2

3

Index

Abstract in English 5

Riassunto in Italiano 7

State of the art and Objectives

Chapter 1.1 The Plant Vacuole 11

Chapter 1.2 Ion transport in plant vacuoles 12

Chapter 1.3 Flavonoid biosynthesis, localization and transport 25

Chapter 1.4 Polyunsaturated fatty acids 30

Chapter 1.5 Objectives 31

Materials and methods

Chapter 2.1 Methods for localization of flavonoids in plant leaves 35

Chapter 2.2 Electrophysiology: the patch-clamp methodology 38

Results

Chapter 3.1 Fluorescence imaging and flavonoid localization 57

Ch. 3.2 Macroscopic Slow Vacuolar currents 61

Ch. 3.3 Inhibition of the SV channel by Naringenin added at

the cytosolic side 63

Ch. 3.4 Quantitative analysis of SV channel inhibition

by Naringenin 66

Ch. 3.5 Voltage dependence characteristics of the SV channel

in the presence of cytosolic Naringenin 69

Ch. 3.6 Naringenin effects on the SV single channel conductance 72

Ch. 3.7 Simultaneous Fluorescence measurements

and patch-clamp recordings 75

Ch. 3.8 pH and DTT effects on the modulation of

the SV channel by Naringenin 80

Ch. 3.9 Effects of vacuolar Naringenin on the SV channel 82

Ch. 3.10 Naringin does not inhibit the SV currents 83

Ch. 3.11 Other cytosolic flavonoids also inhibit the SV currents 85

4

Ch. 3.12 Effects of Naringenin on the SV channel from

transparent testa mutant plants 87

Ch. 3.13 Effects of polyunsaturated fatty acids on the SV currents 89

Ch. 3.14 Voltage dependence characteristics of the SV channel

in the presence of cytosolic Arachidonic Acid 92

Ch. 3.15 Quantitative analysis of SV channel inhibition

by Arachidonic Acid 95

Ch. 3.16 Arachidonic Acid effect on the SV

single channel conductance 97

Discussion 99

References 107

Acknowledgments 127

Abbreviations:

AA Arachidonic Acid

ABC transporter ATP-Binding Cassette transporter

At Arabidopsis thaliana

CICR Calcium Activated Calcium Release

DHA DocosaHexaenoic Acid

DMSO DiMethyl SulfOxide

ER Endoplasmic Reticulum

Fou2 Fatty acid oxygenation up regulated 2

AtGt Arabidopsis thaliana Glucosyl transferase

GST Glutathione S Transferase

MATE Multidrug And Toxic Efflux

MRP Multidrug Resistance associated Protein

NAADP Nicotinic Acid Adenine Dinucleotide Phosphate

Nar Naringenin

PUFAs PolyUnsaturated Fatty Acids

QCT Quercetin

SV channel Slow Vacuolar channel

TPC1 Two Pore Channel 1

tt transparent testa

YFP Yellow Fluorescent Protein

5

Abstract in English

The Slow Vacuolar channel, ubiquitous in plant cells, has been investigated for almost 25

years; nevertheless its role remains still elusive. In search of compounds that are able to

change its properties providing useful information for its physiological role, we

investigated the modulation of the SV channel by the flavonoid Naringenin (Nar) using

the patch-clamp technique. Experiments were conducted on vacuoles obtained from

carrot roots and from mesophyll cells of Arabidopsis thaliana leaves. Firstly, the

presence of non-glycosylated Nar in the cytoplasm was verified by localization studies,

then patch-clamp experiments in the excised patch configuration revealed that Nar

inhibits the SV channel in a reversible dose-dependent manner, shifting the threshold of

activation of the SV currents towards more positive membrane potentials. Cytosolic Nar

did not change the amplitude of the single channel conductance but decreased the mean

open time of the channel. We also demonstrated that Nar inhibits equally well the

permeation of both K+ and Ca

2+ through the SV channel. Flavonoids sharing structural

similarities with Nar, induced a comparable inhibition of the SV currents. Inside-out

patch experiments performed in vacuoles from Arabidopsis thaliana mesophyll cells and

carrot roots revealed that Nar inhibits the SV channel also when added at the luminal

side. Naringin, the glycosylated form of Nar, was unable to change the channel activity

when added at both sides of the membrane.

Finally, in accordance with a higher hydrophobicity of the flavonoids at acidic pHs, Nar

inhibition increases decreasing the pH of the cytosolic bath solution. All these evidences

point to the possibility that flavonoid interaction with the SV channel is mediated by the

membrane phospholipids.

To further investigate this possibility we studied the interaction of polyunsaturated fatty

acids (PUFAs) with the SV channel in vacuoles from carrot roots. Similarly to Nar, both

Arachidonic Acid (AA) and Docosahexaenoic Acid (DHA) added at the cytosolic side

induced a dose-dependent reversible decrease in the SV channel activity. Also AA shifted

the SV activation to more positive voltages without changing the amplitude of the single

channel conductance.

These studies suggest that important endogenous molecules, such as flavonoids and

PUFAS, are able to change the properties of the slow vacuolar channel, possibly through

a lipid mediated interaction. This biophysical characterization opens new perspectives to

investigate the properties of this enigmatic channel in different plant species and tissues.

6

7

Riassunto in Italiano

Il canale Slow Vacuolar (SV), un canale ubiquitario in sistemi vegetali, è studiato da

circa 25 anni ma il suo ruolo fisiologico rimane tuttora elusivo. Alla ricerca di molecole

che potrebbero essere in grado di modularne le proprietà fornendo informazioni utili alla

comprensione del suo ruolo fisiologico, mediante metodologia di “patch-clamp abbiamo

studiato le modificazioni indotte da Naringenina (Nar) su detto canale in vacuoli estratti

dalla radice di carota o dal mesofillo della foglia di Arabidopsis thaliana. La presenza nel

citoplasma di Naringenina, un flavonoide non-glicosilato, è stata preventivamente

verificata mediante studi di localizzazione. Successivamente esperimenti di patch-clamp

nella configurazione “excised-out” hanno dimostrato che Nar inibisce il canale SV in

modo reversibile e dipendente dalla concentrazione di flavonoide mentre la soglia di

attivazione del canale viene spostata verso potenziali più positivi.

L’aggiunta di Nar alla soluzione citosolica non cambia l’ampiezza di singolo canale ma

diminuisce aumenta il tempo di vita media del canale in stato aperto. Abbiamo anche

dimostrato che Naringenina inibisce allo stesso modo sia il trasporto di K+ che di Ca

2+

attraverso il canale SV. Inoltre i flavonoidi con struttura simile a quella di Naringenina

inducono inibizioni confrontabili delle correnti SV. Esperimenti effettuati in

configurazione “inside out” su vacuoli ottenuti da Arabidopsis e da carota hanno

dimostrato che Nar inibisce il canale SV anche quando essa viene aggiunta dal lato

vacuolare. Invece Naringin, la forma glicosilata della Naringenina, non è in grado di

cambiare l’attività del canale sia che essa venga aggiunta al lato citosolico che al lato

vacuolare. Infine, in accordo con l’aumento di idrofobicità dei flavonoidi utilizzati a pH

acidi, l’inibizione delle correnti SV mediate da Nar aumenta diminuendo il pH della

soluzione citosolica.

Tutte queste evidenze suggeriscono che l’interazione tra flavonoidi e canale SV possa

essere mediata dai lipidi di membrana.

Per sviluppare ulteriormente questa ipotesi di lavoro abbiamo studiato l’interazione di

alcuni acidi grassi polinsaturi (PUFAs) con il canale SV del vacuolo di carota. In maniera

del tutto simile a quanto accade all’aggiunta di Nar, sia l’acido Arachidonico (AA) che

l’acido Docoesaenoico (DHA), aggiunti alla soluzione citosolica, inducono una

diminuzione reversibile e dose-dipendente dell’attività del canale. Come la Naringenina,

anche AA sposta il potenziale di attivazione delle correnti SV verso valori di potenziali

più positivi senza cambiare l’ampiezza di singolo canale.

Questi studii suggerisconoo che alcune importanti molecole endogenee, quali i flavonoidi

ed i PUFA, sono in grado di modifcare le proprietà del canale Slow Vacuolar,

probabilmente attraverso un’interazione mediata dai lipidi di membrana. Questa

caratterizzazione biofisica apre nuove prospettive per lo studio delle proprietà di questo

enigmatico canale in specie vegetali ed in tessuti diversi.

8

9

State of the art and Objectives

10

11

Chapter 1.1. The Plant Vacuole

Most plant cells contain liquid-filled membrane-bound organelles called vacuoles. A

vacuolar membrane called tonoplast surrounds the plant vacuole. In mature plant cells,

the vacuole can occupy from 30% to 90% of the cell volume. Classification of the

vacuoles is based on the nature of their soluble proteins and by the class of aquaporins

present on the tonoplast.

Fig. 1. 1. Schematic representation of an idealized plant cell with a large central vacuole.

Mature plant cells may contain two large vacuoles with different functions (Martinoia et

al. 2007). For example (1) the central vacuole plays a role in the maintenance of turgor

pressure, in cell homeostasis and in the storage of metabolic compounds, (2) defence or

signal compounds may be stored in the vacuoles of some cell types (Marty 1999), (3)

proteins are stored in the vacuoles of reserve tissues of fruits and seeds (Herman and

Larkins 1999), (4) vacuoles can be involved in programmed cell death (Paris et al. 1996).

Transport across the tonoplast is essential for cell homeostasis and to maintain the turgor

pressure which is fundamental in regulation of cell expansion and growth (Maeshima

2001). As the central vacuole occupies most of the space in the cell, it stores all the

12

essential compounds, minerals, nutrients, metabolites and different ions required for cell

homeostasis. Different systems of ion transport were identified in the vacuolar

membrane, which allow the ions to selectively permeate according to their

electrochemical gradients (channels) or at the expenses of other energy sources (active

transporters).

Chapter 1.2. Ion transport in plant vacuoles

1.2.1 Vacuolar Transporters

Fig. 1.2. Schematic representation of the vacuolar transporters (Shiratake and Martinoia

2007).

In plant cells compounds are transported across the tonoplast for storage and some

compounds present in the cytoplasm may reach up to 7 fold higher concentrations in the

vacuole. For example, the pH in the lumen can reach values up to 3 units whereas in the

cytoplasm it remains around 7 units. It is obvious to note that proton (H+) fluxes are

required to maintain this pH difference and two proton pumps, namely a V-ATPase and a

V-PPase responsible for this phenomenon (Sze et al. 1999; Maeshima 2000), are present

in all types of vacuoles (Hedrich et al. 1989). By pumping the protons into the lumen they

not only remove them from the cytosol but also create a voltage difference across the

tonoplast of about -30 mV (cytosol minus vacuolar side). The electrochemical pH

gradient and the voltage difference are used by channels (see channels section below) and

13

secondary active transporters to transfer electrically charged ions, in accordance or

against their chemical or electrochemical potential difference, respectively. It is shown

that a potential difference of -30 mV would allow ~5 fold vacuolar accumulation of

monovalent anions, (such as chloride or nitrate) and ~10 fold accumulation of divalent

anions (such as sulfate) relative to the cytosol concentrations (Martinoia et al. 2007).

Vacuolar H+-ATPase

V-ATPase uses the energy of ATP hydrolysis to pump the protons across a variety of

biological membranes. The existence of a V-ATPase was reported way back in 1962

(Kirshner 1962), in plants in 1973 (Rungie and Wiskich 1973). More recently, it was

found out to be universally present in all the membranes of different internal acidic

organelles in eukaryotic cells (Martinoia et al. 2007).

Fig. 1.3. Schematic representation of a vacuolar H+-ATPase. The V-ATPase complex is

composed of a peripheral domain (V1), which interacts with ATP, ADP and inorganic

phosphate. The V1 domain is a hexamer of A and B subunits. The V0 domain is composed

of a ring of proteolipid subunits (c, c' and c"), adjacent to subunits a and e. This complex

mediates the transport of H+. The V1 and V0 domains are connected by a central stalk,

composed of subunits D and F of V1 and subunit d of V0, and multiple peripheral stalks,

composed of subunits C, E, G, H, and a. From Carraro-Lacroix et al (2009).

Their multi subunit composition was revealed by purification of vacuolar ATPases from

animals, plants and fungi. They are made up of 14 different polypeptides, which assemble

as two major ring structures (Fig. 1.3): (1) a peripheral V1 complex that interacts with

ATP, ADP and inorganic phosphate, and (2) an integral membrane V0 complex

that

14

mediates the transport of H+. In eukaryotic V-type H

+ ATPases, the V1 complex is

invariably present in the cytoplasm. In plants V-ATPase activity is transcriptionally

regulated and it has been demonstrated that post translantional modifications by a WNK

(With No K = lysine) kinase are involved in this regulation (Hong-Hermesdorf et al.

2006). V-ATPase activity is inhibited by bafilomycin (Bowman et al. 1988). For a critical

review on the structure, function and physiological role of V-ATPases see Beyenbach

and Wieczorek (2006).

Vacuolar H+-pyrophosphatase

The V-PPase proton pump uses PPi as its energy source, the compound that is

synthesized in many metabolic reactions. This type of pump is found in archaebacteria,

protozoa, algae, photosynthetic bacteria and plants but not in fungi or mammals

(Martinoia et al. 2007). The molecular identity of a V-PPase from Arabidopsis thaliana

plant was reported by Kieber and Signer (1991). It consists of a single polypeptide with a

molecular mass of about 80 kDa. It has been suggested that the V-PPase activity is

regulated by sodium (Rea and Poole 1985), calcium (Ca2+

) (Rea et al. 1992) and

magnesium (Leigh et al. 1992; Baykov et al. 1993) from the cytosolic side. Two isoforms

of this pump are found in plants: a potassium-dependent type which is moderately

sensitive to calcium and a potassium-independent form that is extremely calcium

sensitive (Belogurov and Lahti 2002). Both types of isoforms require magnesium as a

cofactor. V-PPase activity is essential for maintaining the acidity of the large central

vacuole in plant cells (Maeshima 2000). Generally, its activity is high in young tissues,

but in grape berries it is expressed even in mature plant cells (Terrier et al. 1998). For

reviews on V-PPases see Baltscheffsky et al. (1999), Maeshima (2000) and Gaxiola et al.

(2007).

Calcium ATPase

Calcium can constitute up to 5% of the dry weight of a plant (Broadley et al. 2004). The

large central vacuole is the most important free Ca2+

storage organelle in plant cells

(Martinoia et al. 2007). The free Ca2+

concentration inside the vacuole is around ~ 1000 –

fold higher than in the surrounding cytosol (Bush 1995), where the free Ca2+

concentrations are around 100 nM-200 nM at rest (Wheeler and Brownlee 2008). Ca2+

is

one of the most important second messengers in plant cells; an increase in intracellular

15

calcium is believed to be a major pathway in the plant stress response. The huge Ca2+

concentration differences (between the store and surrounding cytosol) are the basis for

the function of Ca2+

as second messenger (Pottosin and Schonknecht 2007). Ca2+

-

ATPases and Ca2+

/H+ antiporters perform the active Ca

2+ transport into the vacuole

(Sarafian et al. 1992).

The Ca2+

-ATPases are one of the best-characterized ion transport ATPases in the

organisms. The plant Ca2+

-ATPases are divided into two groups: type IIA and IIB. In

Arabidopsis thaliana four isoforms of IIA Ca2+

-ATPase (AtECA) (Geisler et al. 2000)

and six isoforms of type IIB Ca2+

ATPase (AtACA) (Sze et al. 2000) have been cloned.

Ca2+

/H+ Antiporter

The Ca2+

/H+ antiporter is driven by the transmembrane pH gradient generated by the V-

ATPase and V-PPase. This antiporter was reported to be present in various plant species

(Schumaker and Sze 1986; Chanson and Pilet 1987; Berkelman et al. 1994). The first

Ca2+

/H+ antiporter from Arabidopsis thaliana was cloned in 1996 (Hirschi et al. 1996). It

has 11 transmembrane domains and a highly acidic motif between the sixth and the

seventh transmembrane domains.

Na+/ H

+ Antiporter

The Na+/ H

+ antiporter can export sodium (Na

+) from the cytosol to extracellular space as

well as into the vacuole using proton gradients (Blumwald et al. 2000). The first Na+/H

+

exchanger identified in plants was Arabidopsis thaliana AtNHX1 protein (Gaxiola et al.

1999). These are proteins of about 550 residues in length and have 10–12 transmembrane

domains with a hydrophilic C-terminal tail in the luminal side (Yamaguchi et al. 2003). It

has been shown that the Na+/ H

+ antiporter plays a major role in salt tolerance (Blumwald

and Poole 1985). For review see Pardo et al. (2006).

