis were studied using ena1-4 nha1 kha1 mutant (LMB01) (Maresova and Sychrova 2005). Although the GFP localization of PpCHX2p in W∆6 yeast mutant was present in the plasma membrane (Fig. 11), it wasn’t exclusively confined to it as in the case of P. Patens protoplasts (Fig. 12). Thus, we wanted to test the possibility of any existing activity of the PpCHX2 transporter in the plasma membrane. The trk1 trk2 W∆6 mutant has been routinely used in the past years to characterize several high affinity K+ or Na+ transporters such as HAK (Garciadeblás et al. 2007, Benito et al. 2012). Our results showed no complementation of any of the P. Patens CHX transporters used in our study either in high or low affinity K+ or Na+ uptake. It is worth noting that W∆6 mutant has the inconvenient that it is only successfully complemented with plasma membrane plant transporters, as described in many previous works. This was also clearly shown in the CHX family where the plasma membrane AtCHX13 transporter successfully suppressed the defective growth of trk1 trk2 mutant (Zhao et al. 2008) while the endomembrane transporter AtCHX17 did not (Cellier et al. 2004, Maresova and Sychrova 2006). Many explanations, although non-conclusive, can be considered to explain why PpCHX2 is inactive at the plasma membrane. These might include a faulty protein processing, missing components required for the transporter activation, or retention of the majority of the protein in internal organelles such as the endoplasmic reticulum. Based on our results, it is risky to draw any conclusions regarding these possibilities. Another disadvantage when using the W∆6 mutant, is that it is unlikely that a low affinity transporter would suppress its defect, as it maintains a very rapid intrinsic low affinity 131

K+ uptake (Santa-María et al. 1997). This could explain why we were unable to detect any differences in the low affinity range either (Fig. 17). However, we cannot negate for sure the possibility that these transporters could mediate low affinity K+ transport. It was reported in previous works that the plasma membrane transporter AtCHX21, in addition to its important role in pollen K+ homeostasis (Lu et al. 2011), may be directly or indirectly involved in Na+ transport in planta (Hall et al. 2006). To test whether any of the PpCHXs of our study are involved in Na+ or K+ efflux, we expressed these in the Na+ and K+ efflux defective mutant (ena1-4 nha1) B3.1, finding no complementation (Fig. 18). This makes the implication of the PpCHXs in either Na+ or K+ efflux, at least in yeast, highly unlikely. However, once again, these results are not conclusive and should be taken with care due to the lack of a clear plasma membrane localization in any of these transporters. On the other hand, both PpCHX1 and PpCHX2 suppressed the defect of the kha1 mutation in S. cerevisiae mutant LMB01 (Fig. 19), which coincides with previously reported results for several Arabidopsis CHX transporters (Maresova and Sychrova 2006, Padmanaban et al. 2007, Chanroj et al. 2011). Kha1 is a K+ transporter residing in the Golgi apparatus membrane of S. cerevisiae. Although a single KHA1 gene mutation does not result in an evident phenotype, nevertheless, when additional mutations in ENA1-4 and NHA1 genes are added to kha1 mutant, an evident growth defect in media with high pH and low K+ is seen (Maresova and Sychrova 2005). Evidence suggests, thus, that K+ is the most preferred substrate for Kha1 tranporter, as the inability of the kha1 mutant to grow at higher pH can be suppressed by the addition of moderate concentrations of K+. The role of Kha1 is believed to be the regulation of intraorganellar K+ and pH homeostasis (Ariño et al. 2010). Also, it is worth mentioning that substrate specificity of PpCHX1 and KHA1 seem to be similar, in contrast to when comparing KHA1 with PpCHX2 or PpCHX2.1. When predicted in terms of the observed phenotype of yeast cells growing in the presence of Na+ (Fig. 21); PpCHX1 seems to transport Na+ similar to KHA1 (Maresova and Sychrova 2005, Ariño et al. 2010). Assuming that the KHA1 gene of S. cerevisiae encodes a K+/H+ antiporter, a tentative interpretation of our results obtained with the heterologous expressions (Fig. 13 and 19) would be that CHX transporters mediate K+-H+ symport in E. coli and K+/H+ antiport in S. cerevisiae. However, an alternative explanation would be that K+-H+ 132

symporters and K+/H+ antiporters fulfill similar functions in endosomal compartments and that these two mechanisms show a certain degree of functional redundancy and reciprocal substitution. This means that a K+-H+ symporter could substitute for KHA1, even if KHA1 mediates K+/H+ antiport (see models of action below). This reasoning would not only apply to CHX transporters but also to PpHAK2, which also suppresses the kha1 mutation of S. cerevisiae (Haro et al. 2013). The basis of this notion is that these two mechanisms participate in the pH control of organelles.

