- Open Access
Protocol: optimised electrophyiological analysis of intact guard cells from Arabidopsis
© Chen et al.; licensee BioMed Central Ltd. 2012
- Received: 13 August 2011
- Accepted: 10 April 2012
- Published: 6 May 2012
Genetic resources available for Arabidopsis thaliana make this species particularly attractive as a model for molecular genetic studies of guard cell homeostasis, transport and signalling, but this facility is not matched by accessible tools for quantitative analysis of transport in the intact cell. We have developed a reliable set of procedures for voltage clamp analysis of guard cells from Arabidopsis leaves. These procedures greatly simplify electrophysiological recordings, extending the duration of measurements and scope for analysis of the predominant K+ and anion channels of intact stomatal guard cells to that achieved previously in work with Vicia and tobacco guard cells.
- K+ channel (voltage-gated)
- Cl- channel, Voltage-gated
- Membrane conductance
- Mutant analysis
Stomata are pores, commonly found in the epidermis of leaves, and are surrounded by a pair of specialised cells known as guard cells. Guard cells regulate the size of the stomatal pore to balance the exchange CO2 for photosynthesis with the need to conserve water . The acquisition of stomata and the leaf cuticle are considered to be key elements in the evolution of advanced terrestrial plants  allowing plants to inhabit different and often fluctuating environments while controlling water content. Stomatal pores typically occupy less than 5% of the leaf surface, but they provide for over 90% of the CO2 entering the leaf and over 70% of water loss from the plant as a whole . Guard cells respond to a number of well-defined signals – including hormones, light and atmospheric CO2 concentration – integrating these signals to regulate stomatal aperture [4, 5].
In the past few decades, the combination of physiological and molecular biological methods in the model plant Arabidopsis thaliana has greatly advanced our understanding of stomata [1, 4–7]. Among these, voltage clamp methods have proven powerful in connecting the molecular and physiological frameworks in an understanding of stomatal function. The voltage clamp itself lies at the core of a toolchest of techniques and provides the essential utility to bring the driving force of membrane voltage under experimental control. By so doing, it enables the dissection, identification and monitoring of ionic currents carried by individual ion transporters – ATP-dependent pumps, ion-coupled carriers and ion channels – across biological membranes . Classic voltage clamp methods rely on impalements with two microelectrodes (or a single microelectrode with two separate barrels) that are used to measure membrane voltage and to pass current for voltage clamping, respectively [8, 9]. Because a defined spatial geometry is essential for quantifying current spread under clamp conditions [8–10], these methods have proven highly successful for work primarily on a small number of single-celled species as well as cell types that are easily isolated from their surrounding tissues [11–17].
Since its wider introduction in the 1980's [18, 19], the patch clamp variant of the voltage clamp has been widely used in studies of plant ion channels [8, 20]. The patch clamp offers a number of advantages for work on plant cells, the most important being the facility for electrical recordings from single cells isolated from almost any surrounding tissue, thereby avoiding the difficulties of electrical coupling via plasmodesmata between cells in situ . It also presents some difficulties. For patch clamp recordings from plant cells it is essential to remove the cell wall, commonly by enzymatic digestion, and to stabilise the protoplast against osmotic swelling in the absence of turgor. Both manipulations affect the underlying homeostatic properties of the cells and must influence their physiological behaviour [22, 23]. Additionally, obtaining electrically and mechanically robust seals between the patch electrode and protoplast, and retaining stable measurements without significant “rundown” of currents over long periods of time are often challenging [20, 24].
By contrast with many plant cell types [but see Chen et al. ], guard cells at maturity do not retain electrical connections with their neighbours [11, 25]. They are easily separated by mechanical peeling of leaves  and recovered intact with their cell wall within the monolayer of epidermal cells. These features greatly simplify their handling for voltage clamp recordings and analysis, avoiding the need to isolate protoplasts and the technical challenges of the patch clamp. Despite the obvious advantages, only a very few studies [26–28] have made use of microelectrode impalements and classic voltage clamp methods with intact Arabidopsis guard cells. A major difficulty in this case has been to obtain reliable measurements over 20–30 min or more, time periods long enough for physiological and pharmacological studies with single cells. Thus, many researchers have relied on statistical approaches in patch recordings from populations of guard cell protoplasts, often without an internal reference for comparisons; simply put, impalement methods have not offered significant benefits in overcoming the problem of ‘rundown’ in channel activities common to patch clamp recording [20, 24].