ATP-Binding Cassette Transporters

The ABC transporter super family is the largest protein-family known, and its members

are capable of a numerous transport functions (Henikoff et al. 1997). The ABC

transporters are reported in several species ranging from archaebacteria to humans

(Higgins 1992; Volkl et al. 1996). They are capable of transporting alkaloids, lipids,

16

peptides, steroids, sugars, inorganic anions, and heavy metal chelates with the exception

of inorganic anions; MgATP directly energizes the transport of all of these substances

(Rea et al. 1998). All ABC transporters are constituted of one or two copies each of two

basic structural elements a hydrophobic, integral transmembrane domain containing four

to six transmembrane helices, and a cytoplasmically oriented ATP-binding domain. In

plants herbicide detoxification is performed by Multidrug Resistance associated Proteins

(MRP) that belongs to the ABC transporter family. From Arabidopsis thaliana plants

four MRP genes have been cloned: AtMRP1 (Lu et al. 1997), AtMRP2 (Lu et al. 1998),

AtMRP3 (Tommasini et al. 1998) and AtMRP4 (Sanchez-Fernandez et al. 1998). For

review on plant ATP-binding transports see Rea (2007).

Multidrug And Toxic compound Extrusion Transporters

The MATE transporter was first reported from bacteria (Morita et al. 1998) and its

homologs were later reported from plants and animals. Arabidopsis thaliana plant

contains 56 MATE transporter genes in its genome (Yazaki et al. 2008). MATE

transporters are involved in the transport of plant secondary metabolites. They consist of

400-700 amino acids with 9-12 transmembrane domains (Yazaki et al. 2008). In

Arabidopsis thaliana plants MATE Transporter TT12 acts as a vacuolar flavonoid/H+-

Antiporter (Marinova et al. 2007b). For review on MATE transporters see Yazaki et al.

(2008).

Aquaporins

Aquaporins are small (23-31 kDa) membrane proteins containing six transmembrane

domains that belong to the family of Major Intrinsic Proteins (MIPs). They are present in

microbes, animals and plants. They not only transport water but also small neutral solutes

and gases (Tyerman et al. 2002). On the basis of sequence homology plant aquaporin

family can be subdivided into four groups: the Plasma membrane Intrinsic Proteins (PIP),

Tonoplast Intrinsic Proteins (TIP), Nodulin-26–like Intrinsic membrane Proteins (NIPs)

and Small basic Intrinsic Proteins (SIPs). In Arabidopsis thaliana plant 10 homologs of

TIPs have been reported (Quigley et al. 2002). For classification and functions of plant

aquaporins see Maurel et al. (2008).

17

1.2.2 Vacuolar Channels

The electrochemical potential existing across the vacuolar membrane is very important

for any ion to be transported via channel mediated transport (Allen and Sanders 1997).

Differently from transporters, ion channels can conduct a large number of ions/sec with

high conductances ranging from 1 pS (pico siemens) to several thousand pS. Vacuolar

channels can be broadly divided into two major groups: anion channels and cation

channels. Though less studied anion channels have equal importance in the homeostasis

of the cell.

Anion Channels

The most abundant anions in plants are malate and nitrate. Malate plays a major role in

plant carbon metabolism and has many functions in plant cells, in charge balancing in the

vacuole, in the carboxylate and glyoxylate cycle, in CO2 temporary storage in C4 plants,

and in stomata and pulvini movements. In many of these processes, malate is

accumulated into the vacuole (Martinoia and Ratajcsak 1997). The malate channel is the

most studied anion channel in plant vacuoles, and it has been found in all plants studied

till now (Hafke et al. 2003; Hurth et al. 2005). These channels are selective for malate as

well as for other organic acids such as fumarate but typically impermeable to citrate. The

accumulation of malate mediated by channels is driven by the electrochemical gradient

across the tonoplast. This gradient is generated by malate concentrations, and by the

transtonoplastic potential created by the proton pumps. The molecular identity of malate

channels has been revealed by the identification of the AtALMT9 protein (Arabidopsis

thaliana Aluminium-activated Malate Transporter) (Kovermann et al. 2007).

The other major anion in plants is nitrate. Nitrate is an important source of nitrogen for

plants and most plants devote a significant portion of their energy for its uptake and

assimilation. Nitrate serves both as a nutrient and as a signal transducer and has profound

effects on plant metabolism and growth. The cytoplasmic nitrate has two major destinies:

(1) to be assimilated in organic compounds, and (2) to be accumulated into the vacuole.

The unassimilated nitrate is transported and stored in the vacuole up to concentration

values of 50-80 mM (Martinoia and Wiemken 1981). The accumulated nitrate is then

redistributed to cytoplasm under starvation.

18

It has been shown that in Arabidopsis thaliana plants the members of ChLoride Channel

(CLC) family, specifically AtCLCa is involved in nitrate homeostasis (De Angeli et al.

2006; De Angeli et al. 2009). Actually AtCLCa is not a channel (as initially

hypothesized) but a transporter that uses the proton gradient to actively accumulate

nitrate into the vacuole.

Cation channels

Potassium (K+) is the most important and abundant cation present in plant cells. Its

concentration may reach up to 10% of the plant dry weight. Potassium plays a role in

cellular processes like cell homeostasis, stomatal and leaf movements. Plants take up

potassium from the soil and plant cells accumulate K

+ to regulate the membrane potential

and turgor pressure. The cytoplasmic K

+ concentration is tightly controlled

at values in

the order of 100 mM. Vacuoles are the major sub cellular storage organelles for K+ in

plant cells (Very and Sentenac 2002). In Arabidopsis thaliana at least 35 genes encode

for K+ transport systems, out of these 15 encode for vacuolar and plasma membrane

channels (Maser et al. 2001). The potassium channels are structurally classified mainly

into three different families. They are shaker like potassium channels, K+ inward

rectifying like channels and two pore potassium channels. Although shaker like

potassium channels are not localized at the vacuolar membrane, the reasons for including

them in this part are the following (1) to give a general view on potassium channels; (2)

the slow vacuolar channel (see SV channel section below, which is the channel of interest

in this thesis) shares some structural similarities with shaker like channel.

Shaker like potassium channels

The first evidence of the existence of a shaker like plant potassium channel was reported

in 1992 (Sentenac et al. 1992). The name shaker like is given to this family of channels as

they resemble the shaker potassium channel cloned from Drosophila melanogaster

(Papazian et al. 1987). The shaker channels are tetramers i.e. four subunits tetramerize to

form functional ion channel with a central pore forming unit. They have six

transmembrane domains and one pore loop. The first four transmembrane domains form

the voltage sensor unit, and the last two domains

19

Fig. 1.4. Topology and functional types of potassium channels in plants: the

transmembrane domains and pore regions of three families (shaker like, TPK, Kir like)

are shown on the upper panel. Abbreviations ext: external, mb: membrane and cyt:

cytoplasmic side; P, CNBD, Anky, KHA, EF, respectively, pore domain, putative cyclic

nucleotide-binding domain, ankyrin domain (i.e. domain rich in hydrophobic and acidic

residues and Ef hand domain; +++: positively charged amino acids. The lower panel

shows the current-voltage (I-V) relationships illustrating the functional types forming

inward rectifying, outward- rectifying and weakly rectifying conductances of shaker type

channels. TPK channels, missing the first four segments, are not regulated by voltages

and show leak like conductance. Similar behavior is expected from Kir like channels.

However, none of them has been functionally identified, so far. From Lebaudy, Very et al.

(2007).

form the pore region. Shaker type channels show inward, outward and weakly inward

rectifying properties (see Fig. 1.4). There are excellent reviews available in the literature

on shaker channels (Gambale and Uozumi 2006; Lebaudy et al. 2007; Borjesson and

Elinder 2008).

Two-Pore K+ channels

Five different genes encoding TPK-type channels are present in Arabidopsis thaliana

plants. AtTPK4 is localized at the plasma membrane (Becker et al. 2004) while the other

four (AtTPK1, AtTPK2, AtTPK3 and AtTPK5) are localized at

the vacuolar membrane

(Voelker et al. 2006). TPK channels have four transmembrane domains and two pore

loops and do not contain the voltage sensor domain (see Fig. 1.4). Consequently,

membrane voltages do not regulate TPK channels and therefore they show a leak-like

conductance. Some members of this family have EF hand domains (helix-loop-helix

20

motif binding Ca2+

ions) at the cytoplasmic side. It has been demonstrated that 14-3-3

proteins enhance the calcium-dependent TPK1 channel activity (Latz et al. 2007).

K+ Inward Rectifying like channels

The name Kir-like was given to this family as they resemble animal K+ Inward

Rectifying channels. The first Kir-like channel in plants was identified in Arabidopsis by

sequence related genome search (Czempinski et al. 1999). The Kir-like channel has two

transmembrane domains and one pore loop. This channel has an EF hand domain at the C

terminal end and does not contain a voltage sensor domain.

Fast Vacuolar channel

FV channel was first functionally identified in beet root vacuoles (Hedrich and Neher

1987), later in broad bean guard cell vacuoles (Allen and Sanders 1996) and in Vicia faba

guard cell vacuoles (Allen et al. 1998). They are active at sub millimolar calcium

concentrations at the cytoplasm. FV channel mainly conducts K+ due its availability but it

is poorly selective between monovalent cations (Bruggemann et al. 1999). It has been

reported that FV channel can be modulated by cytosolic and vacuolar potassium (Pottosin

and Martinez-Estevez 2003). The molecular identity of FV channel is not yet revealed.

21

The Slow Vacuolar channel

The SV channel is one of the most extensively studied vacuolar channels. The SV

channel was shown to be Ca2+

activated in vacuoles from beet storage tissues (Hedrich

and Neher 1987) and has subsequently been identified in vacuoles from many plant cell

tissues and types (Pantoja et al. 1989; Maathuis and Prins 1990; Colombo et al. 1994;

Allen and Sanders 1997; Carpaneto et al. 1997; Paganetto et al. 2001; Scholz-Starke et al.

2004; Scholz-Starke et al. 2005a; Dziubinska et al. 2008). Nowadays it is generally

accepted that the SV channel is ubiquitous in higher plants.

Features of the SV channel are the slow time of activation, and the outward rectification

at elevated cytoplasmic Ca2+

concentration. For reviews see Scholz-Starke et al. (2005b)

and Pottosin and Schonknecht (2007). The SV channel is a non-selective cation channel

permeable to both monovalent and divalent cations (Pantoja et al. 1992; Ward and

Schroeder 1994; Allen and Sanders 1996; Gambale et al. 1996; Ivashikina and Hedrich

2005).

It displays a main conductance, which depends on potassium concentration. The whole

vacuolar currents of the SV channel investigated by patch-clamp method are in the order

of 10 to 100 pA pF-1

in red beet storage root and 100-500 pA pF-1

in guard cells at +100

mV potentials (Allen and Sanders 1997). More precisely, SV single-channel conductance

determined in sugar beet vacuoles are 40−300 pS at K+ concentrations ranging from 50 to

600 mM (Gambale et al. 1996) and 95 pS at 150 mM K+ (Scholz-Starke et al. 2005b). It

has been suggested that the SV channel is permeable to both K+ and Ca

2+ ions and that it

may act as a Calcium Induced Calcium Release (CICR) channel (Ward and Schroeder

1994). Interestingly cytosolic magnesium also plays a role in the modulation of this

channel (Pei et al. 1999; Carpaneto et al. 2001).

With densities of ~1 channel per µm2, or higher, the SV channel is the most abundant

vacuolar channel in plants (Schulz-Lessdorf and Hedrich 1995; Pottosin et al. 1997). On

the basis of conductance of single channel transitions the SV-channel pore-size is

estimated to be around 7 Aº (Amodeo et al. 1994; Paganetto et al. 2001).

In physiological conditions the tonoplast membrane potentials are slightly negative;

consequently the physiological role of the SV channel is not clear so far, as the channel is

mainly open at unphysiological positive tonoplast potentials. Therefore many studies are

aimed at verifying whether any cellular parameter is able to modify or shift the voltage

22

activation threshold of this channel towards more physiological negative potentials.

Several possible agents such as cytosolic Ca2+

(Hedrich and Neher 1987; Reifarth et al.

1994), cytosolic pH, oxidising and reducing agents (Schulz-Lessdorf and Hedrich 1995;

Carpaneto et al. 1999), cytosolic magnesium (Pei et al. 1999; Carpaneto et al. 2001),

vacuolar Ca2+

(Pottosin et al. 2004), potassium gradient across the tonoplast (Ivashikina

and Hedrich 2005), and cytosolic metals (Hedrich and Kurkdjian 1988; Corem et al.

2009) were shown to modulate the SV channel properties. It was reported that also

pharmacological compounds, like TEA (Weiser and Bentrup 1993; Dobrovinskaya et al.

1999) and neomycin (Scholz-Starke et al. 2006) can modulate the channel activity.

Structure of the SV channel

In 2005 it was demonstrated that the gene encoding for the SV channels in Arabidopsis

thaliana is AtTPC1 (At4g03560) (Peiter et al. 2005), whose molecular identity and

topology were already reported in 2001 (Furuichi et al.). AtTPC1 is a unique gene in

Arabidopsis thaliana plants where it is highly expressed in all tissues (Furuichi et al.

2001).

Fig. 1.5. Putative topology of AtTPC1 showing transmembrane segments (S1 to S12),

pore domains (P1 and P2) and EF hands.

AtTPC1 gene encodes a protein of 733 amino acids, which is targeted to the vacuolar

membrane. It has two conserved homologous domains both of which contain six

transmembrane segments (S1 to S6), a pore loop (P) between the S5 and S6 segments and

a EF hand domain at the cytosolic side (see Fig. 1.5). The protein reminds two Shaker

units which are fused together.

Vacuolar side

Cytosolic side

23

001 MEDPLIGRDS LGGGGTDRVR RSEAITHGTP FQKAAALVDL AEDGIGLPVE ILDQSSFGES

061 ARYYFIFTRL DLIWSLNYFA LLFLNFFEQP LWCEKNPKPS CKDRDYYYLG ELPYLTNAES

S1

121 IIYEVITLAI LLVHTFFPIS YEGSRIFWTS RLNLVKVACV VILFVDVLVD FLYLSPLAFD

S2 S3

181 FLPFRIAPYV RVIIFILSIR ELRDTLVLLS GMLGTYLNIL ALWMLFLLFA SWIAFVMFED

S4 S5

241 TQQGLTVFTS YGATLYQMFI LFTTSNNPDV WIPAYKSSRW SSVFFVLYVL IGVYFVTNLI

P1 S6

301 LAVVYDSFKE QLAKQVSGMD QMKRRMLEKA FGLIDSDKNG EIDKNQCIKL FEQLTNYRTL

EF

361 PKISKEEFGL IFDELDDTRD FKINKDEFAD LCQAIALRFQ KEEVPSLFEH FPQIYHSALS

EF

421 QQLRAFVRSP NFGYAISFIL IINFIAVVVE TTLDIEESSA QKPWQVAEFV FGWIYVLEMA

S1 S2

481 LKIYTYGFEN YWREGANRFD FLVTWVIVIG ETATFITPDE NTFFSNGEWI RYLLLARMLR

S3

541 LIRLLMNVQR YRAFIATFIT LIPSLMPYLG TIFCVLCIYC SIGVQVFGGL VNAGNKKLFE

S4 S5

601 TELAEDDYLL FNFNDYPNGM VTLFNLLVMG NWQVWMESYK DLTGTWWSIT YFVSFYVITI

P2

661 LLLLNLVVAF VLEAFFTELD LEEEEKCQGQ DSQEKRNRRR SAGSKSRSQR VDTLLHHMLG

S6

721 DELSKPECST SDT

Fig. 1.6. Protein sequence of Arabidopsis thaliana TPC1 (AtTPC1) presenting putative

transmembrane domains S1 to S6 (highlighted in yellow), putative pore domains P1 and

P2 (in light green), and EF hands (in blue).

Arabidopsis thaliana TPC1 (AtTPC1) protein sequence is shown in Fig.1.6.

24

Some possible hypothesis on the physiological relevance of the SV channel

Even though the SV is the most extensively studied channel in plant vacuoles, its

physiological role is not fully understood. It’s well known that the SV currents recorded

in a typical patch-clamp experiment require unphysiologically high cytosolic and low

vacuolar calcium concentrations for full activation. It was also reported that the channel

is involved in calcium signalling thus suggesting that the SV channel is directly involved

in Calcium Induced Calcium Release (CICR) in Arabidopsis thaliana where it might be

involved in germination and stomatal movements (Peiter et al. 2005). However it has also

been observed that SV CICR is incompatible with the drastic decrease of the open

probability of the channel in conditions that favour the calcium release from the vacuole

to the cytosol (Pottosin et al. 1997). Interestingly, the knock out plants of tpc1-2 do not

show morphological differences compared to wild type plants. It seems that these plants

do not suffer the loss of AtTPC1 gene.

Interestingly a point mutation in the vacuolar side of the channel determines different

properties of the SV channel and even a diverse plant morphology, compared to wild type

plants (Bonaventure et al. 2007). Indeed the Arabidopsis mutant fatty acid oxygenation

up regulated 2 (fou2) is shown to up regulate oxylipin in the Jasmonate pathway. For a

brief introduction to this pathway see section Ch. 1.4. In the fou2 mutant plant

metabolites of 13-LOXs, and specifically oxylipin biogenesis strongly increased in

response to wounding. In the fou2 mutant plants not only SV channel is active at more

negative voltages but also the plants are more resistant to fungal pathogen Botrytis

cinerea (Bonaventure et al. 2007). Moreover inhibition by vacuolar calcium is less

pronounced in comparison with wild type plants as reported by Beyhl et al. (2009).

