molecular basis of salt tolerance in physcomitrella patens...
TRANSCRIPT
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
i
To my family
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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
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“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!
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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
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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.
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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
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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
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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 Grams 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 Vmax 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
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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 Vmax 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
1. INTRODUCTION
3
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+
4
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.
5
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
9
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
10
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(SO4)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
11
these different groups have diverse physicochemical and biological properties besides
having high electrolyte content. Although some agronomic and engineering practices
may address this form of salinity, genetic improvement of crops provides the main 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
man-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
12
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,
exposure of plant cells to the low water potential of a saline environment (Ψow) results
in the equilibration of the internal cell water potential (Ψiw) by cell-water loss and an
accompanying decrease of the cell osmotic potential (Ψiπ) and turgor (Ψp) according to
the equation Ψow = Ψi
w = Ψiπ + 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
13
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 non-
selective 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
some important anions such as NO3− and PO4
−3, as their transport is energized by the
H+-ATPase activity (Shabala and Cuin 2008).
14
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
in (Ca2+)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
15
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
16
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.
17
On the other hand, the implication of some 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
18
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
19
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
20
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
21
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
22
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
23
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
nutrients like K+, Ca2+, 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
24
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
25
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
26
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
27
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
28
importance of K+ retention and maintaining a high cytosolic K+/Na+ ratio and
homeostasis under normal and saline conditions.
29
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 time. 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
30
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
31
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
32
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
33
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+
34
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 Damon 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
35
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
36
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 may 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
37
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,
38
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 non-
selective 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
39
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
40
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
41
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
42
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).
43
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
44
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 non-
selective 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.
45
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
46
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 may 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.
47
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,
48
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.
49
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).
50
Table 1. Scientific classification of Physcomitrella patens.
Taxon Name
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
51
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
52
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
53
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
57
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
DH5α
F- endA1- hsdR17(rK- mK+) supe44
thi-1 lambda- recA1 gyrA96 realA1
80dlacZAM15
(Hanahan 1983)
TKW4205 thi rha lacZ nagA recA Sr::Tn10
∆kdpABC5 trkA405 kup1
(Schleyer and Bakker
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.
58
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
W∆6 Mat a ade2 ura3 trp1 trk1∆::LEU2 trk2∆HIS3 (Haro and Rodríguez-
Navarro 2003)
B3.1 ena1∆::HIS3::ena4 nha1∆::LEU2 (Bañuelos et al. 1998)
LMB01
MATa ade2-1 can1-100 his3-11,15 leu2-3,112
trp1-1 ura3-1
mall0 ena1Δ::HIS3::ena4Δ nha1Δ::LEU2
kha1Δ::kanMX
(Maresova and
Sychrova 2005)
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.
59
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
Bacto-agar 2% Yeast
extract 1% (w/v) H3PO4 8 mM
Glucose 2% Peptone 2% (w/v) MgSO4 2 mM
Yeast nitrogen base
w/o amino acids or
ammonium
0.7% Glucose 2% (w/v) CaCl2 0.2 mM
(NH4)2SO4 0.5% Bacto-
agar 1.5% Arginine 10 mM
Trace
elements 1 ml
Vitamins 1 ml
Glucose 2% (w/v)
60
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).
61
Table 7. Media used for growing Physcomitrella patens.
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.
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
CoCl2·6H2O 0.055 mg/l
KI 0.028 mg/l
Na2MoO4·2H2O 0.025 mg/l
1 ml l-1
1 ml l-1
H3PO4 1.8 mM 1.8 mM
NH4 tartrate (4.6 mM)
pH 5.8 6.5
62
Table 8. Stock solutions for BCD medium.
Solution B (for 1 L) Solution C (for 1 L) Solution D (for 1 L)
MgSO4 7H20
………….……25 g
KH2PO4
….……………….25 g
pH adjusted to 6.5 with 4M
KOH
KNO3
………………101 g
FeSO4.7H20
.………………...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.
63
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.
64
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.
65
Table 9. The plasmids used and their characteristics.
Plasmid Characteristics Reference
pCR2.1-TOPO Bacterial multicopy Plasmid for PCR product
cloning. Amp R km R Invitrogen
pBAD24
Bacterial multicopy plasmid of inducible
expression. pBAD promoter, araC regulator
dependant on arabinose concentration; Amp R
(Guzmán et al.
1995)
pYPGE15
Multicopy plasmid for expression in yeast.
