dedicated to Julius von Sacks, the founder of plant physiology on the centenary of his death.

Part of his studies and lectures concentrated on movement in plants.

Since plants cannot leave their natural habitat, they only survive by adapting to their surroundings. In the greenhouse, by using gas analyzers, the interrelation between photosynthesis and the water status of the plant

can be studied in detail. Carbon dioxide uptake and loss of water vapor is recorded online and displayed on the screen.

For the experiment, a leaf is mounted into a recording chamber. An air stream with precisely adjusted CO2 content and humidity is passed over the leaf surface. By continuously monitoring the changes in the gas composition,

the photosynthetic CO2 uptake and transpiration can be determined. During the day, we follow the relation between CO2 uptake and water vapor loss.

At sunrise, CO2 fixation is induced, often reaching a maximum at noon, to drop again by sunset. These fluctuations are accompanied by a rise and fall in leaf water loss. For every CO2 molecule gained, about 600 water molecules are lost.

The plant is, however, equipped to avoid this dilemma. By taking advantage of adjustable pores, plants are able to adapt CO2 uptake and water loss to the availability of water in the soil.

In the leaf, the gas exchange between the intercellular spaces and the atmosphere is controlled by the external boundary layer, the epidermis. Within the lower epidermis, the stomata can be recognized as microscopically small pores.

When they are open, stomata mediate the influx of CO2 and release of water vapor.

The epidermis, covered with an almost gas-impermeable wax, is now isolated for microscopic analysis. Following an increase in their volume, guard cells are pulled apart and the stoma is opened.

The molecular mechanisms involved in stomatal movement will now be explained. Stomatal opening requires iron and water uptake from the extracellular space as well as synthesis of malate following starch degradation in the chloroplast.

Potassium and chloride are transported into the guard cells and together with malate accumulate in the vacuole. The uptake of these ions and synthesis of malate generate an osmotic gradient, which in turn drives the influx of water.

The accumulation of osmotically active particles, the turgor and volume increase, pulls the two guard cells apart.

During stomatal closure, this process is reversed. Through release of potassium salts and water, guard cells reduce their volume. Potassium transport proteins in the plasma membrane represent essential elements in volume regulation and thus in the control of stomatal aperture.

In order to measure the activity of these transport proteins in individual guard cells, epidermal strips are peeled off and incubated with cell wall degrading enzymes.

During gentle decomposition of the cell wall, the cytoskeleton reorganizes within a few hours.

Meanwhile, the guard cells begin to be released from the epidermal tissue. Organelles such as the nucleus, the vacuole and the chloroplasts remain intact, though rearranged by cytoplasmic streaming.

With progressing cell wall degradation, the cell attains a nearly spherical shape. Protoplasts appear. The plasma membrane is now freely accessible for biophysical analysis by the patch

clamp technique, a method developed to monitor the activity of a single ion channel molecule. For this purpose, protoplasts are perfused with an isotonic potassium salt solution.

After the reference electrode is introduced into the electrolyte solution, the recording electrode is attached to the protoplast. Gentle suction pulls an omega-shaped plasma membrane vesicle into the tip of the electrode. If this membrane patch is magnified beyond the resolution limit of light and electron

microscopy, shown here as a model, the molecular structure of the unit membrane is evident. Embedded in the phospholipid bilayer, the channel protein. When closed, the channel is impermeable to potassium ions.

With its instantaneous opening, the channel conducts potassium ions across the membrane barrier into the cytoplasm. The movement of positively charged ions creates an electric current.

Upon channel closure, the current is interrupted immediately. So far, these gating processes have been shown in slow motion. Now, they will be stepwise adapted close to their real-time kinetics. The individual open times last from a few to almost 30 milliseconds.

Within a millisecond, a channel conducts about 10,000 potassium ions into the guard cell. As a result, the opening and closing of the channel is accompanied by characteristic current fluctuations and is therefore measurable.

The resolution of currents through single ion channels in the picoampere range is only possible by the patch-clamp technique.

This biophysical method requires high technical effort, an extremely low-noise current amplifier and fast computer-based analysis. In the following, the prerequisites for a patch-clamp experiment will be explained in detail.

In the initial step, microelectrodes are prepared. Glass capillaries of about 1 mm diameter are centered within a heating coil.

The glowing filament melts the glass and a weight pulls it out to a very fine tip. Its inner diameter is now only in the order of 1 micrometer. In the following step, this capillary is further optimized by smoothing and adding a special coat under the microscope.