Recently it has been reported that Nicotinic Acid Adenine Dinucleotide Phosphate

(NAADP) mobilizes calcium from acidic organelles through a two-pore channels from

mammalian systems (Calcraft et al. 2009; Zong et al. 2009). NAADP is now an emerging

molecule to study the calcium release from acidic stores through two-pore channels (for

reviews see (Galione and Churchill 2002; Brailoiu et al. 2009; Guse 2009). The

modulation of plant two-pore channel by this molecule still deserve further investigation.

25

Chapter 1.3. Flavonoid biosynthesis, localization and transport

Flavonoids are pigments or secondary metabolites present in all plants. They are one of

the largest group of secondary metabolites, with more than 8000 different compounds,

reported till now (Harborne and Williams 2000; Ververidis et al. 2007). The common

structure of all the flavonoids is a C15 phenylchromane core composed of C6-C3-C6

backbone. The chromane (benzopyran) motif is made up of 2 condensed rings, an

aromatic (benzo) A-ring (C6) and a heterocyclic (pyran) C-ring (C3) associated with

another aromatic B-ring (C6). In the majority of flavonoids (flavanols, anthocyanidins,

flavanones, flavones, and flavonols) the B-ring is attached at position 2, whereas in

isoflavones the B-ring is instead attached at the position 3.

Fig. 1.7. Basic structure of flavonoids. C15 phenylchromane core composed of C6-C3-

C6 backbone. The chromane motif is made up of 2 condensed rings, an aromatic A-ring

(C6) and a heterocyclic C-ring (C3) associated with another aromatic B-ring (C6). In the

majority of flavonoids the B-ring is attached at position 2 (A), whereas in isoflavonoids

the B-ring is attached at position 3 (B).

Each flavonoid subclass comprises numerous members, differing in the degree of

hydroxylation or methoxylation of A and B rings. Modifications at single or multiple

positions of flavonoid molecules via hydroxylation, methylation, acylation, or

glycosylation reactions provide unique features and characteristics to most of the

currently known flavonoids (Fig. 1.8). For review on flavonoid biosynthetic pathway see

Winkel-Shirley (2001), Lepiniec et al. (2006) and Winkel-Shirley (2006).

A B

26

Fig. 1.8. Flavonoid biosynthetic pathway. ACCase, acetyl CoA carboxylase; ANS,

anthocyanidin synthase; AS, aureusidin synthase; DFR, dihydroflavonol 4-reductase;

DMID, 7,2′-dihydroxy, 4′-methoxyisoflavanol dehydratase; F3H, flavanone 3-

hydroxylase; F3′H, flavonoid 3′-hydroxylase; F3′5′H, flavonoid 3′5′ hydroxylase; FLS,

flavonol synthase; FSI/FS2, flavone synthase; I2′H, isoflavone 2′-hydroxylase; IFR,

isoflavone reductase; IFS, isoflavone synthase; IOMT, isoflavone O-methyltransferase;

LAR, leucoanthocyanidin reductase; LDOX, leucoanthocyanidin dioxygenase; OMT, O-

methyltransferase; RT, rhamnosyl transferase; UFGT, UDP flavonoid glucosyl

transferase; VR, vestitone reductase. From Lepiniec et al. (2006).

27

Flavonoids are secondary metabolites unique to plants, where they play very important

roles in many processes such as defense against insects (Simmonds 2003; Lev-Yadun et

al. 2004; Treutter 2006), UV protection (Li et al. 1993; Kootstra 1994), antioxidant

activity (Rice-Evans et al. 1997; Hernández et al. 2009), temperature stress (Coberly and

Rausher 2003; Kirakosyan et al. 2003), heavy metal tolerance (Cocker et al. 1998; Kidd

et al. 2001), chemical signals for pollination (Grotewold 2006), pollen growth (Taylor

and Grotewold 2005; Yang et al. 2008), seed dormancy and seed protection (Debeaujon

et al. 2000), microbial interactions (Cohen et al. 2001; Shaw et al. 2006) and regulation

of auxins for root development (Imin et al. 2007; Santelia et al. 2008).

It is generally believed that flavonoids are synthesized through the phenylpropanoid

pathway, by cytosolic multienzyme complexes at the cytoplasmic surface of the

Endoplasmic Reticulum (ER) (Stafford 1974; Winkel-Shirley 1999). Some enzymes,

participating in flavonoid metabolism, were also reported to co-localize at the ER,

tonoplast, nucleus, secondary cell wall and plastid (Saslowsky and Winkel-Shirley 2001;

Tian et al. 2008). The flavanone Naringenin plays a central role in the biosynthetic

pathway (see Fig. 1.8) and is present in all the plants; in plants such as Citrus is one of

the major flavonoids (Rouseff et al. 1987). The glycosylase transferase gene, which

modifies Naringenin into Naringin, was also reported to be present in Citrus fruits

(Frydman et al. 2004). It has been reported that Naringenin is involved in the fungal

resistance (Padmavati et al. 1997), in lateral root growth (Webster et al. 1998) and in

lignin biosynthesis (Deng et al. 2004) in plants. Naringin is evolved in microbial

interactions (Cohen and Yamasaki 2000). Flavonoid deficient mutants are useful tools to

study the importance of flavonoids in plants. They are grouped into transparent testa (tt)

phenotype mutants, where the seed coat is affected at various steps (see Fig. 1.9) by a

deficiency in the flavonoid biosynthetic pathway (Shirley et al. 1995; Buer and Muday

2004; Buer and Djordjevic 2009).

28

Fig. 1.9. Schematic representation of flavonoid biosynthetic pathway. Enzymatic steps

affected in specific transparent testa mutants are indicated with putative regulatory loci

indicated in parentheses. From Shirley et al. (1995).

29

Flavonoids are transported to different locations from the sites of their synthesis. Most of

the flavonoids are accumulated in the vacuole; others may be secreted and incorporated

into the cell wall. Even though the major steps in flavonoid biosynthetic pathway are

known, there are still unanswered questions concerning the mechanism(s) by which the

flavonoid end-products are properly sorted and transported into the diverse intracellular

compartments.

Fig. 1.10. Hypothetical scheme of flavonoid transport pathways in grapevine. Flavonoids

could be conjugated with glutathione (GSH) through a reaction catalysed by glutathione

S transferases (GSTs). The main transporters localized in grapevine vacuole and plasma

membrane are the ATP-binding cassette (ABC) proteins and the bilitranslocase-

homologue (BTL-homologue). The multidrug and toxic compound extrusion (MATE)

protein, shown to be involved in flavonoid transport in other plant species, has also been

added. Transport mediated by vesicle trafficking is indicated by circles (AVIs,

anthocyanic vacuolar inclusions; ACPs, anthocyanoplasts). Question marks indicate the

lack of information or still hypothetical components and steps in the process. From

(Braidot et al. 2008).

It has been suggested that the flavonoids are transported to the vacuolar membrane

through different pathways (Kitamura 2006; Braidot et al. 2008; Passamonti et al. 2009)

(see Fig. 1.10). In one of the mechanisms it is assumed that flavonoid-specific

Glutathione S Transferases (GSTs) carry flavonoids to the vacuole via non-covalent

30

activity (Grotewold 2001). Alternatively, the second mechanism requires a complex

vesicle-trafficking network, possibly involving the Golgi apparatus (Grotewold 2004;

Grotewold 2006; Poustka et al. 2007). Specific vacuolar transporters were also reported

from several species such as maize (Zea mays), petunia (Petunia hybrida), Arabidopsis

thaliana and grapevine (Vitis vinifera) (Kitamura 2006). These transporters can be

broadly divided into two types: those, which are activated by the hydrolysis of ATP, and

those dependent on a secondary activation, modulated by the trans-membrane proton

gradient across the vacuolar membrane. The ATP-binding cassette (ABC) proteins were

shown to be involved in the transport of flavonoids into the vacuolar lumen (Yazaki

2005; Klein et al. 2006; Yazaki 2006; Rea 2007). In contrast proton/flavonoid antiporters

use trans-membrane proton gradients to transport flavonoids into the vacuole. Multidrug

And Toxic Efflux (MATE) proteins mediate this mechanism as reported in Arabidopsis

thaliana (Debeaujon et al. 2001; Marinova et al. 2007b) and in barley (Hordeum vulgare)

(Marinova et al. 2007a). It has been suggested that Glutathione-linked flavonoids would

be primarily recognized by ABC proteins, in particular Multidrug Resistance associated

Proteins (MRP) (Yazaki 2005) and glycosylated flavonoids would be preferentially

transported by secondary active MATE-like antiporters.

Ch.1.4. Polyunsaturated fatty acids

Membrane lipids are classified into phospholipids, cholesterol and glycolipids and their

ratio may differ from membrane to membrane. All of them possess polar regions tpically

facing towards the aqueous compartments and non-polar regions which constitute the

interior of the lipid bilayer. Membrane phospholipids are built upon the three carbon

backbone of glycerol and have a polar phosphate group or phosphate plus a polar,

hydrophilic group in the sn-3 position and two fatty acid tails. The fatty acid tail

esterified to the first carbon of the glycerol backbone (sn-1 position) is generally a

saturated fatty acid, lacking carbon–carbon double bonds. The tail esterified to the second

carbon (sn-2 position) is generally an unsaturated fatty acid with one (monounsaturated)

or more (polyunsaturated) carbon–carbon double bonds.

31

The mammalian membrane phospholipids mainly contain long chain ω-3

(eicosapentaenoic acid, EPA, C20:5) and ω-6 (Arachidonic Acid, AA, C22:6) fatty acids.

As they serve as signalling molecules many studies are aimed at verifying modulation of

mammalian ion channels by polyunsaturated fatty acids (PUFAs). We know for almost

20 years that lipids and lipid bioproducts are able to fine-tune voltage-gated ion channels

in animal cells. For review on this topic see Boland and Drzewiecki (2008). Little or no

Arachidonic Acid is present in plants however plants use other polyunsaturated fatty

acids such as linoleic acid and Docosahexaenoic acid. For example the plant

phospholipids of tonoplast mainly contain ω-6 (linoleic acid, C18:2). In fact a study

demonstrated that more than 50% of the total fatty acid composition in the vacuolar

membrane of the plants parsnip, parsley and carrot is made of linoleic acid (Makarenko et

al. 2007). The interaction of PUFAs with plant ion channels is less studied so far, but the

downstream products of PUFAs, such as jasmonate members of oxylipins, (which lies in

the wound response pathway) are in focus for the last few decades. The biosynthesis of

most plant oxylipins are initiated by the action of lipooxygenases (LOX), which are

capable of introducing molecular oxygen at either C19 or C13 position of the C18 fatty

acids linoleic acid and α-linolenic acid (Feussner and Wasternack 2002). A mutation in

the vacuolar side of the AtTPC1 gene (fou2 mutant) was shown to up regulate oxylipin

and this mutant plants were more resistant to the fungal pathogen Botrytis cinerea

(Bonaventure et al. 2007). For reviews on jasmonate pathway see Wasternack (2007) and

Bottcher and Pollmann (2009).

Ch 1.5. Objectives

The Slow Vacuolar channel is one of the most extensively studied ion channels in plants.

The SV channel was shown to be calcium activated in vacuoles from beet storage tissues

(Hedrich and Neher 1987) and was subsequently identified in vacuoles from many tissues

and plants (Allen and Sanders 1997; Carpaneto et al. 1997; Scholz-Starke et al. 2005a;

Dziubinska et al. 2008), thus demonstrating that it is ubiquitous in higher plants.

Features of the SV channel are the slow activation and the outward rectification at

elevated unphysiological cytoplasmic calcium concentrations. Indeed it is well known

32

that the SV currents recorded in a typical patch-clamp experiment require

unphysiologically high cytosolic and low vacuolar calcium concentrations for activation.

Furthermore, it has been demonstrated that the SV channel, besides being permeable to

potassium ions, is also permeable to calcium (Gradogna et al. 2009).

It is well known that the Slow Vacuolar channel is modulated by several parameters and

compounds, internal and external to the vacuole; beside calcium, Mg2+

, protons, reducing

agents, metals (like Zn2+

and Ni2+

) and the glycosidic antibiotic Neomycin are able to

modify the properties of the channel. However, none of these parameters is able to

activate the SV channel at physiological membrane potentials and calcium

concentrations. For this reason we aim at investigating unexplored working conditions or

compounds, which are able to further modulate this channel. Specifically we plan to find

out endogenous plant molecules, such as flavonoids and PUFAs, which might be able to

modify the voltage activation threshold of this channel towards more physiological

conditions or that may provide information on the role and the nature of this enigmatic

channel.

33

Materials and methods

34

35

Chapter 2.1. Methods for localization of flavonoids in plant leaves

Plant material

Arabidopsis thaliana seeds from Columbia (Col-0) and Landsberg erecta (Ler) ecotypes

were sowed in soil pots. After 5 days of vernalization at 4°C they were transferred to the

growth chamber for germination (light period was 12h dark and 12h light), 8-10 days

after they were individually transferred in pots and grown under controlled conditions in

the green house. The light period was 16h dark and 8h light, with a light intensity of 179

µmol m-2

s-1

and a temperature of 22°C. Plants were weekly watered with nutrient

solution: 18% Nitrogen (12% Nitrate, 6% Ammonia), 6% Phosphorus Anhydride, 26%

Potassium Oxide, 2% Magnesium Oxide and oligoelements <1%.

Nicotiana tabacum plants were grown on soil in 9 cm pots and kept under controlled

conditions in growth chamber. The light period was 16h dark and 8h light, with a light

intensity of 70 µmol m-2

s-1

and a temperature of 22°C.

Construction of recombinant plasmids

The AtGT (At1g06000) was amplified by PCR using the Phusion® DNA Polymerase

(Finnzymes, Finland) and Arabidopsis thaliana genomic DNA was used as template,

since no intrones are predicted in the gene sequence. The primers used for the PCR were

the following: Nar-NcoI-For (5'-CATGccatggccATGACAACAACAACAACGAAGA-

3', right before ATG start codon with NcoI site) and Nar-NcoI-Rev (5'-CATGCcatggcTgc

CAAACACATCTCTGCAACGAGCT -3', after the stop codon with NcoI site). The stop

codon TAA was mutated into GCA and, to maintain the frame, two extra CC where

introduced between the AtGT and the Yellow Fluorescent Protein (YFP) coding

sequences; in this way two alanines were introduced at the junction of the fusion protein.

The PCR product was digested with NcoI enzyme and cloned in the vector pGreen 0029

in which previously multiple cloning sites was replaced with 2x35S promoter, YFP and

nos terminator using KpnI and SacI sites (Zottini et al. 2008). The right orientation was

confirmed by digestion and sequencing.

36

Western Blot

For protein extraction, tobacco leaf pieces, corresponding to agroinfiltrated areas, were

harvested, frozen, powdered in liquid N2 and homogenized in two volumes of protein

extraction buffer [0.3 M sucrose, 0.1 M Tris pH 7.5, 1 mM ethylenediaminetetraacetic

acid (EDTA), 5 mM dithiothreitol (DTT) containing 2 mM phenylmethylsulfonyl

fluoride (PMSF), 1 mM benzamidine, 5 mM ε-aminocaproic acid, 5 µg ml−1 leupeptin, 2

µg ml−1 aprotinin and 0.7 µg ml−1 pepstatin]. The homogenate was centrifuged at 1500

g for 15 min at 4°C to eliminate debris. Protein concentration was determined by the

Bradford method, using the Bio-Rad protein assay (Bio-Rad, Segrate, Italy). Total

extracted proteins (3 and 6 µg) were separated by 12% (w/v) SDS–PAGE, transferred to

a nitrocellulose membrane (Sartorius) and hybridized with a polyclonal antibody raised

against YFP (Invitrogen Molecular Probes). The blot was then hybridized with an

alkaline phosphate conjugated secondary antibody and the detection was performed with

the NBT/BCIP colorimetric assay.

Transient expression

A. Mesophyll protoplasts from Arabidopsis thaliana

The transient transformation was performed following the Sheen protocol (2002). Briefly

leaves from soil grown 4/5 weeks old Arabidopsis thaliana plants were cut into narrow

strips and put in freshly prepared enzymatic solution (1.25% cellulase R10, 0.03%

macerozyme R10, 0.4 M mannitol, 20 mM KCl, 20 mM MES, 10 mM CaCl2 and 0.1%

BSA, pH 5.7) under vacuum for 30 minutes. The digestion was then carried out for 3

hours at 25°C in the dark. The protoplasts were then filtered with 50 m nylon mesh and

centrifugated at 90 g. The pellet was washed twice with W5 solution (154 mM NaCl, 125

mM CaCl2, 5 mM KCl and 2 mM MES, pH 5.7) and the protoplasts were kept in ice for

30 min before the PEG transformation. PEG-mediated protoplast transformation was

achieved on protoplasts aliquots of 105 protoplasts/ml, with 4 μg of DNA. Transformed

protoplasts were incubated overnight in W5 solution in the dark before confocal analysis.

B. Tobacco leaves

Competent cells of Agrobacterium tumefaciens GV3101 strain were prepared as

previously described (Zottini et al., 2008) and the binary vectors were introduced by

37

freeze/thow method. For the use of pGreen-derived vectors we cotransformed the

Agrobacterium tumefaciens cells with the pSoup vector (Hellens et al. 2000a). The

Agrobacterium is grown in YEP medium (Yeast Extract, 10 g/l; Bacto-Peptone 10 g/l;

NaCl 5 g/l; pH 7.2) supplemented with specific antibiotics (rifampicin 25 mg/l,

gentamycin 50 mg/l, kanamycin 50 mg/l and tetracyclin 5 mg/l) on a rotary shaker at

28°C.