PGK promoter, CYC terminator, URA marker,
Amp R
(Brunelli and
Pall 1993)
pMF6-GFP
Plasmid of expression in plants for obtaining
gene fusions with GFP. 35S promoter, adh1
intron, mGFP5 gene, 3´NOS terminator, AmpR
(Rubio-Somoza
et al. 2006)
pTN186
Plasmid used for the construction of PpCHX1
gene disruption. AmpR for E.coli and HygR for
Physcomitrella. Hygromicin cassette under the
CamV35S promoter and terminator.
Dr.Hasebe, Dr.
Nishiyama and
Dr. Hiwatashi,
Japan
P35S-Zeo
Plasmid used for the construction of PpCHX2
gene disruption. AmpR for E.coli and ZeoR for
Physcomitrella. Zeocin cassette under the
CamV35S promoter and terminator.
Dr.Hasebe, Dr.
Nishiyama and
Dr. Hiwatashi,
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.
66
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
M, CTAB 2%, Na2EDTA 20 mM, PVP-40 2%) supplemented with 1 mg ml-1 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
67
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).
68
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 re-
suspended 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.
69
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.
70
Table 11. Primers used to amplify the different cDNAs.
Primers Sequence 5´-3´ Observations
PpCHX1-ATG1 TATAAGTTCTACAAAGCCTC Cloning of PpCHX1
cDNA in pCR2.1-
TOPO plasmid
PpCHX1-ATG2 CAAGATGGCGGACGCTGTGG
PpCHX1-ATG3 GGCGTGCAAAACTATGTCGG
PpCHX1-STOP TTAGCCAGCCAAGACCTTCGCA
PpCHX1-1R AAAGCTCCCTCTGGTGCAAT Sequencing of PpCHX1
5’ end PpCHX1-2R GCAACGAAAGGCAGAGTGAT
PpCHX1-3R AATGGACATAGCTTGCTTTC
PpCHX1-ATGBamHI2 CGGATCCTAAAAAATGGCGGACGCT
GTGGC Cloning of PpCHX1
cDNA in pYPGE15
plasmid PpCHX1-StopKpnI GGGGTACCTCACACGCGGTGGTCAG
GAGAA
PpCHX1-639 TTGCATGGGTTCTGTTGTGT Primers used to confirm
fidelity of amplification
of the PpCHX1 cDNA
cloned in the different
plasmids used
PpCHX1-1050 TCTGGGTAAAATTCTCGGAAC
PpCHX1-1050- R GTTCCGAGAATTTTACCCAGA
PpCHX1-1107 CAAAGCTTTAACTCTCGGCATC
PpCHX1-1146 ATTGGTGGAGCTTATTGTTCTG
SpeIATG1-PpCHX1 TACTAGTAGGAGGAGATAAGATGGC
GGACGCTGTG Cloning of PpCHX1
cDNA in pBAD24
plasmid PpCHX1-StopKpnI GGGGTACCTCACACGCGGTGGTCAG
GAGAA
PpCHX2.2 AGTGTTGCAAGATGGCGACCGGA Cloning of PpCHX2
cDNA in pCR2.1-
TOPO plasmid
PpCHX2-stop {2B in fig.
S4} GTTTTCAAGATGGGTCTAGGATC
PpCHX2-1R GAAGACAACAAACGGGCCGAATT
PpCHX2-2R AACGGGCCGAATTTCACATCACC Sequencing of PpCHX2
5’ end
PpCHX2-3R TATTAAGGAGGACAAAGGAGACC
CHX2-int-2 {1A-B in fig.
S4}
TTCTTGAACAGATACCATATCGCCAC
C
Intron removal for
construction of
PpCHX2.1 version
CHX2-int-2R {2A-B in
fig. S4}
TATGGTATCTGTTCAAGAAGCGGATC
AC
PpCHX2-ATG-BamHI GTGGATCCTACACAATGGCGACCGG
AAATGC
Cloning of PpCHX2
cDNA in pYPGE15
plasmid
71
PpCHX2-Stop-KpnI GAGGTACCAGGATCTAGACATGAGG
CGCCGA
PpCHX2-622 TGCCTCCTCGCTCTCGCTGT
PpCHX2-1259 TGACTACTCCCCTTGTGATG
Primers used to confirm
fidelity of amplification
of the PpCHX2 cDNAs
cloned in the different
plasmids
PpCHX2-1715 TCCACAAAACCCAAAGACTC
PpCHX2-1969 GTGATCCGCTTCTTGAACAG
SpeIATG-PpCHX2 TACTAGTAGGAGGAGATAAGATGGC
GACCGGAAAT
Cloning of PpCHX2
cDNA in pBAD24
plasmid
PpCHX2-Stop-KpnI GAGGTACCAGGATCTAGACATGAGG
CGCCGA
PpCHX3-2 GCGAGAAAGCGAAGACGGTG
Verification of
existence of PpCHX3
gene in P. patens
genome
PpCHX3-2R GATGCTGCACAAGTGATTTCG
PpCHX4-2 TCAAGGCGAAATAGCAGAGAC
Verification of
existence of PpCHX4
gene in P. patens
genome
PpCHX4-2R GTGCTGAGAATAATTGGTGGG
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.