The rims of the open tip are smoothed by holding it close to a hot filament. The smooth surface is essential for the formation of a tight contact between the glass and cell membrane in the experiment. Coating the outer surface of the capillary with silicon rubber reduces the conductance of the glass.

Thus, the electrical properties of the membrane become measurable. Prior to the recording, the capillary is filled with an isotonic electrolyte.

It contains potassium ions as the main charge carrier. The contact between the electrolyte of the capillary and the amplifier is established through insertion of a silver electrode.

The recording electrode is now ready for the experiment. Together with the reference electrode and the guard cell protoplast, an electric circuit is formed in the recording chamber. The protoplasts are at the focal plane of the microscope. Using micromanipulators, the patch pipette is carefully attached to the protoplast.

The contact between the pipette and the plasma membrane is greatly improved by gentle suction. As a result, the electrical resistance increases continuously. An acoustic signal underlines this process.

After the membrane has covered the tip of the patch pipette completely, the resistance between the recording and reference electrodes increases to the gigaohm range. The flow of current is now almost completely suppressed.

The resistance is now essentially determined by the membrane patch. The contribution of the large membrane area outside the pipette can be neglected since resistance is reciprocally dependent on the surface. As a result of the tight mechanical contact between the membrane and the glass walls of the micro pipette tip,

an intact patch can be excised from the cell. The intracellular surface of the membrane is now experimentally accessible. Thus, the electric potential difference between the internal and external membrane surface can be adjusted.

The electric current through the membrane patch is shown in the equivalent circuit. The potassium flow, and thus the electric current I, is obstructed by the membrane resistance R.

The potential difference between U0 and UM drives an electric current I, which is proportional to the voltage drop between U0 and UM and reciprocal to the resistance R.

At 10,000-fold amplification of the current signal, however, fluctuations in the picoampere range become visible, indicating the opening and closing of single potassium channels in the guard cell protoplast. The potassium flux through an open channel, the current amplitude, depends on the membrane potential.

By increasing the electric potential on the cellular side of the membrane towards more negative values, the driving force for the current of positively charged potassium ions rises.

In a graph, the voltage dependence of the current through an open potassium channel can be demonstrated. The current amplitude is related to the voltage applied.

The current through the open channel is a function of the voltage applied and its conductance, G. In analogy to an ohmic resistor, the potassium conductance G of the channel protein can be deduced from the slope of the linear current-voltage relation.

In comparison to potassium channels in nerve cells, the guard cell channels represent only one-third of the conductance, that is, a conductance of 5 to 8 picoSiemens.

Within half an hour, 100 to 1,000 channels transport so many potassium ions into the guard cells that potassium salts accumulate. The events in a membrane patch characterized by pulsations in the potassium current thus reflect the opening and closing of potassium channels.

This principle of a microvalve is not only encountered in stomatal movement, but represents a general mechanism for potassium transport in plants as well as other organisms.

Hubert Ziegler's research focuses on solute fluxes in plants, their molecular mechanisms, regulation and functional importance.

For plants, potassium is an essential nutrient.

It is taken up from the soil and utilized for growth, development and movement. Potassium ions contribute significantly to the generation of turgor and in this way drive cell elongation.

Following potassium uptake into the root, the cell wall expands and the cell volume increases. Internodes elongate as a result of increasing accumulation with potassium ions. Leaves develop. In the leaf, turgor-driven microvalves, the stomata, mediate gas exchange between the plant and the atmosphere.

As in the root, shoot and leaf cells, potassium uptake into guard cells is mediated by potassium channel activity. In the following sequence, the strategies and techniques required for the molecular analysis

of a plant potassium channel are demonstrated using guard cells from Visscher Faber. The use of a laser scanning microscope and fluorescent dyes allows clear visualization of the nucleus. In this organelle, the genetic information for the three-dimensional structure of a potassium channel is stored by nucleic acid sequences or DNA.

Following the activation of a particular gene, its DNA sequence is transcribed into messenger RNA and transported to the cytoplasm. This copy process is called transcription.

In the following step, cytoplasmic mRNA is translated into its corresponding amino acid sequence. The potassium channel protein folds into its three-dimensional structure and inserts into the plasma membrane. The structure and function of a potassium channel reflects the role it plays in a specific cell type, tissue or developmental stage.

To isolate the gene for a guard cell potassium channel, first the stomata-rich lower epidermis is peeled off the leaf and frozen in liquid nitrogen.

In this way, a picture of the genes which were active before freezing is captured in the mRNA composition of the frozen tissue.