Agroinfiltration of tobacco leaves

The tobacco agroinfiltration protocol was performed as described in Batoko et al. (2000).

Two days prior to agroinfiltration, single colonies of Agrobacterium tumefaciens growing

on agar plates were inoculated in 5 ml YEP liquid medium supplemented with specific

antibiotics. The bacterial cultures were incubated at 28–30oC at 200 rpm on an orbital

shaker. New bacteria cultures were started two days later by inoculating fresh medium

with the old suspension cultures (1/200 ratio, v/v). The new cultures were grown under

the same conditions for an additional day. After 24h, 2 ml of each bacteria culture were

transferred to eppendorf tubes and pelletted by centrifugation at 1,500 g for 4 min. in a

micro-centrifuge at room temperature. The pellets were washed twice with 2 ml of

infiltration buffer [50 mM MES pH 5.6, 2 mM Na3PO4, 0.5% glucose (w/v) and 100 M

acetosyringone (Aldrich, Italy)]. The bacteria suspensions were diluted with infiltration

buffer to a final OD600 of 0.2 and allowed to grow for 1-2h at 25oC in the dark, with

continuous gentle shaking before using them for agroinfiltration experiments. The

inocula were delivered to the lamina tissues of tobacco leaves by gentle pressure

infiltration through the stomata of the lower epidermis by using a 1-ml syringe without a

needle. Young leaves prior to full expansion were used. After infiltration the plants were

transferred in a growth chamber under standard growth conditions.

Fluorescence image acquisition

YFP-dependent fluorescence was analyzed after 24 hours in case of Arabidopsis

mesophyll protoplasts and 6 days after agroinfiltration of tobacco leaves. Arabidopsis

protoplasts were mounted on the slides with cover slips for microscopic observations.

Pieces of tobacco leafs were randomly cut from the infected areas and mounted on slides

for microscopic observations. Confocal microscope analyses were performed using a

38

Nikon PCM2000 (Bio-Rad, Germany) laser scanning confocal imaging system. For YFP

detection excitation was at 488 nm and emission between 530/560 nm. For the

chlorophyll detection, excitation was at 488 nm and detection over 570 nm. Image

analysis was done with the ImageJ bundle software (http://rsb.info.nih.gov/ij/).

Ch. 2.2. Electrophysiology: the patch-clamp methodology

Prior to 1976 typical electrophysiological experiments were performed on large cells, like

the squid giant axon, which have a size in the order of about 1 mm x 2 cm (Hille 1978).

Erwin Neher and Bert Sackman introduced patch-clamp technique in 1976 (Neher and

Sakmann 1976) in order to allow the measurement of ionic transport under controlled

potential and ionic conditions in small (~10-100 μm) cells.

A glass pipette with a very small opening (<1 μm) is used to make tight contact with a

tiny area, or patch, of the vacuolar membrane. After the application of a small negative

pressure, the seal between the pipette and the membrane becomes so tight that very few

ions can flow between the pipette and the membrane. In good conditions the electric

resistance between the pipette and the membrane reaches values larger than one gigaohm,

109 Ω (“gigaseal” see Fig. 2.1). This arrangement is usually called the vacuole-attached

patch-clamp recording configuration (Fig. 2.2 A). Consequently, the recorded current

(flowing through the recording pipette) is exclusively the current passing through the

membrane. Thus, all the ions that flow when a single ion channel opens must flow into

the pipette. The resulting electrical current, though small, can be measured with an ultra

sensitive electronic amplifier connected to the pipette.

Once a tight seal is attained, several experimental options are available, according to the

type of currents that need to be studied (Fig. 2.2). If the membrane patch below the

pipette is disrupted by applying a relatively strong suction pulse or a large voltage pulse,

the interior of the pipette becomes continuous with the interior of the vacuole. This

arrangement allows measurements of electrical potentials and currents from the entire

vacuole and is therefore called the whole-vacuole recording configuration (Fig. 2.2 B).

The whole-vacuole configuration also allows diffusion exchange between the pipette and

the interior of the vacuole, producing a convenient way to perfuse substances into the

interior of a “patched” vacuole.

39

Fig. 2.1. Schematic representation of a “gigaseal” during a patch-clamp experiment. A)

When the pipette is still outside of the solution, the current consists of a fast capacitive

transient. B) The pipette is dipped into the bath while applying slightly positive pressure,

in order to keep the tip of the pipette clean while passing through the air/water interface.

C) The pipette is still far from the vacuole, and the current, observed as a response to a

small potential pulse, allows to evaluate the pipette resistance. D) The pipette is carefully

approaching the vacuole, while still keeping a slightly positive pressure inside. A decline

in the current indicates that the pipette is touching the vacuole. E) The pressure is

released and a further decline of the current is observed. F) A weak negative pressure is

applied until the seal is formed. The formation of a good seal is pointed out by an almost

complete decline of the current. The reduction of the leak current reduces also the

registration’s background-noise, allowing single channel registration. A contact

achieved in this way presents high mechanical stability, that allows to perform

registrations in four different configurations.

40

Fig. 2.2. Schematic representation of four configurations that can be achieved during a

patch-clamp experiment. A) “Vacuole-attached” configuration; B) “Whole-vacuole”

configuration, depart from a vacuole-attached configuration; C) “Inside-out” patch

configuration, depart from a vacuole-attached configuration; D) “Outside-out” patch

configuration, depart from a whole-vacuole configuration.

If the pipette is retracted while it is in the whole-vacuole configuration, the membrane

patch obtained has its cytosolic surface exposed to the bath. This arrangement, called the

outside-out recording configuration (Fig. 2.2 D), is optimal for studying how the single

channel activity of one or a few channels is influenced by cytosolic chemical signals.

Once a tight seal has formed between the membrane and the glass pipette, small pieces of

membrane can be pulled away from the cell without disrupting the seal; this yields a

preparation that is free of the complications imposed by the rest of the vacuole. By

retracting a pipette that is in the vacuole-attached configuration one generates a small

patch of membrane with its vacuolar surface exposed to the bath solution. This

arrangement, called the inside-out patch recording configuration (Fig. 2.2 C), allows the

measurement of single-channel currents with the added benefit of making it possible to

change the medium to which the surface internal to the vacuole is exposed.

A B

C D

41

The patch-clamp setup

A patch-clamp setup (Fig. 2.3) consists of a microscope placed on an antivibrational table

within a Faraday cage, that reduces to a minimum the electric disturbances from the

surrounding electronic instruments. The system contains also a patch-clamp amplifier for

voltage-clamping the cell, a micromanipulator holding the amplifier probe and the

attached patch-pipette and a personal computer to generate the stimulation and record the

current. An inverted microscope (Zeiss Axiovert 10) with a magnification of up to 320X

(32X10), allows the visualization of the cell and to monitor the touch-down of the pipette

on the cell.

In order to obtain good seals, it is essential to precisely control the movement of the

pipette in the sub micrometer range. To this purpose two micromanipulators are used: 1)

a mechanically driven system (Microcontrole, France), allowing movements in the order

of mm, to be used when the pipette is still far from the vacuole and 2) a hydraulically

driven system (Narishige, Japan), allowing fine movements in the order of fractions of

micrometer. The stimulation and acquisition system is based on a personal computer

(Macintosh Quadra 700), which uses a specially designed software (Pulse+PulseFit, Heka

Electronics, Germany) to send to the amplifier the stimulation protocols and register the

evoked currents. An A/D/A converter (Instrutech ETC 16) provides the in and out data

conversion.

The membrane potential and the survey of the current signal are controlled by a patch-

clamp amplifier (List EPC-7), which has two parts: the headstage and the central body.

The headstage is equipped with an electrode-holder; this is the physical support for the

glass pipette, and has a double role: firstly, to establish the electrical contact between the

pipette solution and the current/voltage converter of the headstage through an Ag||Ag/Cl

electrode and secondly, to allow the regulation of the pressure applied to the cell through

the glass pipette.

42

Fig. 2.3. Patch-clamp setup at the IBF-CNR laboratories in Genoa. At the right hand

side the computer monitor can be seen. In the center, the electronic equipment, from

bottom: A/D/A converter, filter, central body of the amplifier. At the left hand side:

Faraday cage, where the microscope, the amplifier-headstage and the

micromanipulators are positioned. The microscope is equipped with a video camera

transferring images to the monitor on the top of the Faraday cage.

In the headstage the current signal is converted to a voltage signal and transferred to the

amplifier. The amplifier allows also to change the gain, and to compensate the effects of

the pipette capacity, the membrane capacity and the access resistance. Currents from

several tens of nA to fractions of pA can be recorded by the amplifier. The voltage

signals are filtered by a four-poles Bessel filter (KEMO) and transferred to the computer

for digital display and data storage on the hard disk.

43

Perfusion systems

During the patch-clamp experiments the ionic solution of the bath needs to be changed

from the standard (control) solution to different ionic solutions.

In this work, two types of perfusion systems were used: a “gravity-driven” and a “fast”

perfusion system (Carpaneto et al. 1999; Carpaneto et al. 2001), that differ mainly in the

speed, of changing the external solution.

Fig. 2.4. The registration-dish placed on the microscope, with arrows indicating (1) the

electrode-holder with the glass micro-pipette, (2) the small tubes of the perfusion system,

(3) the suction-tube of the peristaltic pump and (4) the ground-electrode.

A gravity-driven perfusion system allows to change the external solution in a few

minutes by means of 2-6 syringes of 20 ml (depending on the experimental demand). The

syringes deliver the solution by means of a valve and polyetilene tubes (Fig. 2.4). The

bath solution is constantly removed from the registration petri dish by means of a

peristaltic pump.

A fast-perfusion system is used to change the external solution in time intervals in the

order of few ms. The system comprises a part that regulates the velocity of the flux,

which includes a support system with an independent height regulation (with respect to

the registration petri-dish) for 3 capillaries of 200 μl. The capillaries are treated with

44

Fig. 2.5. Schematic representation of the “fast perfusion system” (above); a picture

through a microscope showing 3 pipettes containing different solutions, positioned close

to the patched vacuole (below).

the silanizing agent sigmacote that decreases the surface tension of the liquid bathing the

interior of the pipette. The capillaries are connected to a valve on one side, and to a

flexible tygon tube on the other side. The second part is formed of 3-glass micropipette

with a tip of 20 to 30 μm diameter. On one side they are connected to the rest of the

system with a tygon tube, while on the other side they are soaked in the bath solution

(Fig. 2.5). The 3 micropipettes are fixed to a metal support that allows an independent

regulation of their position in three directions, while they are lined in the same focal

plane and the distance between them is about 10 μm. The perfusion pipette can be

positioned in front of the cell by means of hydraulic micromanipulator (Narishige,

Japan).

Instead of plastic tubings, glass tubes connected to oil-filled syringes controlled by a

stepping motor was used for the perfusion of polyunsaturated fatty acids (PUFAs) to

avoid the binding of lipids to the perfusion system. This system allowed a slow release of

the PUFA solution with a flow rate of about 10 μl/min.

Micropipettes and electrodes

The patch pipettes were pulled by a pipette puller (Micropipette puller mod L/M 3P-A

List Medical) from borosilicate glass capillaries (Clark Electromedical Instruments)

which have an external diameter of 1.7 mm and an internal of 1.2 mm. The capillaries

were cleaned by flame and coated internally with sigmacote.

45

Data acquisition

Patch-clamp experiments were typically made in excised-patch configuration. Patches in

the excised configuration are much more stable than those in the whole-vacuolar

configuration and allow to maintain the patch for a long time under fast or slow perfusion

conditions. Pipette resistance was 1.8-2.4 mOhm. Seals with more than 2 GOhm

resistances were obtained prior to the establishment of the excised-patch configuration.

Patch-clamp recordings were made by an EPC7 (List) amplifier or in alternative by an

Axon 200A amplifier (Axon Instruments). Data were acquired with software Pulse and

Pulsefit (HEKA) and analysed with Igor Pro (Wavemetrics). Currents are typically

evoked in response to series of +10 mV voltage steps ranging from -80 to +120 mV; tail

voltages are the voltages following the main step pulse (Fig. 2.6). The signs of current

and voltage follow the convention proposed by Bertl et al. (1992).

Fig. 2.6. Typical stimulation protocol used to record the ionic current at different step

potentials. The “tail” currents are elicited by “tail” voltages applied after the main

pulse. In our working conditions the tail voltage is typically equal to the holding potential

(-50 mV; see Fig. 3.4 for SV currents elicited by this protocol).

Data analysis

The I-V characteristics of the SV channel is constructed by plotting the average value of

the current recorded during the last 50 ms at each applied voltages.

46

The dose-response analysis of Naringenin block was performed by calculating the

residual SV currents (INar /Icontrol) at +80 mV. (INar /Icontrol) was plotted as a function of

Naringenin concentration and data were fitted with the Hill equation

INar /Icontrol = 1/(1+([Nar]/Kh)n) eq. 2.1

where [Nar] is the Naringenin concentration, Kh and n are the Hill inhibition constant and

Hill coefficient, respectively.

In the dose-response analysis where Gnorm was plotted as a function of Nar or AA

concentrations Gnorm was obtained from the Boltzmann analysis (see below).

Boltzmann analysis:

1. We define It as the current at the beginning of the tail protocol (see protocol in Fig.2.6

and Fig 3.4). The equation for It is the following:

It (Vp)=N g (Vt) (Vt-Vrev) P(Vp) eq. 2.2

where Vp=voltage of main pulse, N=number of channels, g=single channel

conductance,Vt=tail potential, Vrev=reversal potential, P(Vp)=open probability at the

voltage of main pulse.

2. Evaluation of It: the tail current was fitted with a single exponential function. The fit

was performed 1-3 ms after the onset of the tail voltage in order to remove capacitance

artefacts.

3. We fitted It (Vp) with the following Boltzmann function:

It=Ioff +Imax /(1+exp (-(Vp-V1/2)/s)) eq. 2.3

where Ioff, Imax,V1/2 and s are the free parameters of the fit. Ioff represents the current

offset, Imax the maximum of the current, Vp the applied voltage, V1/2 half activation

voltage, and the s is RT/ Fz where z is the apparent gating charge.

4. Normalized conductance was calculated by

47

Gnorm= (It-Ioff)/Imax(control) eq. 2.4

where Imax (control) is the maximum value of the Boltzmann fit in control conditions.

5. Averages of Gnorm for at least three different vacuoles in control and at varying

concentrations of Naringenin and Arachidonic Acid were calculated.

Half activation time is the time at which the current assumes the half of its maximum

value. The maximum value is evaluated from the difference between steady state current,

and the initial current, where the channels are still closed.

Detection of fura2 fluorescence and data analysis

The recording chamber was mounted on the stage of an inverted microscope (IM35,

Zeiss, http://www.zeiss.com/) with a 100× Nikon Fluor objective (Nikon,

http://www.nikon.com). For fura-2 excitation, the tip of the recording pipette was

illuminated using a xenon lamp via a light fiber. The light beam passed through a rotating

wheel with six interference filters, four centered at 340 nm and two centered at 380 nm.

A computer-driven spectrophotometer system (Cairn Research, http://www.cairn-

research.co.uk) controlled both rotor speed and acquisition of emission light through a

photomultiplier (Fig. 2.7). The mean emission relative to each set of filters was

calculated. The speed was set at 32 rotations per second (rps), and the final recording

frequency was one point every second. Part of the experiments was repeated using a new

set of interference filters, one centered at 340 nm and one centered at 380 nm.

48

Fig. 2.7. Schematic representation of the experimental set up for the fluorescence

measurements.

In this case, the speed was set at 20 rps, the final recording frequency was one point

every 50 msec, and the traces reported in the figures were filtered off-line at 1 Hz. There

was no difference between the results of the two sets of experiments. The voltage applied

through the patch–clamp system was recorded simultaneously with the fluorescence

signals by the A/D board of the computer controlling the spectrophotometer. Fig. 2.8

illustrates the raw data of the fluorescence signals after background correction.

49

Fig. 2.8. Normalisation of fura-2 fluorescence signals. Raw data of the fluorescence

signals after background correction. All segments of the 380 nm component in the

absence of transients throughout the experiment (marked in black) were fitted with a

polynomial function (black line). Amplitudes of both fluorescence components were then

normalised to the resulting polynomial.

All segments of the 380 nm component in the absence of transients throughout the

experiment (marked in black) were fitted with a polynomial function (black line).

Amplitudes of both fluorescence components were then normalised to the resulting

polynomial.

50

Step by step procedure

Pulling of the patch pipette and bending of the very tip by 45° using a flame. Bending

serves to compensate for the 45° of the holder in order bring the pipette tip in a quasi-

vertical position and to obtain a symmetrical pipette orientation in the focal plane.

Back-filling of the pipette with pipette solution containing the fluorophore fura-2 (at a

concentration of 100 µM) in the dark. Generally, all operations handling fura-2 are done

in red ambient light to limit photobleaching.