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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
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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
KOH (2 M)
H2O (Final volume) 500 ml
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
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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
75
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 time
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 re-
suspended 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
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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
M). Sterilized by filtration 0.22 or
0.45 µm Millipore filters.
pH adjusted to 7.5 with KOH (1 M), and left at
room temperature all night. Sterilized by filtration
0.22 or 0.45 µm Millipore filters.
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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
Tartrate 0.5M
(100x)
5 ml 1 ml 200 µl
Trace elements
(1000x)
0.5 ml 0.1 ml 20 µl
Mannitol 30 g 6 g 1.2 g
Agar (plant
Agar, Duchefa)
0.55% 0.4g (0.4%)
CaCl2 (1 M) 200 µl
Solution sterilized in
autoclave. Once
sterilized, 5 ml of 1 M
CaCl2 is added.
For each DNA
transformed, 3 plates of
20-25 ml were prepared
covered with cellophane
paper discs.
Solution sterilized in
autoclave. Once
sterilized 5 ml
CaCl2 1 M is added.
The regeneration of
protoplasts requires
a higher calcium
concentration (10
mM) than normal
growth (1 mM)
Final volume
taken to 20ml.
Sterilized by
filtration through
0.22 or 0.45 µm
Millipore filters
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
78
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
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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).
80
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
knock-out PpCHX1-5’-HindIII AAGCTTAGCAAAATGAACGGGCACATCTCC
PpCHX1-3’-XbaI-2 TATTCTAGATATTTTGGTTGGCCGATCGAG Cloning of the 3´
fragment in pTN186
for PpCHX1 gene
knock-out PpCHX1-3’-SacI-2 TTGAGCTCACAGATCCATTCCATCAAAACA
PpCHX2-5’EcorI2 TGAATTCTATCCCCTTGCCACACGCC Cloning of the 5´
fragment in p35S-
Zeo for PpCHX2
gene knock-out PpCHX2-5’XhoI2 ACTCGAGAATTTCATCACCTGCCTAC
PpCHX2-2-3’(1680) AACTGACAAACGAGCCGCAATGATC Cloning of the 3´
fragment in p35S-
Zeo for PpCHX2
gene knock-out PpCHX2-3’ NotI-2 GCGGCCGCGACCAATCCACTTCACTAAA
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
81
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/).
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Table 16. Primers used for the verification of Physcomitrella knockouts.
Primers Sequence 5´-3´ Observations
PM35S-1R {2D in
fig. S2-A&B} GCCCTTTGGTCTTCTGAG
Anneals to 35S promoter of
p35S-Zeo and pTN186
cassettes of resistance to
antibiotics
35SPA-1 {1C in fig.