5.3. Proposed models of action for PpCHXs NHX, HAK, and CHX transporters are probably present in the membrane of most organelles (see below in section 5.4). Therefore, the possibility that the functional mechanisms associated to these transporters might be either K+-H+ symport or K+/H+ antiport raises the question about how these two mechanisms can participate in the pH control of the organellar lumen. In the most likely model the organelle pH is established by the steady state that results from H+ pumping into the organelle and return to the cytosol, in parallel with K+ and Cl- conductances (Demaurex 2002, Paroutis et al. 2004, Casey et al. 2010, Ohgaki et al. 2011). H+ pumping and a parallel influx of anions would decrease the organelle pH to very low values but the effective control of the pH requires the return of H+ to the cytosol. H+ pumping is mediated by the electrogenic V- ATPase while the return of H+ can be mediated by multiple systems: H+ passive leaks and fluxes coupled to Cl- or K+ fluxes. In plant cells a specific type of pyrophosphatase might cooperate with the H+ pump V-ATPase (Segami et al. 2010). The question raised above refers to K+ coupling and can be addressed with a simple model including the pump, the coupled H+ efflux, and a K+ channel, if necessary (Fig. 30). Assuming that the membrane potential drives all the other movements, a simple calculation shows that both a K+/H+ antiporters and a K+-H+ symporter are similarly effective to return H+ from the organelle lumen to the cytosol, assuming the existence of a K+ channel for K+ recirculation. If the equilibrium is reached, which is the limit of the gradient that can generate the system, the organelle pH could be higher than the cytosolic pH (ΔpH would be 1 for a ΔΨ of -60 mV). It is worth noting that the three couplings: antiport plus channel, symport plus channel, and antiport plus symport would be similarly effective to return H+ to the cytosol (Fig. 30) but they would be associated to different K+ contents. 133

Fig. 30. Alternative models of action for PpCHXs. Coupling of: a K+-H+ symporter with a K+ channel (a), a K+/H+ antiporter with a K+ channel (b), and a K+-H+ symporter with a K+/H+ antiporter (c). The equations for the systems in equilibrium are used to calculate the ∆pH that the system can attain. ∆Ψ denotes the membrane potential, negative in the cytosolic side; for calculations, 2.3 RT/F = 60 mV.

5.4. Functional analysis of PpCHXs in Physcomitrella patens To analyze the function of PpCHX1 and PpCHX2 in plant cells, we disrupted these genes and also constructed the ∆Pphak1 ∆Ppchx2 double mutant. PpCHX1 localized to the Golgi complex and ∆Ppchx1 plants showed no growth defects when growth was tested in a large variety of conditions of pH, and K+, Na+, and Ca2+ concentrations. This suggests that the function of PpCHX1 may be redundant with other transporters. As already discussed, the mechanism of the redundant transporters might be either K+-H+ symport or K+/H+ antiport. PpHAK3 also localizes to Golgi (Haro et al. 2013), which makes it a candidate substitute of PpCHX1. Most likely there are also other candidates. In Arabidopsis, for example, NHX5 and NHX6 are associated with

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Golgi and TGN (Bassil et al. 2011) and two NHX transporters in P. patens show high sequence homology with AtNHX5 and AtNHX6 (Chanroj et al. 2012). The Golgi complex is an important metabolically active organelle performing important functions in the cell including modifying, sorting, and packaging protein and lipid macromolecules for cell secretion or use within the cell. A strict and precise K+ and pH homeostasis is crucial for the functioning of this organelle, thus, it might be possible that CHX, HAK, and NHX transporters are working in parallel in the Golgi complex performing similar functions. Consequently, due to this possible functional redundancy, it was expected beforehand that single ∆Ppchx1 knockout mutant plants of P. patens would hardly display obvious growth phenotypes even under stress conditions. In the case of ∆Ppchx2 plants, no defects were found that connect CHX2 to K+ (or Rb+) uptake and the same conclusion applies to ∆Pphak1 ∆Ppchx2 plants as well (Fig. 26). Furthermore, the involvement of PpCHX2 in K+ uptake through the plasma membrane can be ruled out considering the phenotype of ∆Ppchx2 plants regarding its morphological appearance under normal or stress conditions, in addition to the simple Rb+ influx tests we carried out (Fig. 26 and 27). On the other hand, when ∆Ppchx2 plants were exposed to Rb+ they showed an increased Rb+/K+ ratio but the same K+ loss, which implies a higher cellular retention of Rb+. Considering that Rb+ influx was not increased by the mutation, the increased Rb+ content of ∆Ppchx2 plants must be the consequence of either a slower Rb+ efflux through the plasma membrane or a slower vacuole-to-cytosol Rb+ transfer, which results in a higher vacuolar Rb+ retention in both cases. Because we did not detect differences in K+ contents, and the differences in Rb+ contents between ∆Ppchx2 and wild-type plants were small, the most likely hypothesis is that PpCHX2 mediates K+ and Rb+ movements in parallel with other transporters that exhibit a higher K+/Rb+ discrimination either in the plasma membrane or in the tonoplast. Because PpCHX2-GFP localized to the tonoplast and plasma membrane, either or both of the functions proposed above for PpCHX2 are possible. The functional difference between these two is mechanistic because Rb+ efflux (K+ efflux in physiological conditions) across the plasma membrane must be Rb+/H+ antiport while vacuole-to-cytosol Rb+ transfer (K+ transfer in physiological conditions) must be either Rb+ uniport or Rb+-H+ symport assuming that the vacuole has low pH, high K+ content,