High-resistance seal is obtained on isolated vacuoles. After reaching the whole-vacuole

configuration using combined voltage pulse (700 mV, 700 µs) and slight suction, patch

excision leads to the cytosolic side-out configuration.

The tip of the patch pipette is set into the focal plane of the microscope in bright field

(equipped with a red-light filter). With excitation light and photomultiplier on, the focal

plane is slightly adjusted (usually shifting upwards), in order to gain comparable

fluorescence signals between different experiments.

Simultaneous recordings of ionic currents and fluorescence signals upon voltage

stimulation are performed. Bath solutions are changed by means of a gravity-driven

perfusion system. The chamber volume of 0.1 ml is normally exchanged at least 20 times,

at a speed of about 1 ml*min-1

(slow perfusion procedure).

At the end of the experiment, background fluorescence is measured after setting the focal

plane far from the pipette; voltage offset is measured after removal of the membrane

patch by bringing the pipette tip in contact with the bottom of the recording chamber.

Protoplast and vacuole isolation

From Arabidopsis:

For isolation of mesophyll protoplasts, rosette leaves of 3–5 week old plants were used.

After careful removal of the lower epidermis, leaves were incubated for 30 min at 30°C

in enzyme solution containing 0.3% Cellulase, 0.03% Pectolyase Y-23 (Seishin), 1 mM

CaCl2, 1 mM MgCl2, 10 mM MES pH 5.5 with KOH π=600 mOsm with sorbitol.

Protoplasts were washed twice and resuspended in maintaing solution containing 1 mM

CaCl2, 1 mM MgCl2, 10 mM HEPES pH 7 with KOH, π=600 mOsm with sorbitol, by

centrifugation at 90 g for 1 min. Protoplasts were kept in the maintaing solution at room

temperature until used for experiments.

51

Vacuole release was observed under the microscope after adding EGTA to aliquots of

protoplast solution. The concentration of EGTA needs to be optimised to release vacuole

without compromising their stability. After settling of the vacuoles, solution was

carefully replaced by bath solution. Typically large and clear vacuoles are patched.

From carrots:

Fresh carrots were supplied by a local company and were used on the same day of the

experiments. Carrot taproot parenchymal tissue vacuoles were readily extruded into the

recording chamber by gently slicing the root cortex tissue into the standard bath solution.

Experimental solutions adopted in this work

Excised cytosolic side-out experiments:

Standard control solution 1: For the recordings of macroscopic currents standard, ionic

solutions were as follows: 200 mM KCl, 2 mM MgCl2, 2 mM CaCl2, 10 mM MES/Tris,

pH 5.5 in the pipette; 100 mM KCl, 2 mM MgCl2, 1 mM CaCl2, 1 mM dithiothreitol

(DTT), and 10 mM HEPES/Tris, pH 7.5 in the bath. DTT was added to the bath solution

in order to prevent channel rundown (Carpaneto et al. 1999; Scholz-Starke et al. 2004).

The osmolarity in both solutions was adjusted to 600 mOsm by the addition of D-

sorbitol.

Standard control solution 2: 200 mM KCl, 2 mM MgCl2, 10 mM MES/Tris, pH 5.5 in

the pipette; 100 mM KCl, 2 mM MgCl2, 1 mM CaCl2, 1 mM dithiothreitol (DTT) and 10

mM HEPES/Tris, pH 7.5 in the bath. The osmolarity in both solutions was adjusted to

600 mOsm by the addition of D-sorbitol.

Standard control solution 3: 200 mM KCl, 10 mM MES/Tris, pH 5.5 in the pipette; 100

mM KCl, 1 mM CaCl2, 1 mM dithiothreitol (DTT) and 10 mM HEPES/Tris, pH 7.5 in

the bath. The osmolarity in both solutions was adjusted to 600 mOsm by the addition of

D-sorbitol.

Standard control solution 4: For recordings of single SV channel conductance, standard

solutions were as follows: 100 mM KCl, 20 mM HEPES/5 mM KOH, pH 7.0 and 100

µM fura-2 in the pipette; 100 mM KCl, 2 mM CaCl2, 2 mM dithiothreitol

(DTT) and 20

52

mM HEPES/5 mM KOH, pH 7.0 in the bath. The osmolarity in both solutions was

adjusted to 420 mOsm by the addition of D-sorbitol. Free calcium in this solution

(considering a Ca2+

contamination of 2.5 µM measured by atomic absorption

spectroscopy) was estimated to be 5 nM (http:/maxchelator.stanford.edu).

Excised vacuolar side-out experiments:

Standard control solution 5: For recordings of macroscopic currents in vacuolar side-

out patch configuration, standard solutions were 100 mM KCl, 2 mM MgCl2, 2 mM

CaCl2, 1 mM dithiothreitol (DTT) and 10 mM HEPES/Tris, pH 7.5 in the pipette; 200

mM KCl, 2 mM MgCl2, 1 mM CaCl2, 10 mM MES/Tris, pH 5.5 in the bath. The

osmolarity in both solutions was adjusted to 600 mOsm by the addition of D-sorbitol.

Standard control solution 6: 100 mM KCl, 1 mM CaCl2, 1 mM dithiothreitol (DTT)

and 20 mM HEPES/1 M KOH, pH 7.0 in the pipette; 100 mM KCl, 1 mM CaCl2, 20 mM

HEPES/1 M KOH, pH 7.0 in the bath. The osmolarity in both solutions was adjusted to

420 mOsm by the addition of D-sorbitol.

Different concentrations of the flavonoids Naringenin, Naringin, Apigenin and Chrysin

(from Sigma-Aldrich) were reached by dilution from DMSO stock solutions. Both stock

solution and final solutions were freshly prepared just before the experiments to reduce

the possibility of degradation due to exposure to light.

Polyunsaturated fatty acids (PUFAs): Arachidonic Acid (AA: 5,8,11,14-all-cis-

eicosatetraenoic acid) and Docosahexaenoic acid (DHA: 4,7,10,13,16,19-all-cis-docosa-

hexa-enoic acid) were purchased from Sigma-Aldrich. Pure fatty acids were dissolved in

99.5% ethanol to a concentration of 100 mM and stored at −20°C until usage. Right

before experiments, stock solutions were diluted in pure control solution.

Does DMSO and Ethanol affect the SV currents?

In order to test if dimethyl sulfoxide (DMSO, stock solutions of Naringenin were

prepared in DMSO) has any effect on SV currents, we performed experiments by adding

DMSO at both cytosolic and luminal side. No significant effects were found when up to

53

0.2% (v/v) of DMSO was used (data not shown).

In order to test if Ethanol (stock solutions of PUFA were prepared in ethanol) has any

effect on SV currents, we performed experiments by adding ethanol at cytosolic side. No

significant effects were found when 0.9% (v/v) of ethanol was used (data not shown).

54

55

Results

56

57

Chapter 3.1. Fluorescence imaging and flavonoid localization

It is generally believed that flavonoids are synthesized on the multienzyme complexes at

the cytoplasmic surface of the endoplasmic reticulum; then they are transported to

different cell locations. However the location of the glycosylation process is not well

defined, so far. Therefore before investigating the effects of the flavonoid Naringenin on

the SV channel, we wanted to verify if non-glycosylated Naringenin is present in the

cytoplasm. To this purpose UDP-glucosyl transferase gene, which glycosilates

Naringenin into Naringin, was isolated. Arabidopsis thaliana UDP-glucoronosyl/UDP-

glucosyl transferase family protein (AT1G06000) (AtGT) was amplified using genomic

DNA as a template, digested with NcoI enzyme and cloned in frame with YFP as a fusion

protein in the pGreen vector. The right orientation was confirmed by digestion and

sequencing. Before studying the subcellular localization with this clone, western blot

analysis was performed to confirm that AtGT-YFP is expressed as a fusion protein.

Fig. 3.0. Cloning strategy of AtGT gene in the pGreen Vector. (A) Multiple cloning sites

of pGreen vector. (B) Vector map of pGreen 0029 vector.

A

B

58

Western Blot

To confirm the expression of fusion protein AtGT-YFP, western blot analysis was

performed using total protein extract from infected tobacco leaves. Total proteins were

extracted from the infected areas of the tobacco leaves and separated by 12% (w/v) SDS–

PAGE, transferred to a nitrocellulose membrane and hybridised with a polyclonal

antibody raised against YFP. The blot was then hybridised with an alkaline phosphate

conjugated secondary antibody and the detection was performed with the NBT/BCIP

colorimetric assay. This result confirms that the AtGT-YFP is expressed as a fusion

protein of 79 kDa.

Fig. 3.1. Western blot analysis of AtGT-YFP fusion protein. Total extracted proteins (1,2

and M are 3, 6 µg and protein marker respectively) were separated by 12% (w/v) SDS–

PAGE, transferred to a nitrocellulose membrane and hybridized with a polyclonal

antibody raised against YFP. The blot was then hybridized with an alkaline phosphate

conjugated secondary antibody and the detection was performed with the NBT/BCIP

colorimetric assay. Black arrow indicates the size of the fusion protein.

1 2 M

59

Subcellular localization of ATGT-YFP protein

To study the subcellular localization of AtGT-YFP, two experimental approaches were

used. First approach was based on PEG transformation of Arabidopsis mesophyll

protoplasts with pGreen AtGT-YFP vector; the second was based on agroinfiltration of

tobacco leaves. YFP-dependent fluorescence was analysed after 24 hours in case of

Arabidopsis mesophyll protoplasts and after 6 days after agroinfiltration of tobacco

leaves.

Fig. 3.2. Sub cellular localization of AtGT-YFP protein. (A) In Arabidopsis mesophyll

protoplasts it was observed the expression of fusion protein of AtGT-YFP in the

cytoplasm (left panel), chlorophyll (centre panel) and the merge of the two (right panel).

(B) Transient expression of AtGT-YFP in tobacco leaves showing the expression of

AtGT-YFP in the cytoplasm (left panel) and in the nucleus (centre panel); cytoplasmic

strands were also noticed (right panel).

The results obtained on Arabidopsis mesophyll protoplasts are illustrated in Fig. 3.2 A

showing the expression of AtGT-YFP fusion protein in the cytoplasm (left panel), of

A. Arabidopsis mesophyll protoplasts

B. Tobacco leaves

60

chlorophyll (centre panel) and the merge of the two signals (right panel). Similar results

were obtained in the confocal imaging of agro-infiltrated tobacco leaves as shown in Fig.

3.2 B where the expression of AtGT-YFP is mainly observed in the cytoplasm (left

panel) and in the nucleus (centre panel); cytoplasmic strands are also clearly visible (see

right panel).

These results indicate that the enzyme that glycosilates Naringenin into Naringin is

present in the cytoplasm, thus emphasising that a certain concentration of free Naringenin

should be present in the cytoplasm before its glycosylation.

61

Ch. 3.2. Macroscopic Slow Vacuolar currents

Typical voltage-dependent Slow activated Vacuolar (SV) currents, recorded in the

tonoplast from Arabidopsis mesophyll cells in excised cytosolic side-out patch

configuration, are shown in Fig. 3.3 A. These currents are mediated by the SV channel

and are exclusively activated at positive tonoplast voltages (Bertl et al. 1992). These

outward rectifying currents mainly represent the efflux of K+ from the cytosol to the

interior of the vacuole; here they are recorded in asymmetrical KCl, i.e KClvac=100 mM

and KClcyt =200mM. The asymmetric current-voltage characteristics displayed in Fig. 3.3

B illustrates the channel activation only at positive membrane potentials while almost no

current was detected at negative membrane potentials.

Fig. 3.3. Macroscopic Slow Vacuolar currents recorded in a tonoplast from Arabidopsis

mesophyll cells in excised cytosolic side-out patch configuration. (A) SV currents elicited

by a series of voltage steps ranging from –80 mV to +100 in +20 mV steps. Solutions

used were standard control solution 1; 200 mM KCl, 2 mM MgCl2, 2 mM CaCl2, 10 mM

MES/Tris, pH 5.5 in the pipette; 100 mM KCl, 2 mM MgCl2, 1 mM CaCl2, 1 mM

dithiothreitol (DTT), and 10 mM HEPES/Tris, pH 7.5 in the bath. Holding potential -50

mV, tail potential -50 mV. (B) SV current-voltage characteristics of the traces shown in

panel A. The I-V characteristics is constructed by plotting the average value of the

currents recorded during the last 50 ms (delimited by vertical bars) at each applied

voltage. Inset: schematic representation of the cation fluxes. The dashed circles represent

the cell that in vivo surrounds the vacuole. Positive currents represent cations entering

the vacuole and therefore leaving the cell.

62

Therefore the SV channel encoded by TPC1 (see chapter 1.2.2) is a strongly outward

rectifying channel that, in normal working conditions, only conducts positive currents at

positive trans tonoplast potentials. The positive currents represent cationic charges

flowing out of the cell into the vacuole (see Bertl et al. (1992) for the convention of the

current).

It is interesting to observe that the kinetics of SV current activation may significantly

change from one experiment to another. The reason for this variability (see Fig. 3.4) is

not known so far and may depend on a series of different parameters including the

membrane composition and the state of the plant or the tissue originating the vacuole.

Fig. 3.4. Macroscopic Slow Vacuolar currents recorded in two different carrot root

vacuoles under identical working conditions. A stimulation protocol (see Fig. 2.6 for

explanation) B and C panels show SV currents elicited by a series of voltage steps

ranging from –80 mV to +120 in +20 mV steps. Tail voltage -50 mV, Holding voltage -50

mV. Solutions used were standard control solution 3; 200 mM KCl, 10 mM MES/Tris, pH

5.5 in the pipette; 100 mM KCl, 1 mM CaCl2, 1 mM dithiothreitol

(DTT), and 10 mM

HEPES/Tris, pH 7.5 in the bath.

63

Ch. 3.3. Inhibition of the SV currents by Naringenin added at the

cytosolic side

Looking for compounds able to modulate the SV currents we challenged this channel

with Naringenin, a flavonoid playing an important role in plant cells.

Fig. 3.5 illustrates the results induced by Naringenin on the SV currents. The traces

represent the typical current response of the SV channel stimulated by a voltage step to

+80 mV. When the bath solution was replaced with an identical solution containing 1000

µM Naringenin, we observed a significant inhibition in the SV currents of vacuoles

obtained from either (A) Arabidopsis mesophyll protoplasts or (B) carrot roots. In both

cases when the Naringenin-enriched solution was replaced with a Naringenin-free bath

solution, the original currents were completely restored.

Fig. 3.5. Effects of the flavonoid Naringenin on Arabidopsis and carrot SV currents. (A)

SV currents in response to voltage stimulation at V= +80 mV in control conditions, in the

presence of Naringenin (1000 µM) and after washout (recovery) in vacuoles from

Arabidopsis mesophyll protoplasts and B in vacuoles from carrot roots in identical

working conditions as in A. Holding voltage -50 mV, tail voltage -50 mV. Solutions:

standard control solution 1 (see materials and methods).

Presumably the current inhibition mediated by Naringenin is a characteristic of the SV

channel that does not depend on the tissue and the plant species where this channel is

expressed. More experiments on vacuoles from other plants and tissues are needed to

further support this preliminary evidence.

64

Fig. 3.6. Time course of the inhibition induced by Nar, measured by the fast perfusion

procedure (see materials and methods). Vacuoles were isolated from carrot roots. Panel

A shows the inhibition and the recovery of two typical SV currents (blue and green

traces) when they are perfused with a solution supplemented with 1000 µM Nar

(indicated by the top black line) in identical working conditions. Continuous voltage at

V= +80 mV. The grey trace superimposed to the green trace represents the time needed

by the fast perfusion setup to change the solution bathing the vacuole. This was obtained

in test experiments perfusing the patch pipette with a diluted solution (1:10 by water) and

returning back to standard control solution 1. The times needed for the inhibition and

recovery of the current at different voltages are plotted in panels B (on) and C (off),

respectively. Applied voltages from +40 to +120 mV in +20 mV steps. Solutions used

were standard control solution 1 (see materials and methods). The continuous lines

65

connect different points from the same vacuole. Different experiments are represented by

diverse symbols.

In order to clarify the mechanism of interaction of Naringenin with the membrane phase,

we also measured the time course of inhibition induced by Nar by the fast perfusion

procedure applied to vacuoles isolated from carrot roots under continuous voltages.

The results are reported in the Fig. 3.6. Panel A shows typical SV currents at V= +80 mV

and the inhibition and recovery of these currents while perfusing the membrane patch

with a solution supplemented with 1000 µM Naringenin (black line on the top) in two

different experiments (blue and green traces) under identical working conditions. Clearly,

in some vacuoles the SV currents displayed a faster inhibition and recovery while others

showed a slow time course (compare top and bottom panels of Fig. 3.6 A). Notably, the

variability of the kinetics of current inhibition and recovery induced by Nar spans almost

three orders of magnitudes (Fig. 3.6 B).

Interestingly, the time course of current inhibition was faster compared to the recovery

(compare B and C in Fig. 3.6) in the order of a few seconds. These data suggests that the

absorption/desorbtion of Nar from the membrane phase changes significantly from

vacuole to vacuole as it possibly depends on a series of parameters controlling the

membrane chemico-physical characteristics.