S2-A&B} AGGGCCATGAATAGGTCT
Anneals to 35S terminator
of p35S-Zeo and pTN186
cassettes of resistance to
antibiotics
PpCHX1-ATG3 {1A
& 1D in fig. S2- A} GGCGTGCAAAACTATGTCGG
5´ end verification of
PpCHX2 knock-out
PpCHX1-STOP {2B
& 2C in fig. S2- A} TTAGCCAGCCAAGACCTTCGCA
3´ end verification of
PpCHX2 knock-out
PpCHX1-ATG3 {1B
in fig. S2- A} GGCGTGCAAAACTATGTCGG Amplification of knocked-
out fragment of PpCHX2
gene PpCHX1-1R {2A in
fig. S2- A} AAAGCTCCCTCTGGTGCAAT
PpCHX2-5’-G1 {1A
& 1D in fig. S2- B} CAGCGAACGAACAAACCAAC
5´ end verification of
PpCHX1 knock-out
PpCHX2-3’-GR {2B
& 2C in fig. S2- B} CAAACTGCCAAAACATATGA
3´ end verification of
PpCHX1 knock-out
PpCHX2-4R {2A in
fig. S2- B} CAATGACCCGAGGCTGTTTC Amplification of knocked-
out fragment of PpCHX1
gene PpCHX2- (1680) {1B
in fig. S2- B} AACTGACAAACGAGCCGCAATGATC
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
83
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
with GFP gene PpCHX1-NoStop-BamHI TTGGATCCGCACTCTGTGGTCAGGAGAAGAC
AA
PpCHX2-ATG-BamHI GTGGATCCTACACAATGGCGACCGGAAATGC Cloning of the
PpCHX2 in phase
with GFP gene PpCHX2-NoStop-BamHI TTGGATCCGTACATGAGGCGCTGACTTC
84
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 H2Omq (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
85
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)
86
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 H2Omq 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 ClustalW2 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
89
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
90
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).
91
Fig. 5. PpCHX1-4 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.
PpCHX1 ----MADAVACKTMSATSNGVWQGDVPVHFALPLLIVQIVLVLAITRALAFVLKPLKQPR 56 PpCHX2 MATGNATVTTCKTMPASSNGVWQGDIPIHFALPLLIIQICIVLTITRVLAALFKPLKQPR 60 PpCHX3 -----MASENVTDVSVTSSGVVAGDNPLHHTLPLLIIQVMVIITVSRCVAVLLRPLKQPR 55 PpCHX4 -----MTSNQTSDVSVTSRGVLAGDDPLHHALLLLIIQVIIIICLTRFLALLLRPLKQPR 55 AtCHX18 MAT-NSTKACPAPMKATSNGVFQGDNPIDFALPLAILQIVIVIVLTRVLAYLLRPLRQPR 59 AtCHX20 ------MPFNITSVKTSSNGVWQGDNPLNFAFPLLIVQTALIIAVSRFLAVLFKPLRQPK 54 : .:* ** ** *:..:: * *:* ::: ::* :* :::**:**: PpCHX1 VVAEIIGGILLGPSAFGRNKDYLHTIFPHESVIILEVFADMGLLFFLFMVGLELDMTQIR 116 PpCHX2 VIAEVVGGILLGPSALGKNTAYIANIFPKQSVIILEVFAQMGLIFFLFMVGLELDIRQIR 120 PpCHX3 VVAEILGGILLGPTAFGHIPGFTKNIFPESSLPVLETVAEVGLIFFLFLVGLELDTKQIR 115 PpCHX4 VVAEILGGIILGPTGFGSIPGITDTIFPESSLTVLDTAANVGLIFFLFLVGLELNIKTIV 115 AtCHX18 VIAEVIGGIMLGPSLLGRSKAFLDAVFPKKSLTVLETLANLGLLFFLFLAGLEIDTKALR 119 AtCHX20 VIAEIVGGILLGPSALGRNMAYMDRIFPKWSMPILESVASIGLLFFLFLVGLELDLSSIR 114 *:**::***:***: :* :**. *: :*: *.:**:****:.***:: : PpCHX1 KTGKQAMSIAAAGITLPFVAGVGVSFVLHLTIAPEGA---FGPFLVFMGVAMSITAFPVL 173 PpCHX2 RTGFQALVISAAGIAVPFSTGVGVSFVLLNTIAGDVK---FGPFVVFMGVAMSITAFPVL 177 PpCHX3 RSGVMTLWLSAAGIILPFGLGAVAAFIIFKLQNSLH-HPNFGAFVLFLGVALSVTAFPVL 174 PpCHX4 KSGVISLWMAAAGIIFPFGLGSIVACLISKLLPQMDSHHSFGVFTMFLGVALSITAFPVL 175 AtCHX18 RTGKKALGIALAGITLPFALGIGSSFVLKATISKGVN---STAFLVFMGVALSITAFPVL 176 AtCHX20 RSGKRAFGIAVAGITLPFIAGVGVAFVIRNTLYTAADKPGYAEFLVFMGVALSITAFPVL 174 ::* :: :: *** .** * : :: * :*:***:*:****** PpCHX1 ARILAERKLLTTEVGQLAMSAAAVNDVVAWVLLALAVALSGSG-------RSPAIVAWVL 226 PpCHX2 ARILAERKLLTTEVGQLAMSVAAVDDVVAWCLLALAVALTGTN-------TKPSVVAWVL 230 PpCHX3 ARILTERKLLHTDIGQMAMAAAAINDVVAWVLLALAVAITSSG-------SDPLVALWVL 227 PpCHX4 ARILAERKLLNTEIGQMAMSAAAINDIVAWILLALALALTNSG-------STPVVAIWVL 228 AtCHX18 ARILAELKLLTTEIGRLAMSAAAVNDVAAWILLALAIALSGSN-------TSPLVSLWVF 229 AtCHX20 ARILAELKLLTTQIGETAMAAAAFNDVAAWILLALAVALAGNGGEGGGEKKSPLVSLWVL 234 ****:* *** *::*. **:.**.:*:.** *****:*::... * : **:
MI
MII MIII
MIV MV
MVI
92