The cell walls of guard cells are mechanically robust and resist a turgor pressure of up to 10 bars. A pestle and mortar are used to break the frozen cells. In the following series of steps, the mRNA will be isolated and purified.

The cell lysate is transferred into a reaction vial and incubated with a denaturing buffer.

Under these conditions, the RNA degrading enzymes are inactivated. Cell walls are separated from the soluble fraction by centrifugation. Under the hood, the resulting supernatant, containing RNA and also DNA and proteins, is treated with phenyl chloroform mixture.

The preparation is mixtured and again centrifuged.

This organic extraction separates the nucleic acids from the proteins. In the lower organic phase, accumulation is mainly of proteins. In the interphase of DNA.

The water-soluble supernatant is enriched in total RNA. The mRNA content of this fraction is only about 1 to 2%. The supernatant is transferred to a new vial.

Sodium acetate is added. In the presence of sodium acetate and isopropanol, the now insoluble RNA precipitates.

During centrifugation, order the RNA sediments to the bottom of the reaction vial. Next, the solubilized total RNA is subjected to an affinity purification process.

The RNA fraction contains mRNA, which is characterized by a chain of adenine nucleotides at the 3' end. This poly-A tail is a structural marker, found neither in ribosomal RNA nor in transfer RNA.

On addition of magnetic beads covered with polythymidine chains, the polyadenine tails of the mRNA form hydrogen bonds with the polythymidines on the magnetic beads.

A strong magnet is used to attract this complex to the side wall of the reaction tube.

In this way, the poly-A containing mRNA is separated from the other RNAs. In the lab, the isolation of mRNA from the total RNA is performed in reaction tubes, which are exposed to a magnet implanted in the sample holder.

Non-bound RNA is removed with a pipette.

In the presence of an elution buffer, the mRNA is released from the magnetic bead complex. It will then be separated electrophoretically in accordance with size and charge.

Electrophoresis is performed on an agarose gel. A toothed spacer is used to form pockets in the polymerizing gel where the mRNA can later be loaded. Before loading the mRNA, the two dyes, xylin cyanol and bromphenol blue, are added to the RNA.

This mixture of RNA and dye is pipetted into the pockets of the agarose gel. Due to the high density of the glycerol-containing loading buffer, the RNA mixture sediments to the bottom of the pockets. Total RNA and molecular weight standards are included in the gel as references of known size.

On application of a constant voltage, electrophoresis is initiated. The movement of the two dyes indicates the progressing separation of the still invisible RNA. Within the electrical field, RNAs of different lengths move at different rates, like the differently mobile dyes.

After incubation with ethidium bromide, UV light excites RNA fluorescence. To determine the size distribution of the RNA, the internal molecular weight standards are used.

These encompass bands between 9,000 and 200 bases in length. The total RNA is characterized by two prominent bands, the ribosomal 28S and 18S bands.

The mRNA emits diffuse fluorescence in the range between 400 and 4,000 bases. The identification of the potassium channel gene is accomplished using a DNA copy of the mRNA.

This approach allows the several thousand mRNAs, which are normally expressed simultaneously, to be stored as a cDNA library. Since a large number of genes are always expressed, the isolated fraction of mRNA encodes many proteins of different size and function.

The generation of a cDNA library requires several experimental steps in the lab. These steps will now be demonstrated using a single mRNA molecule as an example.

The process of generating a cDNA library begins with the synthesis of a single-stranded cDNA corresponding to the mRNA.

In the first step, a polythymidine primer is bound to the polyadenine tail of the mRNA. Hydrogen bonds form between the two complementary bases. Symbols now represent the nucleotides.

The mRNA is composed of the nucleotides cytosine, uracil, adenine, and guanine. The reverse transcriptase, which attaches to the poly-T primer, synthesizes the corresponding cDNA

strand by incorporating single nucleotides from the solution, resulting in an RNA-DNA hybrid. The synthesis of the first strand is completed.

To form the second strand, RNase H is initially required. It specifically degrades the template mRNA strand.

Next, the activity of a DNA polymerase is required. The gaps created by the RNase H are filled by the DNA polymerase.

All enzymes involved work hand in hand.

Now a ligase is activated. The ligase repairs gaps which still exist in the now continuous second DNA strand.

All the enzyme reactions which have been shown here one after the other actually take place simultaneously under strict temperature control in a thermocycler. After purification of the now double-stranded DNA, a ligase is again required.