66

Ch. 3.4. Quantitative analysis of SV channel inhibition by Naringenin

In order to gain a better understanding of the effects induced by Naringenin on the SV

currents, we performed dose-response experiments in the concentration range from 100

µM to 5 mM Naringenin in carrot root vacuoles. A dose-dependent decrease of the SV

channel activity was measured. For example at V= +80 mV, on the addition of [Nar]

concentrations as small as 150 µM, the residual SV current (INar/Icontrol) was about 0.2,

while 5 mM Naringenin decreased the SV currents to 5% of the control. This modulation

of the current was fully reversible. These results are illustrated in Fig. 3.7, where

INar/Icontrol is plotted as a function of Naringenin concentration. A curve fitting with the

Hill equation (see eq. 2.1):

INar /Icontrol = 1/(1+([Nar]/Kh)n)

gave a half inhibition concentration of (0.44 ± 0.03) mM. In eq. 2.1 [Nar] is the

Naringenin concentration, Kh and n (=1.4 ± 0.1, in Fig. 3.7) are the Hill half inhibition

constant and Hill coefficient, respectively.

Fig. 3.7. Quantitative analysis of SV current inhibition by Naringenin: INar /Icontrol plotted

as a function of Naringenin concentrations. We obtained a half inhibition constant Kh =

(0.44 ± 0.03) mM with a Hill equation where n=1.4 ± 0.1. These data values were

derived normalizing the steady state current measured in the presence of Nar to the

steady state level obtained in control conditions, i.e. before the addition of Nar.

67

Fig. 3.8. Quantitative analysis of SV current inhibition by Naringenin: Gnorm plotted as a

function of Naringenin concentrations. A) Normalized conductance, with respect to

control conditions plotted vs Nar concentrations in the voltage range from +30 to +120

mV; note that only three voltages are represented for clarity reasons. For all the

potentials under investigation a fit curve was performed and the Hill inhibition constant

(Kh) and Hill coefficient (n) were obtained. B) Kh values, derived from the Hill fit shown

in panel A are plotted against the applied membrane voltages. C) Hill coefficients (n) are

plotted against the applied membrane voltages. The value of n is >1 at all the

investigated voltages.

68

Similar results were obtained when Gnorm, the conductance normalized with respect to the

control conditions in the absence of Nar was plotted as a function of Nar concentration

using the tail currents*. The results obtained in these experiments are illustrated in Fig.

3.8 panel A. Panel B shows Kh values, derived from the Hill fit shown in panel A, plotted

against the applied membrane voltages. With this fit we obtained Hill coefficients n>1 at

all the investigated voltages (Fig. 3.8 C).

These results constitute an indirect indication of a specific interaction between

Naringenin and the SV channel: indeed a positive cooperation between Nar and the

channel suggests that more than one Nar molecule is necessary to inhibit the channel and

the binding of one flavonoid molecule further facilitates Nar binding to the protein.

* Note that data in Fig. 3.8 were retrieved from voltage-dependence analysis illustrated in

Fig. 3.9.

69

Ch. 3.5. Voltage dependence characteristics of the SV channel in the

presence of cytosolic Naringenin

A further insight into the SV channel properties modified by Naringenin can be obtained

investigating the voltage-dependence of the channel in the presence and absence of this

flavonoid. Fig 3.9 A illustrates the currents mediated by the SV channel of carrot root

vacuoles at different voltages ranging from -80 mV to +120 mV in +10 mV steps, in

control conditions (left traces), in the presence of 500 µM Naringenin (centre) and in

recovery conditions (right traces) (note that only one out of two traces are reported in Fig.

3.9 A for clarity reasons). Indeed, macroscopic currents were strongly reduced when 500

µM Naringenin was added to the bath solution, while, upon removal of Naringenin,

current inhibition was fully reversed. In Fig. 3.9 B normalized conductances derived

from instantaneous tail currents are plotted as a function of the main voltage pulse (for

definition of tail potential see Fig. 2.6 and Fig. 3.4) and fitted with a single Boltzmann

function. Data analysis confirmed that the addition of Nar affects the voltage dependence

of the channel shifting the activation to more positive voltages, increasing s and

decreasing the maximum conductance. These parameters are reported in table 1.

Fig. 3.9 C shows the time required for half-activation (t1/2 norm) of the current at different

membrane potentials. The values obtained in control/recovery conditions are comparable

to those obtained in the presence of Nar. Instead, the time constant of deactivation (τnorm)

derived from tails currents in the presence of Nar are faster compared to the values

derived in control/recovery conditions.

These data indicate that Nar, determines a modification of the energy needed for

activation by the voltage sensor of the channel, which, in the presence of the flavonoid,

opens at much more positive membrane potentials. So far is not clear whether this

process occurs via a direct or indirect interaction of the flavonoid with the channel.

70

Fig. 3.9. Inhibition of the SV currents by Naringenin is voltage dependent. (A)

Macroscopic SV currents, elicited by a series of voltage steps ranging from -80 to +120

mV in +10 mV steps in control (left panel), in the presence of 500 µM Naringenin (centre

panel) and after recovery (right panel) (note that only one out of two traces was reported

for clarity reasons). Holding potential -50 mV, tail potential -50 mV. Solutions used were

standard control solution 3 (see materials and methods). (B) Normalized conductances

derived from instantaneous tail currents were plotted as a function of the main pulse and

fitted with a single Boltzmann function (see materials and methods). Note that

normalization is done with respect to the control. Data points represent the mean of nine

experiments in control and at least three experiments at various concentrations of Nar;

error bars represent SEM. (C) Normalized time required for half-activation of the

current (t½ norm) is plotted against the applied membrane potential. Data points represent

mean values (± SEM) determined in control conditions (open circles; n = 9), in the

presence of 300 µM Nar (closed circles; n = 3) and 500 µM Nar (closed triangles; n =

3). Data points are normalized to the value of control at V =+ 60 mV. (D) Normalized

time constants of current deactivation, τnorm are plotted against the tail potential. Data

points represent mean values (± SEM) of τnorm determined in control conditions (open

circles; n = 9) and in the presence of 300 µM Nar (closed circles; n = 3) and 500 µM

Nar (closed triangles; n = 3). Data points are normalized to the value of control at V = -

60 mV.

71

Table 1 V1/2 (n)

(mV ± SEM)

s

(mV ± SEM)

GMax(Nar)/GMax(control)

Control 18 ± 4 (9) 19 ± 1 1

Nar= 300 µM 33 ± 7 (3) 27 ± 4 0.70 ± 0.07

Nar= 500 µM 60 ± 11 (3) 36 ± 4 0.63 ± 0.06

Nar= 1000 µM 62 ± 7 (3) 39 ± 1 0.40 ± 0.07

Table 1: Half-activation potentials,V1/2, s (=RT/zF) and ratios of the maximum

conductances in the presence of and absence of Naringenin Gmax(Nar)/Gmax(control),

derived from eq. 2.4, are reported in the table as function of Nar concentration. Note that

an increase of the “s” means a decrease of the apparent gating charge z.

Nevertheless a comparison of the activation and deactivation kinetics suggests that the

effects mediated by Nar are not due to a simple shift of the activation characteristics of

the channel as the activation and deactivation kinetics are modified in a different manner.

This is incompatible with a single shift of the activation characteristics of the channel to

be ascribed, for example, to a modification of the surface charges at the

membrane/protein-water interface.

72

Ch. 3.6. Naringenin effects on the SV single channel conductance

To investigate whether Naringenin binds within the pore of the SV channel and affects

the single channel conductance, we performed single channel experiments on vacuoles

from carrot roots with 500 µM Nar. Two voltage protocols were used. In the first

approach the current response to continuous voltage stimulation, e.g. at V= -70 mV (like

in the Fig. 3.10) was recorded. At this voltage the open probability of the SV channel is

very low and consequently the single channel signals can be recorded in more appropriate

conditions i.e. the recordings contain one or a few channels. Under these conditions,

control solution (standard control solution 4) was replaced with an identical solution

supplemented with 500 µM Nar (Fig. 3.10).

Fig. 3.10. Naringenin does not change the amplitude of the conductance of the SV single

channel. Single channel current transitions in control conditions, in the presence of 500

µM Nar and after recovery. C, O1 and O2 represent the closed state, one channel and two

channels open respectively. Solutions used were, 100 mM KCl, 20 mM HEPES/5 mM

KOH, pH 7.0 and 100 µM fura-2 in the pipette; 100 mM KCl, 2 mM CaCl2, 2 mM

dithiothreitol (DTT), and 20 mM HEPES/5 mM KOH, pH 7.0 in the bath (standard

control solution 4). Applied potential V= –70 mV.

Control

73

Fig. 3.10 illustrates the single channel recordings in the absence and in the presence of

500 µM Nar. Top panel shows the single channel openings at V= -70 mV. When we

replaced the control solution with an identical solution supplemented with 500 µM Nar

the single channel conductance did not change appreciably (compare C and O1 of control

and Nar), while the channel remained closed most of the time and the mean open time

decreased drastically. The original firing pattern of the channel was completely restored

when we replaced the Nar supplemented solution with the control solution (see recovery

in Fig. 3.10).

Fig. 3.11. A) Single channel current–voltage relationships obtained after the application

of a fast voltage-ramp in control conditions (black trace) superimposed to data points

(filled circles) obtained under continuous voltage stimulation in the presence of 500 µM

Nar. Cytosolic side-out excised patch from the vacuoles of carrot roots. Solutions used

were, 100 mM KCl, 20 mM HEPES/5 mM KOH, pH 7.0 and 100 µM fura-2 in the

pipette; 100 mM KCl, 2 mM CaCl2, 2 mM dithiothreitol (DTT), and 20 mM HEPES/5 mM

KOH, pH 7.0 in the bath (standard control solution 4). B) Voltage protocol applied to the

membrane patch to record single channel characteristics by means of a voltage-ramp.

Voltage frequency= 2.5 V/S.

The second protocol was based on fast voltage ramps in order to get access to the

conductance at far positive voltages where single channels cannot be resolved because of

high number of channels that open simultaneously. If the ramp is sufficiently fast, one

can span the entire voltage range of interest starting from negative voltage, completing

the cycle before more channel superimpose to the channel signals recorded at negative

voltages. In Fig 3.11 the single channel conductance obtained from voltage ramps is

74

represented as a black trace superimposed to the single channel data (filled circles)

obtained from the continuous voltage protocol in the presence of 500 µM Nar. The data

(n=204 mean ± SD) from Fig. 3.10 are also represented at –70 mV.

These data provide further support to the results illustrated in Fig. 3.10 and guarantee that

the single channel conductance does not change in the presence or absence or Nar in the

whole range of the investigated voltages. Moreover ramp signals illustrated in Fig. 3.11

refer to one single protein unit. While single channel recordings obtained under

continuous voltage protocols might be representative of successive openings of one or

more single protein units and therefore request a time consuming statistical analysis.

Note the smaller single channel conductances at positive voltages with respect to the

negative voltages. This is due to the working conditions (i.e. high cytosolic, low vacuolar

calcium concentrations). The reversal voltage is about 0 mV in accordance with

symmetrical KCl solutions that give a VNernst = 0 mV.

75

Ch. 3.7. Simultaneous Fluorescence measurements

and patch-clamp recordings

The SV channel is not very selective to different cations. Indeed, it has been shown that it

is permeable to Na+, K+ and Ca2+ ions (see Chapter 1.2.2). In order to verify whether

Nar affects both K+ and Ca2+ conductances, we performed experiments by combining

the patch-clamp technique with fluorescence measurements using the fluorophore fura-2

on Arabidopsis thaliana mesophyll and carrot root vacuoles. Permeation of calcium was

demonstrated by conductance measurements (Pottosin et al. 2001) and reversal potential

determination in the presence of varying concentration ratios of potassium and calcium:

(Allen and Sanders 1996).

Fig. 3.12. Calcium permeation through Slow Vacuolar channels isolated from carrot

roots. Simultaneous fura-2 fluorescence and macroscopic current recordings on a

cytosolic side-out tonoplast patch. Fura-2 signals resulting from alternate excitation at

340 and 380 nm are shown. B) After reaching the cytosolic side-out excised patch

configuration, 10-s voltage pulses were applied according to the voltage scheme. The

respective current responses are shown in the panel C. Solutions used were standard

control solution 4 (see materials and methods).

76

The experimental approach was based on the idea by Neher (1995) to simultaneously

record whole cell currents (using the patch-clamp technique) and the fluorescence signal

of the calcium indicator dye fura-2. Unless otherwise indicated, our experiments were

conducted in the cytosolic-side-out excised patch configuration; the fluorescence signal

was collected focusing the photomultiplier to the tip of the pipette loaded with the

fluorescent dye. In these working conditions fluorescence signals (Fig. 3.12 A) and Slow

Vacuolar currents (Fig. 3.12 C) were recorded simultaneously upon voltage stimulation

(Fig. 3.12 B). Stimulation of SV currents coincided with characteristic changes in

fluorescence in vacuoles from carrot roots. The emission upon excitation at 380 nm

decreased, while the signal at 340 nm increased increasing the membrane potential (Fig.

3.12 A). This behavior is indicative of calcium permeation through the channel and

subsequent binding of Ca2+

to the fluorescent dye (Grynkiewicz et al. 1985).

To verify whether fluorescence is strictly linked to calcium transport mediated by SV

channel, experiments were conducted in vacuoles from Arabidopsis thaliana wild-type

and tpc1 mutant plants that lack SV channel activity (Peiter et al. 2005). Similar to carrot

root vacuoles, voltage stimulation of a cytosolic-side-out membrane patch derived from

an Arabidopsis wild-type vacuole elicited both macroscopic SV currents and

simultaneous fluorescence changes; instead, vacuoles from tpc1 knockout plants, besides

lacking SV channel activity, did not show any variation of the fluorescence signals,

neither upon prolonged voltage stimulation (Fig. 3.13). These results confirmed that the

changes in fura-2 fluorescence amplitudes indeed reflected Ca2+

currents through SV

channels. Details on this approach and the corresponding results are reported in Gradogna

et al. (2009).

77

Fig. 3.13. The fura-2 fluorescence response is absent in Arabidopsis tpc1 knockout

vacuoles. Changes in fura-2 fluorescence were recorded upon voltage stimulation of a

cytosolic side-out tonoplast patch derived from an Arabidopsis thaliana Col-0 vacuole

(wild type, left panels). No fluorescence response was present when patches from tpc1-2

knockout vacuoles were tested (right panel). The respective current responses are shown

in the lower panels. Vacuoles were isolated from leaf mesophyll protoplasts. Solution

used was standard control solution 4 (see materials and methods).

In order to investigate the effects of flavonoids in our working conditions, control

solution was replaced by solutions containing different concentrations of Nar. Fig. 3.14

illustrates the results obtained in such patch-clamp experiments performed on carrot root

vacuoles. Fluorescence signals observed at 380 (Fig. 3.14 A upper panel) and 340 nm

(Fig. 3.14 A middle panel) are shown together with the applied membrane potential

(protocol, Fig. 3.14 A lower panel). When 200 µM Naringenin was added to the bath

solution (central panel in A) we could observe a reduction in fluorescence (middle panel

A) with respect to the control condition (left panel A). This effect was fully reversible, as,

when we replaced 200 µM Naringenin solution with the bath solution; fluorescence was

completely restored (right panel A).

78

Fig. 3.14. Patch-clamp recordings with fluorescence measurements on carrot root

vacuoles. (A) Changes in fura-2 fluorescence in control conditions were elicited by a

voltage step to +80 mV from a holding potential of -80 mV (see lower panel illustrating

the applied voltage). In the presence of 200 µM Naringenin fluorescence signal was

reduced. The effect was fully reversible after washout of Naringenin (recovery). (B)

Changes in fura-2 fluorescence in control conditions were elicited by a voltage step to

+80 mV from a holding potential of -80 mV (see lower panel). In the presence of 1000

µM Naringenin, fluorescence signal was almost absent. The effect was fully reversible

after washout of Naringenin (recovery). (C) SV currents responses upon voltage

79

stimulation in control and in the presence of 200 µM Naringenin. (D) SV currents

responses upon voltage stimulation in control and in the presence of 1000 µM

Naringenin. Solutions used were standard control solution 4 (see materials and

methods).

The results obtained with 1000 µM Naringenin are illustrated in Fig. 3.14 B.

Fluorescence signal observed at 380 (upper panel) and 340 nm (middle panel B) are

shown together with the applied membrane potentials (lower panel B). On the addition of

1000 µM Naringenin, the fluorescence signal was completely abolished. Current records

reported in panels 3.14 C and D integrate the fluorescence experiments illustrated in

panel 3.14 A and B. Furthermore Fig. 3.14 C illustrates SV channel current response on

prolonged voltage stimulation in control and 200 µM Naringenin. Finally Fig. 3.14 D

illustrates SV channel current response on prolonged voltage stimulation in control and in

the presence of 1000 µM Naringenin.

The substantial equivalence of electrophysiological and fluorescence signals definitely

demonstrate that flavonoid Naringenin affects both K+ and Ca2+ conductances.

As illustrated in the Discussion section this observation provides a substantial evidence of

the fact that Naringenin does not directly bind within the pore or in the proximity of the

ionic pathway.