PpCHX1 LCGIAFCLAIFLVVQPCMQWVAHR-SPDNEPVKEYIVALTLLCVLVAGFCTDAIGVHSIF 285 PpCHX2 LTGIAFIITMFVVVQPVMRWVATR-SADNEPVKEILVCLTFAGVLIAAFTTDLIGIHAIF 289 PpCHX3 LLGTVFIVFMLLVISPITYALAHH----SEPATESVIAMTLMLVLGAAFITDLIGIHVIF 283 PpCHX4 LLGVAFSAFMFVVVSPVMYALANY----RELPPEPVVAMTLVIVLGSAFFSDLIGIHVIF 284 AtCHX18 LSGCAFVIGASFIIPPIFRWISRR-CHEGEPIEETYICATLAVVLVCGFITDAIGIHSMF 288 AtCHX20 LSGAGFVVFMLVVIRPGMKWVAKRGSPENDVVRESYVCLTLAGVMVSGFATDLIGIHSIF 294 * * * .:: * :: : * :. *: *: ..* :* **:* :* PpCHX1 GAFLFGLVIPKEGPFA----AALVEKLEDFVSILLLPLYFASSGLKTNIGAIHSAQSFGL 341 PpCHX2 GAFLFGLIVPKDGPFA----VALVEKIEDFISILMLPLYFASSGLKTNIGAIKTGQSFGL 345 PpCHX3 GAFICGLIIPKDGPFT----AILIEKVEDYVSVLFLPLYFAASGLKTHLSAINNGTAVLI 339 PpCHX4 GAFICGLIVPKDGSFAGTLTSTMIEKVEDYVSVLFLPLYFAISGLKTHLSEVNNGTAVLI 344 AtCHX18 GAFVVGVLIPKEGPFA----GALVEKVEDLVSGLFLPLYFVASGLKTNVATIQGAQSWGL 344 AtCHX20 GAFVFGLTIPKDGEFG----QRLIERIEDFVSGLLLPLYFATSGLKTDVAKIRGAESWGM 350 ***: *: :**:* * ::*::** :* *:*****. *****.:. :. . : : PpCHX1 LVLVISVACLGKILGTFAAAKACRVDARKALTLGILMNTKGLVELIVLNIGLDRGVLNSE 401 PpCHX2 LVLVIAVACFGKMCGVFLAATASKVNPRKALTLGVLMNTKGLVELIVLNIGKDRGVLNEE 405 PpCHX3 LFLVIASACVGKVLGTFIVAKVWGVGSRKAIALGFLMNTKGLVELIILNIGLSKGVLNEE 399 PpCHX4 LFMVIATACIGKVMGTLIVAKIWGVENSKALALGFLMNTKGLVELIVLNIGLSKRVINQE 404 AtCHX18 LVLVTATACFGKILGTLGVSLAFKIPMREAITLGFLMNTKGLVELIVLNIGKDRKVLNDQ 404 AtCHX20 LGLVVVTACAGKIVGTFVVAVMVKVPAREALTLGFLMNTKGLVELIVLNIGKEKKVLNDE 410 * :* ** **: *.: .: : :*::**.***********:**** .: *:*.: 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
MVII MVIII
MIX
MX MXI
MXII
93
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”
94
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.
95
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
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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).
97
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
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
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.
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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.
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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,
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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. We 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
106
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, 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 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. Growth 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 (nmol/mg) 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
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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 small, 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.
5. Discussion
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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.
members 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
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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.
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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 Vmax 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
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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
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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
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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
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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+
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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.
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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
135
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,
136
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
137
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.
138
6- CONCLUSIONS
141
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.
142
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