It attaches adapters to the ends of the cDNA, an essential step for the cloning of the DNA.

During this process, single-stranded overhanging ends, or sticky ends, are added to the double-stranded DNA. In a last step, the individual cDNA strands will be ligated into plasmids which will serve as vectors.

Specific restriction enzymes prepare the plasmids for the subsequent cloning of the cDNA. The free ends of the plasmids now perfectly match the ends of the cDNA.

Ligases link the ends to one another. Following these enzyme reactions, each mRNA is represented by a DNA copy at least once in the cDNA library.

Using the functional assay, the cDNA library will now be screened for potassium channel genes. For this purpose, yeast cells are transformed with the plasmids. Initially, the cDNA plasmid mixture is added to a yeast suspension culture.

For the transformation, this suspension is transferred to a cuvette. Baker's yeast is well suited to screen for a potassium channel gene since

yeast growth, like that of plants, depends on the ability to take up potassium. The transformation process takes place in an electroporator.

Two metal plates attached to the side walls of the cuvette act as capacitor plates across which an electrical field is applied to induce the uptake of the plasmids into the yeast cells. In response to a short voltage pulse, pores a few nanometers in diameter form in the plasma membrane of the yeast cells.

These pores allow the entry of plasmids into the yeast cells. Some of the incorporated plasmids will contain the potassium channel gene.

The transformed yeast cells are now transferred onto a selection medium. For transformation, a yeast mutant, which lacks the ability to take up potassium, has been used.

This allows the identification of those yeast cells which have received a gene encoding a potassium channel.

During the screening process for functional complementation of yeast growth, transformed cells are plated on medium-low in potassium. Yeast growth is monitored over several days.

A multi-step screen finally identifies yeast strains which grow on media containing minimal potassium concentration and therefore must be functionally expressing a plant potassium channel gene. A yeast like this will grow, divide, and form a colony of several million cells.

Growing colonies are isolated and incubated for two to three days in a temperature-controlled chamber.

At the end of the screen, each yeast colony, or each clone, carries a functional potassium channel gene. For the following sequence analysis, a large quantity of plasmids is required, so single colonies are transferred from agar plates into liquid culture.

To further increase the plasmid yield, an intermediate step is included, in which the plasmids are transferred to Escherichia coli.

In the presence of an optimal nutrient supply, the E. coli clone grows in a shaking incubator to produce a dense cell suspension, a basic requirement for the isolation of potassium channel-containing plasmids. The sequence of the potassium channel gene is based on the chain termination method developed by Sanger.

Each of the previously performed sequencing reactions is terminated by a modified form of bases A, C, G, or T.

The resulting mixture of different length DNA is now applied to a high-resolution polyacrylamide gel in an automatic sequencer. In the sequencer, the fluorescently labeled DNA fragments, which differ in length by just a single nucleotide, are separated. A laser scans the gel and detects the fluorescently labeled DNA fragments to determine the sequence of nucleic acids in the potassium channel gene.

The computer assembles the overlapping DNA sequences and unravels the primary structure of the potassium channel gene.

The pattern recognized by the laser is digitized by the computer, making it possible to determine the sequence of the bases automatically.

In one scan, the series of bases in the first four A, C, G, and T lanes is monitored. With each new sequencing reaction, the nucleotide chain of the potassium channel gene continuously increases.

Eventually, the complete sequence is known. The molecular structure of the channel gene is now identified. The potassium channel gene was isolated from guard cell mRNA and identified using a yeast complementation assay.

It is expressed in the nucleus of the guard cell. After transcription, mRNA is shuttled into the cytosol. Here, at the ER, ribosomes translate the ribonucleic acid sequence into an amino acid chain.

The subunits of the potassium channel are assembled and incorporated into the plasma membrane. A guard cell is then able to take up potassium ions to regulate guard cell volume and stomatal movement.

This process is the result of changes in channel activity and density. In this way, the activity of potassium channel genes and other cell-specific genes controls plant movement, growth, and differentiation, as well as adaptations to changes in their environment.

Following the identification of the structure of plant genes, a study of the molecular mechanisms regulating these diverse processes is possible. Modern plant physiology will have a major impact.

Erwin Neher and Klaus Aschke have provided the groundwork for studying biological processes from the phenomenon to the molecule.

In contrast to the spontaneous growth of crystals, the growth, movement, and differentiation of living cells is mediated by protein molecules.

The first plant potassium channels were characterized and cloned from stomatal guard cells.