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and a weak membrane potential that is positive with reference to the cytosol. Considering these observations and the previous discussion about the mechanism of CHX transporters, the most likely possibility is that PpCHX2 mediates K+ (or Rb+)-H+ symport in the tonoplast. If this hypothesis is true, it is highly probable that PpCHX2 contributes to the cell K+ homeostasis by supplying the cytosol with K+, if it were functional in tonoplast. In all cases, it is surprising to detect an evident effect by knocking-out one single vacuolar transporter gene, bearing in mind the redundancy of protein function in plants as well as the enormous adaptive plasticity of plant responses (Maathius 2010). Indeed, other transporters (like members of HAK/KUP/KT family) or channels might fully substitute PpCHX2 for K+ transport, however in this case, only partially for Rb+ transport. Unfortunately, taking into account the large volume of the vacuole, the technical tools to distinguish from a slightly higher Rb+ content in the cytosol or in the vacuole between mutant and wild type plants are practically inexistent. Therefore, at the current level of knowledge, a precise conclusion cannot be reached and it is doubtful that it can be reached from biochemical experiments. Most likely, the identification of the functions of both PpCHX1 and PpCHX2 will come from a genetic approach, by constructing double or triple mutants that show clear defects. Nevertheless, functional redundancy is a common phenomenon in transporters that most probably will make these future studies a difficult task. This phenomenon was clearly demonstrated in the case of CPA2 family members AtCHX21 and AtCHX23, where no apparent phenotype is present in their single mutants but highly evident in double mutants, although both localizing to different membranes (Lu et al. 2011). This also appears to be the case in KEA transporters of Arabidopsis, where kea1 kea2 double mutant shows a clear phenotype in contrast to kea1 or kea2 single mutants (Kees Venema, Personal communication). Functional redundancy in other transporter families is not uncommon either, as in the case of NHX1 and NHX2 (Barragán et al. 2012). It is unlikely, thus, that P. patens double mutant ∆Ppchx1 ∆Ppchx2 might show a clear phenotype, given that PpCHX3 and PpCHX4 might also be playing a role in such redundancy. Moreover, the problem of this approach is that the number of “non-CHX” transporters that can mediate K+/H+ exchange and K+/H+ exchange or K+-H+ symport is high, and the possibility of obtaining triple or quadruple mutants may not be always possible. As discussed above, the presence of HAK and NHX transporters in endomembranes might substitute 136

PpCHX functions, but also if PpCHX2 were functional in plasma membrane as a K+/H+ antiporter, it might be replaced by the PpENA1 K+ ATPase in Physcomitrella (Fraile- Escanciano et al. 2009). Thus, this complicates even more the detection of any abnormal phenotype, even when multiple mutations are achieved. In addition to the previous discussion about NHX transporters in the Golgi complex, P. patens have five NHX transporters showing high sequence homology to the NHX1-4 transporters of Arabidopsis (Chanroj et al. 2012), which play vital K+/H+ exchanges in vacuoles (Barragan et al. 2012, Chanroj et al. 2012). All in all, it is apparent that K+ homeostasis in plants is extremely complex and several transporter families are involved in the pH homeostasis of organelles by mediating either K+/H+ antiport or K+–H+ symport. Although these systems might not be essentially redundant considering their functional conditions, they might replace each other in mutant plants.

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6- CONCLUSIONS

6. Conclusions 1. The CHX family of transporters has suffered numerous events of duplication and diversification during the course of evolution of non-vascular plants into flowering plants. Physcomitrella patens has four CHX genes; two of them are expressed at normal levels (PpCHX1-2) while the remaining two show very low levels of expression (PpCHX3-4). 2. Despite the phylogenetic distance between Arabidopsis and P. patens, the functional basis of CHX transporters in both species seems to be very similar. As in the case of other Arabidopsis CHX transporters, PpCHX1 localizes to the Golgi complex while PpCHX2 localizes to vacuolar compartments and plasma membrane. 3. Both PpCHX1 and PpCHX2 are implicated in K+ transport. They successfully rescued the defective phenotype of yeast mutants unable to grow at low K+ and alkaline pH, and also rescued the phenotype of E. coli mutants defective in their K+ influx transporters. 4. Experiments carried out in E. coli mutants strongly suggest that PpCHX1 and PpCHX2 mediate “active” K+ uptake, which might be a K+-H+ symport. K+-H+ symporters and K+/H+ antiporters are both effective for controlling the pH of the organellar lumen. 5. The in planta studies carried out in this thesis suggest the implication of PpCHX2 in transporting Rb+ from either the vacuole to the cytosol or from the cytosol to the external medium. Our knowledge on the function of PpCHX1 is still lacking. 6. Taken together, present and other studies about CHX, HAK, and NHX transporters in P. patens and Arabidopsis provide compelling evidence about the existence of a complex series of K+ transporters in plant organelles. Although these systems might not be essentially redundant considering their functional conditions, they might replace each other in mutant plants. 7. Future works on the construction of double mutants involving two different transporter families seems the only way to unravel the complexity of the individual functions of endomembrane K+ transporters in plant cells.

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