80

Ch. 3.8. pH and DTT effects on the modulation of the SV channel by

Naringenin

It has been reported that, owing to its hydrophobic nature Nar can partition into the

hydrophobic core of the lipid bilayer (van Dijk et al. 2000) modulating the membrane

fluidity (Arora et al. 2000). It has also been demonstrated that the flavonoid Quercetin

(QCT) can penetrate into the lipid bilayer to different extents at different pHs. This

depends on the neutral form and liposolubility of QCT at acidic pHs and to its

deprotonated form at more alkaline pHs (Movileanu et al. 2000). An increasing value in

the pH of the bath solution also promotes a more hydrophobic character of Naringenin

(van Dijk et al. 2000). On this basis we decided to verify whether a variation of the pH

has any effect of the inhibition of the SV channel; therefore we changed the pH of

external solution from 7.5 to 6.5.

Under these conditions we challenged the SV currents with 500 µM Nar.

Fig 3.15. pH and DTT modify the inhibition of SV currents by Naringenin. (A) The graph

represents the residual SV currents (INar /Icontrol) when 500 µM Nar was added to the bath

at pH 7.5 and pH 6.5 (n=3). Solutions used were standard solution 1 (see materials and

methods). (B) Residual SV currents (INar /Icontrol) when DTT was increased from no DTT

to 1 mM DTT and 5 mM DTT in the presence of 500 µM Nar (n=3). Error bars indicate

SEM.

81

Panel 3.15 A represents the residual SV currents (INar /Icontrol) when 500 µM Nar was

added at pH 7.5 and pH 6.5 (n=3). Our results demonstrate that a decrease of the pH

determines an increase of the inhibition of the SV currents by Nar; thus it supports the

possibility that Nar interacts with the SV channel through the phospholipid bilayer.

It has been reported that DTT increases the open probability of the SV channels, possibly

interacting with sulphydryl groups of cysteine residues located at the cytoplasmic side of

the channel (Carpaneto et al. 1999).

As the control standard solution used in normal conditions contains 1 mM DTT (that

prevents SV current rundown) and flavonoids have well known antioxidant properties,

we decided that it was worthwhile to investigate the effects induced by different DTT

concentration on SV currents in the presence of Naringenin. Fig. 3.15 B illustrates the

results obtained varying DTT concentration. The graph represents the residual SV

currents (INar /Icontrol) when DTT was absent or in the presence of 1 mM DTT or 5 mM

DTT and 500 µM Nar.

Although these experiments produced data with no significant differences, one can

observe a general trend where the inhibition of the SV currents induced by Nar decreases

increasing DTT (n=3).

More experiments are needed, in order to understand the effects of reducing agents that

may compete or interact with other antioxidants, changing their modulation of the SV

channel properties; however this detailed investigation goes beyond the present scope of

this thesis. Therefore these preliminary studies on the role of DTT on Nar action are

preliminary to a detailed characterization of the effects of flavonoids and PUFAs (see Ch.

3.13) in diverse redox conditions (Angelova et al. 2009).

82

Ch. 3.9. Effects of vacuolar Naringenin on the SV channel

We also investigated the effects induced on the SV channel by Naringenin added at the

luminal side, in comparison with those induced by Nar at the cytosolic side. Fig. 3.16 A

illustrates SV currents in response to a voltage step to +80 mV in control, in the presence

of 300 µM Naringenin and in recovery conditions in vacuoles from Arabidopsis

mesophyll cells. The average residual SV current (INar/Icontrol) was 0.55 ± 0.03 SEM

(n=5). Fig. 3.16 B illustrates similar experiments performed on a vacuole from carrot

roots using 1000 µM Nar. Macroscopic currents were strongly reduced when Naringenin

was added to the bath solution also in these working conditions; upon removal of

Naringenin, current inhibition was fully reversed.

Fig. 3.16. Luminal Naringenin decreased the SV current. Excised luminal side-out patch

showing voltage dependent outward currents mediated by the SV channel at positive

membrane potential. (A) SV current response to voltage stimulation at V= +80 mV

(control), in the presence of Naringenin (300 µM) and after washout (recovery).

Vacuoles were isolated from Arabidopsis thaliana mesophyll cells. Solutions used were

standard solution 5 (see materials and methods). (B) SV current response to voltage

stimulation at V= +80 mV (control), in the presence of Naringenin (1000 µM) and after

washout (recovery). Holding voltage -50 mV, tail voltage -50 mV. Vacuoles were isolated

from carrot roots. Solutions used were standard solution 6 (see materials and methods).

These data demonstrate that Nar presents similar modulation characteristics of the SV

channel activity at both sides of the membrane. Therefore, presumably the binding site is

not located at the membrane water interface (either at the cytosolic or the luminal side),

but it is readily accessible by both membrane sides. These considerations lead us to

investigate whether the glycosylated form of Naringenin has also similar consequences

on the channel.

83

Ch. 3.10. Naringin does not inhibit the SV currents

As the glycosylated form of Naringenin, Naringin, has different hydrophobicity

characteristics with respect to the non-glycosylated form, we investigated whether

Naringin also has the ability to change the transport properties of the SV channel.

Fig. 3.17. Structure of Naringenin and Naringin

Fig. 3.18 A illustrates the consequences of the addition of Naringin at the cytosolic side

of the vacuole: it is shown the SV current response to voltage stimulation at V= +80 mV

in control conditions, in the presence of 1000 µM Naringin and after the recovery. In

vacuoles isolated from Arabidopsis thaliana mesophyll cells, Naringin displayed a

negligible inhibition of the SV currents (n=3). Fig. 3.18 B illustrates the effects of

Naringin at the vacuolar side; it is shown the SV channel current response on voltage

stimulation at +80 mV in control, in the presence of 1000 µM Naringin and in recovery

conditions.

Also in these conditions Naringin did not show any modulation of the SV currents (n=5).

Naringin Naringenin

84

Fig. 3.18. The glycosylated flavonoid Naringin only slightly inhibits the SV currents in

vacuoles isolated from Arabidopsis thaliana mesophyll cells. (A) SV channel response to

voltage stimulation at V= +80 mV (control), in the presence of cytosolic Naringin (1000

µM) and after washout (recovery). Holding potential -50 mV, tail potential -50 mV.

Solutions were 200 mM KCl, 2 mM MgCl2, 2 mM CaCl2, 10 mM MES/Tris, pH 5.5 in the

pipette; 100 mM KCl, 2 mM MgCl2, 1 mM CaCl2, 1 mM dithiothreitol (DTT), and 10 mM

HEPES/Tris, pH 7.5 in the bath (standard control solution 1). (B) SV channel response to

voltage stimulation at V= +80 mV (control), in the presence of vacuolar Naringin (1000

µM). Solutions were 100 mM KCl, 2 mM MgCl2, 1 mM CaCl2, 1 mM dithiothreitol

(DTT), and 10 mM HEPES/Tris, pH 7.5 in the pipette; 200 mM KCl, 2 mM MgCl2, 1 mM

CaCl2, 10 mM MES/Tris, pH 5.5 in the bath solution (standard control solution 5).

In conclusion only the non-glycosylated form, Naringenin does affect the channel

activity. We hypothesize that the difference in the effects produced by Naringenin and

Naringin depends on the higher hydrophobicity of the non-glycosylated flavonoid with

respect to the glycosylated form that carries a relatively large hydrophilic sugar terminal

attached to the Naringenin scaffold.

B. Vacuolar side A. Cytosolic side

85

Ch. 3.11. Other cytosolic flavonoids also inhibit the SV currents

In order to investigate whether other flavonoids sharing structural similarities with Nar

produce effects on the SV channel currents, we challenged a few vacuoles with the

flavonoids Apigenin and Chrysin (Fig. 3.19).

Fig. 3.19. Structure of flavonoids used in this study.

The results obtained in such experiments are illustrated in Fig. 3.20. Panel A shows the

typical response of the SV channel after a voltage stimulation to V= +80 in the presence

of 300 µM Nar and in recovery conditions. Panels B and C show the responses of SV

channel (still at V= +80) in the presence of 300 µM Apigenin and Chrysin respectively as

well as in recovery conditions. These results indicate that flavonoids, sharing structural

similarities with Nar, determine a similar modulation of the SV currents.

Naringenin

Chrysin

Apigenin

86

Fig. 3.20. Other cytosolic flavonoids modulate the SV currents similarly to Nar. A shows

the typical response of SV channel at V= +80 in control conditions and in the presence of

300 µM Nar. B and C show the responses of SV channel (V= +80) in the presence of 300

µM Apigenin and Chrysin. Holding potential -50 mV, tail potential -50 mV. All the

experiments were performed on vacuoles from carrot roots. Solutions used were standard

control solution 1 (see materials and methods).

87

Ch. 3.12. Effects of Naringenin on the SV channel

from transparent testa mutant plants

Interestingly the seed coats of Arabidopsis thaliana transparent testa mutant plants are

affected by a deficiency in the flavonoid biosynthetic pathway (Shirley et al. 1995; Buer

and Muday 2004; Buer and Djordjevic 2009). In order to verify whether SV currents in

these flavonoid deficient plants display different modulation by Nar, we performed some

experiments on these plants. Two mutant plants were investigated, namely tt4 and tt5,

which lack the flavonoid biosynthetic pathway and Naringenin, respectively. As the tt

mutant plants belong to the Landsberg erecta ecotype, these plants were also included in

this study.

Fig. 3.21. Cytosolic Naringenin inhibits SV currents from transparent testa mutant

plants. A) The SV channel response to voltage stimulation at V= +80 mV in control, in

the presence of 1000 µM Naringenin and in recovery conditions in vacuoles from

mesophyll cells of Landsberg erecta plants. B) and C) illustrate the results obtained in tt4

and tt5 mutant plants respectively; the same working conditions as in A. Holding

potential -50 mV, tail potential -50 mV. Solutions used were standard control solution 1

(see materials and methods).

A. Landsberg erecta

B. tt4 mutant plant

C. tt5 mutant plant

88

Fig. 3.21 illustrates the results obtained in such experiments. Panel A illustrates the SV

channel response to voltage stimulation at +80 mV in control, in the presence of 1000

µM Naringenin and in recovery conditions in vacuoles from mesophyll cells of

Landsberg erecta plants. B and C illustrate the results obtained in tt4 and tt5 mutant

plants respectively in the same working conditions as in A.

We observed a reversible decrease in the activity of the currents when 1000 µM Nar was

added to the cytosolic side, thus demonstrating that Nar displays similar modulation

capability of the SV current also in vacuoles isolated from plants that lack flavonoids.

89

Ch. 3.13. Effects of polyunsaturated fatty acids on the SV currents

It has been reported that Naringenin can partition in the hydrophobic core of the lipid

bilayer (van Dijk et al. 2000) modulating the membrane fluidity (Arora et al. 2000). It has

also been demonstrated that flavonoids could penetrate into the lipid bilayer to different

extents at different pHs. For example flavonoid Quercetin is deeply embedded in planar

lipid bilayers at acidic pH (Movileanu et al. 2000) while, on the contrary, it interacts at

the water-lipid interface at physiological pHs (Terao et al. 1994; Movileanu et al. 2000).

Indeed our pH experimental data are consistent with these results (see Ch. 3.8), thus it

supports the hypothesis that Nar interacts with the SV channel through the phospholipid

bilayer. To verify this possibility, we investigated the interaction of the SV channel with

polyunsaturated fatty acids (PUFAs). Indeed many studies are aimed at verifying

modulation of mammalian ion channels by PUFAs. For review on this topic see Boland

and Drzewiecki (2008).

Two PUFAs, namely Arachidonic Acid (AA: 5,8,11,14-all-cis-eicosatetraenoic acid) and

Docosahexaenoic acid (DHA: 4,7,10,13,16,19-all-cis-docosahexaenoic acid), were used

in this study (Fig. 3.22).

Fig. 3.22. Structure of PUFA included in this study. Arachidonic Acid (AA) and

Docosahexaenoic acid (DHA).

These experiments, performed on vacuoles from carrot root vacuoles, are illustrated in

Fig. 3.23. Top Panel in A shows the typical response of the SV channel to voltage steps

at V= +80 in control conditions and in the presence of 1 µM AA. Middle and bottom

panels show the inhibition of the currents on the addition of 3 µM AA and 10 µM AA,

respectively. In all the cases original currents were restored when the AA-supplemented

solutions were replaced with the control solution. Similar results were obtained when

DHA (see panel B) was used in place of AA in the same experimental conditions as

shown in Fig. 3.23 A.

Arachidonic acid Docosahexaenoic acid

90

Fig. 3.23. Cytosolic PUFAs inhibit the SV currents. (A) Top panel shows the SV current

in control conditions, in the presence of 1 µM AA and after washout (recovery). Middle

panel shows the SV channel response in control conditions, in the presence of 3 µM AA

and after washout (recovery). Bottom panel shows the SV channel in control conditions,

in the presence of 10 µM AA and after washout (recovery). (B) Top panel shows the SV

current in control conditions, in the presence of 3 µM DHA and after washout (recovery).

Bottom panel shows the SV channel response in control conditions, in the presence of 10

µM DHA. Vacuoles were isolated from carrot roots. Applied voltage V= +80 mV holding

potential -50 mV, tail potential -50 mV. Solutions used were standard control solution 3

(see materials and methods).

Panel B top traces shows the typical responses of the SV channel to voltage stimulations

at V= +80 in control conditions and in the presence of 3 µM DHA. The bottom panel B

91

shows the current inhibition mediated by 10 µM DHA. In both cases original currents

were restored when the DHA solutions were replaced with control solution.

In conclusion our data strongly suggest that the SV channel is modulated by flavonoids

but also by other compounds, such as PUFAs, that possibly interact with the protein

through the lipid double layer. However, so far it is not clear whether these compounds

modify the SV channel properties directly or changing the organization and fluidity of the

bilayer.

More experiments are needed to clarify this point. For example capacitance measurments

may provide indications on the presence of PUFA layers at the membrane-water

interface.

92

Ch. 3.14. Voltage dependence characteristics of the SV channel in the

presence of cytosolic Arachidonic Acid

A further insight into the channel properties modified by Arachidonic Acid can be

obtained investigating the voltage-dependence of the channel in the presence and absence

of AA. In Fig 3.24 A the normalized conductances obtained from instantaneous tail

currents were plotted as a function of the main voltage pulse and fitted with a single

Boltzmann function.

Data analysis confirmed that also the addition of AA affects the voltage dependence of

the channel shifting the activation to more positive voltages, increasing “s” and

decreasing the maximum conductance. The values of these parameters in different

working conditions (i.e. in the absence and in the presence of different AA

concentrations) are illustrated in table 2.

Fig. 3.24 B shows the time required for half-activation (thalf norm) of the current at different

membrane potentials. The values obtained in the presence of AA are faster compared to

the values derived in control/recovery conditions. Fig. 3.24 C shows that the time

constant of deactivation, derived from conductances in the presence of AA, is faster

compared to the values derived in control conditions. AA possibly interacts with the

voltage sensor of the channel or changes the membrane properties in a manner that

determines the activation of the SV channel at much more positive membrane potentials.

93

Fig. 3.24. The inhibition of the SV current induced by Arachidonic Acid is voltage

dependent. (A) Normalized instantaneous tail currents were plotted as a function of the

activating potential and fitted with a single Boltzmann function. Data points represent the

mean of eight experiments in control and three experiments from different vacuoles at

various concentrations of AA as indicated. Error bars represent SEM. (B) Normalized

half-time of the current activation is plotted against the applied potential. Data points

represent mean values (± SEM) determined in control conditions (open circles; n = 8), in

the presence of 1 µM AA (closed red circles; n = 3) and 3 µM AA (closed gray triangles;

n = 3). Data points are normalized to the values obtained at V =+ 60 mV. C) Time

constants of current deactivation τ are plotted against the tail potential. Data points

represent mean values (± SEM) of τ determined in control conditions (open circles; n =

8) and in the presence of 1µM (closed red circles; n = 3) and 10 µM AA (closed gray

triangles; n = 3).

94

Table 2 V1/2 (n)

(mV ± SEM)

s

(mV ± SEM)

GMax(AA)/GMax (control)

Control 39 ± 5 (8) 21 ± 1 1

AA= 1 µM 58 ± 2 (3) 32 ± 2 0.83 ± 0.06

AA= 3 µM 48 ± 14 (3) 36 ± 4 0.51 ± 0.13

Table 2: Half-activation potentials, V1/2, s (=RT/zF) and the ratios of the maximum

conductances Gmax(AA)/Gmax(control), in the presence and absence of AA derived from

eq. 2.4, are reported in the table as a function of AA concentration. Note that an increase

of the “s” implies a decrease of the apparent gating charge, z.

However, differently from what observed on the addition of Naringenin, a comparison of

the activation and deactivation kinetics shows that AA slows down both the activation

and deactivation times. Nevertheless, also in this case (AA mediated inhibition) the

slower kinetics of current activation is incompatible with a shift of the activation

characteristics due to surface charge modifications at the membrane-water interface.

95

Ch. 3.15. Quantitative analysis of the SV channel inhibition

by Arachidonic Acid

In order to gain a better understanding of the effects induced by Arachidonic Acid on the

SV currents, we performed a dose-response characterization in the concentration range

from 1 to 10 µM of AA in carrot root vacuoles. The results obtained in such experiments

are summarized in Fig. 3.25 where Gnorm is plotted as a function of AA concentrations.