Ion channels, proteins of the plasma membrane, mediate the exchange of solutes and information between cells and their environment. The molecular blueprint, and thus the structure of these proteins, is stored in the nucleus. Following identification of the gene structure of the first potassium channels,

large-scale genome projects continuously deliver DNA sequences encoding new potassium channels. This has become possible with the aid of a new generation of automatic sequencers and ultra-fast computers. It is possible to generate an evolutionary tree of potassium channels, which includes bacteria, fungi, plants, and even humans.

The distance between individual branches of the tree indicates the genetic diversity between the various potassium channels and symbolizes the relationships within the three groups of organisms.

When comparing the amino acid sequences of all the potassium channel proteins, an alternation between variable and conserved regions is visible, the latter indicated by vertical bars.

One characteristic is shared by all potassium channels, irrespective of their origin. From a mixture of cations of almost similar size, they are able to selectively transport the potassium ion. The smallest common element is represented by the amino acid sequence glycine-tyrosine-glycine, a motif found in all potassium channels, including those of bacteria and even viruses.

Within this motif and the adjacent region, amino acids will be replaced to test their function in the guard cell potassium channel.

The replacement of a single amino acid residue, for example a charged by an uncharged or a hydrophobic by a hydrophilic, is called site-directed mutagenesis.

This is done at the level of the DNA. A plasmid carrying the potassium channel cDNA serves as a template in a polymerase chain reaction. The single molecular steps within a mutagenesis cycle are now demonstrated.

In the first step, the plasmid DNA is denatured, hydrogen bonds break, and the double-stranded DNA separates or melts. After the reaction temperature has been lowered, an oligonucleotide, which carries the desired mutation, attaches to the single-stranded DNA.

The free 3' end of the oligonucleotide serves as a primer for a DNA polymerase, which copies the circular DNA leading strand.

This increase in DNA chain length is called elongation. A plasmid hybrid consisting of mutated and non-mutated DNA evolves. The first mutagenesis cycle is completed.

The second cycle is initiated by melting of this hybrid.

During the second annealing process, another primer attaches to the mutated single strand. It fits perfectly. During the elongation process, the DNA polymerase again elongates the free 3' end of the primer.

The product of the second mutagenesis cycle is a mutated double-stranded potassium channel cDNA. This process is repeated many times to amplify the selectively mutated potassium channel cDNA.

It is transcribed in vitro into a copy cRNA and is ready to be used for a functional analysis.

To test the function of transport proteins, an expression system is used which itself possesses no or only a very few ion channels. In this respect, oocytes of the South African clawed toad Xenopus levis have become well established.

After the viability of the oocytes has been checked, they are prepared for injection with cRNA, encoding the mutated potassium channel gene. With the help of micromanipulators and an automatic injection system, a small capillary impales the oocyte and injects the cRNA.

In this way, a population of oocytes that express the mutated potassium channel gene is generated. In a second oocyte population, the unmodified potassium channel gene of the wild type is injected.

In the following analysis, a comparison between both populations is possible. Inside the cell, the potassium channel mRNA is recognized by ribosomes and translated into protein subunits.

They assemble and fuse with the plasma membrane of the oocyte. As early as one to two days after injection of the potassium channel cRNA, the electrical properties of the oocyte membrane are dominated by potassium channels.

Oocytes are placed in the cavity of a perfusion cuvette and exposed to a potassium solution. Using the two electrode voltage clamp technique, potassium currents flowing through many hundred thousand potassium channels expressed simultaneously are monitored.

For this purpose, a voltage and current electrode is carefully inserted into the oocyte. This is accomplished with micromanipulators. The voltage electrode on the left measures the membrane potential.

The current electrode on the right injects exactly as much current as is needed to maintain a given test potential. The potassium currents resulting from the changes in membrane potential are recorded by an amplifier and displayed. A computer controls the measurement and analysis of the data.

Now we follow the potassium current into the oocyte using the wild-type channels as an example. In response to stepwise changes in membrane potential to negative values, with the channel selected here, a time-dependent potassium current is elicited.

On replacement of potassium in the test solution by sodium, no currents are observed, even with strong polarization of the membrane to minus 160 millivolts.

This indicates that the potassium channel is impermeable to sodium ions. On repeating this experiment with different channel mutants, one finds that among them some mediate ionic currents even in a sodium solution.

The selectivity filter is, as it were, leaky and allows sodium to move through the channel in addition to potassium. This experiment shows that quite a restricted region of the protein forms the selectivity filter of the ion-conducting pore.