Panel A shows the dose-response inhibition for selected voltages from + 60 to + 120 mV.

Note that only four voltages are represented for clarity reasons. For all the potentials

under investigation a fit curve was performed, which allows to determine a series of Hill

constants, Kh and Hill coefficient, n. Panel B displays the Kh values, plotted against the

applied membrane voltages. The Hill coefficient (n) was plotted against the applied

membrane voltages; as shown in panel C, n>1 at all the investigated voltages.

One can observe that the data illustrating AA modifications of the SV currents, in Figs.

3.24 and 3.25, qualitatively remind the results obtained on the addition of Naringenin,

illustrated in Figs. 3.8 and 3.9. Only the Hill inhibition constant, Kh, is significantly

smaller (more than two order of magnitude) in the case of AA with respect to Nar.

Clearly, much lower concentrations of AA (in the micromolar range) are sufficient to

induce an inhibition of the current comparable to that observed at millimolar

concentrations of Naringenin.

96

Fig. 3.25. Quantitative analysis of the SV current inhibition by Arachidonic Acid: Gnorm

plotted as a function of Naringenin concentrations. A) Dose-response inhibition in the

voltage range from + 50 to + 120 mV; note that only four voltages are represented for

clarity reasons. For all the potentials under investigation a fit curve was performed,

which allows to fit the data sets with a series of Hill coefficients and Hill functions. B) Kh

values derived from the Hill fit (shown in panel A) are plotted against the applied

membrane voltages. C) The Hill coefficient (n) was plotted against the applied membrane

voltages; note that n>1 at all the investigated voltages. Error bars indicate SEM.

97

Ch. 3.16. Arachidonic Acid effects on the SV single channel conductance

To investigate whether AA binds within the pore of the SV channel, we performed single

channel experiments on vacuoles from carrot roots with 3 µM AA. The protocol was

based on fast voltage ramps in order to get access to the conductance at far positive

voltages where single channel cannot be resolved because of high number of open

channels. In Fig 3.26 the single channel currents obtained from voltage ramps in the

absence (black trace) and in the presence of 3 µM AA (red trace).

Fig. 3.26. Single channel current–voltage relationships obtained after the application of

a fast voltage-ramp in control (black trace) superimposed to data (red trace) obtained in

the presence of 3 µM AA in cytosolic side-out excised patch from the vacuoles of carrot

roots. Solutions used were standard control solution 3 (see materials and methods).

These data demonstrate that the amplitude of the single channel conductance does not

change in the presence or absence or AA.

Control

98

99

Discussion

100

101

The majority of plant cells contains organelles called vacuoles surrounded by a

membrane called tonoplast. In mature plant cells, the vacuole occupies up to 90% of the

cell volume. The transport across the tonoplast is crucial for cell homeostasis and to

maintain the turgor pressure which is fundamental in the regulation of cell expansion and

growth (Maeshima 2001). The vacuole stores the essential compounds, minerals,

nutrients, metabolites and different ions required for cell homeostasis. Different proteins

of the tonoplast allow the ions to selectively permeate according to their electrochemical

gradients (channels) or at the expense of other energy sources (active transporters).

The Slow Vacuolar channel

The patch-clamp technique offers the possibility of a direct examination of ion channel

properties in isolated vacuoles. The SV channel was shown to be calcium activated in

vacuoles from beet storage tissues (Hedrich and Neher 1987) and was subsequently

characterized in vacuoles from many plant species (Allen and Sanders 1997; Carpaneto et

al. 1997; Scholz-Starke et al. 2005a; Dziubinska et al. 2008). Nowadays it is generally

accepted that the SV channel is ubiquitous in higher plants.

Typical features of the SV channel are the outward rectification at elevated

unphysiological cytoplasmic Ca2+

concentrations and the slow activation. Indeed it is

well known that the SV currents recorded in a patch-clamp experiment require

unphysiologically high cytosolic and low vacuolar calcium concentrations for full

activation. The SV channel is a non-selective cation channel, permeable to both

monovalent and divalent cations (Pantoja et al. 1992; Ward and Schroeder 1994; Allen

and Sanders 1996; Gambale et al. 1996; Ivashikina and Hedrich 2005). It has been

demonstrated that the SV channel, besides being permeable to potassium ions, is also

permeable to calcium (Gradogna et al. 2009). It has also been demonstrated that the SV

channel is encoded by the TPC1 gene (Peiter et al. 2005), comprising 12 alpha-helices

and two pore segments.

It is well known that the SV channel is modulated by a series of different parameters and

compounds, internal and external to the vacuole; beside calcium, Mg2+

, protons, redox

potentials and various metals like Zn2+

and Ni2+

and the glycosidic antibiotic Neomycin

(Scholz-Starke et al. 2006) are able to modify the properties of the SV channel. However,

as none of these parameters was able to transform the SV into a channel that activates at

physiological potentials and calcium concentrations, we aim at investigating unexplored

102

working conditions or compounds, which are able to further modulate this channel.

Specifically we plan to find out endogenous plant substances (for example flavonoids,

glycosidic compounds, polyamines and other charged or uncharged molecules, reducing

and oxidizing molecules, etc...) which might be able to modify the voltage activation

threshold of this channel towards more physiological conditions or that provide

information on the role and the nature of this enigmatic channel.

Flavonoids and polyunsaturated fatty acids

Flavonoids are plant pigments or secondary metabolites present in all plant species. They

are one of the largest group of secondary metabolites represented by more than 8000

different compounds (Harborne and Williams 2000; Ververidis et al. 2007). It was

reported that the flavanone Naringenin (Nar) is present in all plants where it plays a

central role in the biosynthetic pathway (Lepiniec et al. 2006).

Flavonoid localization

It is generally believed that flavonoids are synthesized on the multienzyme complexes at

the cytoplasmic surface of the ER from where they are transferred to different locations.

However their glycosylation site is not well known so far. Therefore we firstly

investigated the presence of non-glycosylated Nar in the cytoplasm and then studied the

effects of Naringenin on the SV channel by the patch-clamp technique. In order to

confirm that free Naringenin is present in the cytoplasm, UDP-glucosyl transferase gene,

which glycosilates Naringenin into Naringin, was isolated. Arabidopsis thaliana UDP-

glucoronosyl/UDP-glucosyl transferase family protein (AT1G06000) (AtGT) was

amplified using genomic DNA as a template, cloned in frame with YFP as a fusion

protein in the pGreen vector. The subcellular localization was investigated by PEG

transformation of Arabidopsis mesophyll protoplasts with pGreen AtGT-YFP vector and

by agroinfiltration of tobacco leaves. It was shown that the AtGT-YFP fusion protein is

mainly expressed in the cytoplasm, thus confirming that non-glycosylated Naringenin is

present in the cytoplasm.

Modulation of SV channel by flavonoids

We investigated the modulation of the SV channel by the flavonoid Nar in vacuoles from

carrot roots and Arabidopsis thaliana mesophyll cells by using the patch-clamp

technique. When Nar was added to the cytosolic bath solution in carrot root vacuoles we

recorded a dose-dependent reversible decrease in SV channel activity described by the

103

Hill equation with n>1 and a half block concentration of about 0.4 mM. Similar effects in

the channel activity were observed in vacuoles from mesophyll cells of Arabidopsis

thaliana.

Naringenin changed the characteristics of SV channel activation “unfortunately” shifting

toward more positive membrane potentials the activation threshold and decreased the

deactivation times. This implies a diverse modification of the voltage sensor during the

opening and the closure processes.

In accordance with the macroscopic current measurements we also verified that Nar did

not change the amplitude of the single channel conductance but modified the opening

probability of the channel favoring its closed state. Indeed, the mean open time decreased

and the mean closed time of the channel increased significantly in the presence of

cytosolic Nar.

In order to verify whether Nar affects both potassium and calcium conductance, we

performed experiments by combining the patch-clamp technique with fluorescence

measurements using the fluorophore fura-2. When 200 µM Nar was added to the bath

solution we could observe a reversible decrease in the current as well as in the

fluorescence signals. In the presence of 1000 µM Nar the current and fluorescence

signals were completely and reversibly abolished. Therefore Nar affects both K+ and Ca

2+

permeation.

When we investigated the effects induced by 1000 µM Nar added at the luminal side we

observed a reversible inhibition of the initial SV current of about 40% in vacuoles from

both carrot roots and mesophyll cells of Arabidopsis thaliana.

We also performed experiments on SV currents from carrot root vacuoles with other

flavonoids, like Apigenin and Chrysin, which share similar structural properties with

Naringenin. We observed a reversible decrease in the activity of the channel when 300

µM Apigenin and Chrysin were added at the cytosolic side.

Finally, in order to verify the specificity of Nar inhibition, we performed experiments

using Naringin, the glycosylated form of Naringenin, on vacuoles from Arabidopsis

thaliana mesophyll cells. Interestingly Naringin did not induce any significant decrease

of the channel activity even at a concentration of 1000 µM added both at the cytosolic

side or the luminal side. In conclusion only the non-glycosylated form, Naringenin did

affect the channel activity. We hypothesize that this difference depends on the higher

104

hydrophobicity of the nonglycosylated flavonoid with respect to the glycosylated form

that carries a relatively large hydrophilic sugar terminal.

Indeed it has been reported that Nar can partition in the hydrophobic core of the lipid

bilayer (van Dijk et al. 2000) modulating the membrane fluidity (Arora et al. 2000). It has

also been demonstrated that flavonoids could penetrate into the lipid bilayer to different

extents at different pHs. For example the flavonoid Quercetin (QCT) is deeply embedded

in planar lipid bilayer intercalating the flexible acyl chains at acidic pH (Movileanu et al.

2000) where QCT is neutral and highly hydrophobic. Instead at more alkaline pHs, where

QCT is deprotonated, the flavonoids binding site is restricted to the hydrophilic domain

of the membrane (Terao et al. 1994; Movileanu et al. 2000). In order to further

investigate these aspects we changed the pH of the external solution from 7.5 to 6.5.

Under these conditions, we challenged the SV currents with 500 µM Nar. Interestingly at

pH 6.5 the inhibition of SV the channel was larger compared to the inhibition at pH 7.5.

All these indications suggest that Nar might interact with the SV channel through the

phospholipid bilayer. More studies are needed to support this possibility.

Modulation of the SV channel by PUFAs

For this reason we investigated the interaction of the carrot root SV channel with

polyunsaturated fatty acids (PUFAs). Arachidonic acid (AA, 5,8,11,14- all-cis-

eicosatetraenoic acid) and Docosahexaenoic acid (DHA, 4,7,10,13,16,19-all-cis-

docosahexaenoic acid) were added at the cytoplasmic side of the vacuole to investigate

the response of the SV channel. Similarly to Naringenin micromolar concentrations of

PUFAs induced a dose-dependent reversible decrease in SV channel activity.

In conclusion our data strongly suggests that the SV channel is modulated by flavonoids

and by other compounds, such as PUFAs, that interact with the protein through the lipid

bilayer. It is not clear whether these compounds modify the SV channel properties

directly or changing the organization and fluidity of the bilayer.

Indeed a specific drug might modulate the channel properties either interacting with the

protein at the water-membrane interface or, in alternative, binding either at a site internal

to the protein or deeply embedded into the lipid bilayer. In general the modifications of

channel properties may typically occur as a consequence of a direct binding and

occlusion of the pore or by a modification of either the permeation pathway or the gating

mechanisms. The first process is typically fast and is influenced by the nature of the

105

permeating ions present in the water solution when the channel is challenged with the

drug. Instead the second is a slower process that depends on the physico-chemical

characteristics of the protein as well as on the membrane structure. Indeed it is well

known that several hydrophobic compounds, such as dihydropyridines, are able to block

the L-type calcium channels via a lipid mediated interaction (Gambale and

Dellacasagrande 1994). This block and the reversibility of the blocking mechanism

mediated by different dihydropyridines, in some cases, may give rise to long lasting

inhibition of the channel.

Several cases are reported where phospholipids, PUFAs, phosphoinositides, cholesterols

and other endogenous compounds participate, alone or together with beta subunits, in the

modulation of the transport characteristics of the channel.

For example it was reported (Roberts-Crowley and Rittenhouse 2009) that Arachidonic

Acid modulates the properties of the L-type calcium channel CaV 1.3b in superior

cervical ganglion neuron in a manner that highly resembles the action of M1 muscarinic

receptor. AA decreases the open probability of CaV 1.3 (Liu and Rittenhouse 2000) and T

type CaV 1.3 (Talavera et al. 2004) channels, increasing the dwell time in the closed state

without affecting the single channel conductance amplitude (Liu and Rittenhouse 2000).

Notably, this behavior strongly reminds the action of Naringenin and AA on the voltage-

dependent calcium permeable SV channel (see Materials and Methods, Chs. 3.6 and

3.13). It was also demonstrated that AA inhibits and changes the kinetics of CaV 1.3

channel directly and not through metabolites or the intermediates of CaV ß subunits.

Interestingly palmitic acid may compete with AA for an inhibitory site, responsible for

the AA-mediated current-decrease, located in the most hydrophobic part of the channel.

This indicates that the membrane composition and AA may have competing or

synergistic effects on the channel.

Another example of modulation of channel properties by intracellular AA is the

inhibition of the voltage-dependent K+ channels KV 1.4 or KV 4.2 which is prevented by

antioxidants (Angelova et al. 2009). This suggests the presence of diverse oxidative sites

playing important roles in the properties of these K+

voltage-dependent channels. As it is

well known that also Slow Vacuolar currents are sensitive to the redox potential, in future

experiments the possibility that AA affects the SV channel properties through a

106

modification of redox reaction sites should be monitored. Likewise one should also

investigate any potential interaction between flavonoids and antioxidant AA.

The polyunsaturated fatty acids AA and its amide, Anandamide, produce a rapid

inactivation of KV channels (Oliver et al. 2004). The relatively slow modification induced

by AA and Anandamide strongly suggest that the interaction is mediated by the lipid

membrane and prevented by Cs+ and NH

4+ present in the permeation pore.

CONCLUSIONS

Naringenin effects on the SV channel, namely the kinetics and the voltage dependence of

macroscopic currents (Chs. 3.3 and 3.5) and single channel data (Ch. 3.6), the

comparable inhibition of both Ca2+

and K+ permeation (Ch. 3.7), the dependence of

inhibition on the pH of the cytosolic solution (Ch. 3.8), the comparable activity when

Naringenin is added either at the cytosolic or at the vacuolar side of the tonoplast (Ch.

3.9), the absence of inhibition by Naringin (Ch. 3.10) as well as PUFAs and Naringenin

comparable effects point to a significant role played by the membrane composition,

organization and fluidity in modulating the interaction of the SV channel by flavonoids

and polyunsaturated fatty acids.

More experiments are needed to quantify the interaction between Naringenin and

Arachidonic Acid with specific protein binding sites as well as the contribution of the

phospholipid hydrophobic core and the role of the surface charges and/or the organization

of the lipid polar heads at the membrane water interface. For example, the Arabidopsis

mutant plant fou2 presents a single aminoacid substitution on the SV channel that shifts

the threshold of channel activation towards negative voltages. fou2 has been shown to

upregulate oxylipins in the jasmonate pathway where linolenic acid is released from the

plasma membrane to be converted in jasmonic acid.

On the other side, PUFAs inhibit the SV channel with high affinity and in a voltage

dependent manner. We can speculate that, if linolenic acid acts similarly to AA and

DHA, its removal from the vacuolar membrane too might unlock the SV channel. In any

case a mechanism of lipid mediated feedback involving the SV channel may occur.

These results are surprising and interesting as they open new perspectives on the role of

the SV channel in vivo in different membranes with diverse phospholipid compositions.

Therefore biophysical characterization opens new perspectives on the physiological

properties of the SV channel in different plants and tissues.

107

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Acknowledgments

Firstly I would like to thank GOD Almighty for giving me the opportunity to come to

Italy.

I would like to express my gratitude to my supervisor Dr. Franco Gambale for being very

understanding, patient, friendly, for his tremendous support and for believing in me and

trusting that I would learn the “patch-clamp” technique and do some good work. I

appreciate his vast knowledge and skills in many areas and his assistance in writing

reports (Ph.D. annual reports and this thesis).

A very special thanks to Dr. Armando Carpaneto without whose motivation and

encouragement I would not have reached this point. It was through his persistence,

understanding and kindness that I was able to complete my Ph.D. thesis.

I would like to thank Prof. Fiorella Lo Schiavo for giving me an opportunity to register

with her in Padua and work here in Genoa. Many thanks go to all my lab mates and

friends who helped me a lot during my stay in Italy. I would like to thank Prof. B.C.

Tripathy for helping me out during the hard times of my life.

A special thanks to Stella de Robertis for creating a friendly atmosphere and for helping

in establishing myself in Genoa. I would like to thank the administrative staff for being

kind, and for putting up with my bad Italian at crucial times. I would like to thank all the

technical staff who helped me in innumerous ways directly or indirectly.

I would also like to acknowledge the European Union Marie-Curie RTN Research

Network VaTEP that provided financial support for my study.

I would like to thank all my family members and friends in India for all their support

during these years. Finally and most importantly, I would like to thank my sweet wife,

Anila Sunandini (who takes away my pressures just by smiling) for her tremendous

support, love and understanding.