Now we will determine the three-dimensional structure of the channel protein by crystallization. Using microorganisms, which can be grown to a dense suspension in fermenters, the protein in question is enriched million-fold.

Here a membrane protein of the purple bacterium Rhodobacter spheroidis. In the first step, the bacteria have been sedimented in a flow-through centrifuge to separate them from the surrounding growth medium.

On the plastic membrane during centrifugation, a bacterial sediment several millimeters thick has formed. In the sediment, the density of the bacteria, and thus of the membrane protein to be isolated, is increased many-fold.

With a spatula, the cell concentrate is transferred to a beaker and weighed.

For the subsequent isolation and crystallization of the membrane protein, one needs several grams of sediment. The following steps take place in a cold room to prevent the extremely labile membrane proteins from disintegrating.

By chromatographic methods, they will now be separated from one another. After solubilizing the cell membrane in a detergent-containing buffer, the solution

contains the protein in question, as well as many other membrane proteins. In the last step, the highly enriched membrane protein is separated from other proteins by ion exchange chromatography. It appears as a dark band, and while moving through the column, shown here in time-lapse, separates from the other proteins, which are discarded.

The channel protein is eluted from the column and directed to a fraction collector.

Now the basic requirements for the subsequent crystallization of the protein are fulfilled. The protein concentrate is pipetted into special crystallization dishes.

Afterwards, at constant temperature, water is gradually removed from the protein suspension.

In this way, protein molecules come into close contact with one another and aggregate into protein crystals. A time-lapse sequence helps to illustrate this process, which takes several days.

The crystal consists of many millions of identical channel proteins. When the protein in the suspension is used up, the crystal growth stops. Crystals that are now 1 to 3 mm in diameter are prepared for the analysis of the crystal structure.

A selected crystal is drawn into a measuring capillary. The ends of the capillary are sealed with wax to protect the very fragile protein crystal from disintegration caused by air humidity.

The three-dimensional structure of the protein crystal is now determined by X-ray analysis.

For this purpose, the glass capillary containing the crystal is aligned with the X-ray source. The detector is positioned right next to it. Subsequently, the X-ray beam hits the crystal. In response, the protein crystal diffracts the X-ray beam.

A detector screen monitors the diffraction pattern. The crystal is now rotated stepwise around its axis. The X-rays, which are normally not visible, are illustrated here. For each angle, the diffraction patterns are computed in relation to each other.

Like the orthopedist, who identifies the exact position of bones and vertebrae from an X-ray image, the crystallographer calculates the position of every individual amino acid within the protein crystal from the diffraction pattern. In contrast to the doctor, however, the crystallographer is not satisfied with millimetre differences.

He has to reach a resolution of one to two angstrom, which is a ten-millionth of a millimetre, to solve the three-dimensional structure of a potassium channel. Here is the result of such an analysis on the potassium channel from the bacterium Streptomyces lividans.

As visible from the top view, the channel is made up of four identical subunits. In the centre of the channel complex is situated the iron-conducting pore, in which the potassium ions are located. After rotation through 90 degrees, it can be seen that these four subunits are made up of alpha helical stretches

of about 17 to 21 amino acids and anchor the membrane protein within the membrane. Following removal of the front and rear subunits, the potassium-selective region of the pore is visible. The constriction in the channel pore is formed by the three amino acids glycine, tyrosine, glycine,

the motif that all potassium channels have in common. This essential selectivity filter for potassium ions is stabilised by two pore helices.

The path of a potassium ion through the channel pore is exemplified on a simple model. The negative membrane potential draws a potassium ion into the pore where it binds to the selectivity filter

until a second potassium ion binds, which, due to electrostatic repulsion, causes the first ion to be released. Negative charges at the entrance to the pore prevent the entry of anions, here shown in silver. The selectivity for potassium in relation to other cations is the result of specific binding sites at the channel filter.

Due to repulsion amongst the potassium ions, fluxes of the order of 10 to the power of 7 ions per second are obtained. When mutations occur, ion channel function is impaired. Sometimes, the channels even lose their activity.

In plants, these genetic defects can result in malfunction of guard cells, growth defects in the shoot and root, and in humans, they are related to hereditary diseases like arrhythmia, epilepsy,

cystic fibrosis. Thus the structural analysis of iron channels is an essential prerequisite in, for example, the development of new approaches to therapy and the optimization of plant breeding.

The information for realization of these projects can be gained from protein crystals. The structural analysis, together with biophysical and molecular genetic analyses, will facilitate the step from molecule to application.