We have revisited the problems of voltage clamp recording from intact Arabidopsis guard cells and offer here a few simple procedures that enable classic, two-electrode voltage clamp recordings. Included with this protocol are summaries of results demonstrating its utility in characterising the major ion channel currents and their stability over time periods of one hour or more. The impalement approach greatly simplifies experimental access to these currents and enables physiological studies to be carried out on a cell-by-cell basis.
Arabidopsis thaliana. For purposes of demonstration, we included with wild-type (Col0) the nitrate reductase-null mutant nia1-1/nia2-5 (nia1nia2) , the ABA-receptor quadruple mutant pyr1/pyl1/pyl2/pyl4 (QC3) , the vesicle-trafficking mutant syp121 (=syr1/pen1) and its complementation with SYP121[31, 32], the dehydroascorbate reductase mutant dhar1-3, and the K+ channel mutant kc1-2.
KCl, Ca(OH)2, NaOH, HCl, CsCl, tetraethylammonium chloride (TEA-Cl), potassium acetate (K+-Ac), and 2-(N-morpholino)ethanesulfonic acid (MES) analytical grade.
Opening Buffer (OB) for pretreating the stomatal guard cells, comprising 50 mM KCl and 10 mM MES, titrated to its pH 6.1 with NaOH, without added Ca2+.
Recording Buffer 1 (RB1) for voltage clamp measurements of K+ channel currents, comprising 10 mM KCl and 5 mM MES, titrated to pH 6.1 with Ca(OH)2 ([Ca2+] = 1 mM).
Recording Buffer 2 (RB2) for voltage clamp measurements of the Cl-/anion channel currents, comprising 15 mM TEA-Cl, 15 mM CsCl and 5 mM MES, titrated to pH 6.1 with Ca(OH)2([Ca2+] = 1 mM).
Environment-controlled growth room
Refrigerator for stratifying seeds at 4°C
Narashige PD5 multi-purpose microelectrode puller or equivalent, modified for multibarrelled microelectrodes .
Light microscope with a total magnification at least 400× or higher
12-volt battery for DC power to supply microscope
Huxley-type micromanipulator with carrier (see below) incorporating light-weight micropositioner (e.g. Narishige C2-type micromanipulator)
Gravity-feed system for switching between experimental solutions 
Fine-tipped forceps, dressing forceps and razor blades
Glass capillaries for double-barrelled microelectrodes 
Two-ml polypropylene pipettes, silicon rubber and 0.5-mm diameter Ag wire for halfcells (see  and below)
Key steps for growing Arabidopsis plants and selecting guard cells for voltage clamp
Pretreat compost with Intercept 70WG (Scotts, Ipswich, UK), a systemic insecticide.
Sow seeds onto the nutrient-rich Levington F2 + S 3 compost (Coulders, Glasgow, UK) in 60 mm pots covered with polyester mesh (Remnant Kings, Glasgow, UK Figure 1A) to avoid soil contact of the abaxial leaf surface and soil-borne stress factors.
Stratify seeds at 4°C, once sown, for 48 hours and leave the seed to germinate under a plastic lid (>95% RH) for one week.
Cultivate plants in a controlled environment growth room under long day conditions with 100 μmol m-2 s-1 light and a light/dark cycle of 16 h/8 h, 22/18°C, and 55/70% RH. Evenly and regularly water plants from below.
Transfer pots after one week to propagators. We use propagators with NITEX mesh fabric (mesh opening 200 μm diameter; Sefar, Heiden, Switzerland) over the sides of the covers to permit free air exchange while keeping out insects.
In preparation for experiments excise either the 5th or 6th true leaf of three-week-old plants; these leaves display an elliptical shape and are more serrated compared to the older leaves. NOTE: There is a correlation between stomatal responsiveness and stomatal age, the most responsive stomata often occur on leaves with higher stomatal densities, many stomatal primordia and smaller epidermal cells (Figure 1 B and C). Successful impalements yield similar currents under voltage clamp when recorded from guard cells of plants grown under long- and short-day conditions. Nonetheless, we favour plants grown under long days, as growth under short days gives lower stomatal densities (Figure 1 D).
Pretreat the glass of the measuring chamber, coating it with Dow-Corning silicon prosthetic adhesive (Factor II, Tucson, USA; see ). NOTE: Silicon adhesive is pressure-sensitive and optically clear. Once dried, it remains useable for many weeks, even under water. However, the solvent used in the adhesive must evaporate before use or it will kill the cells.
Excise the epidermis of the leaf by wrapping the leaf over a finger, adaxial side down, cut into the mesophyll near the base of the mid-vein with forceps, and lift the abaxial epidermis away from the mid vein towards the leave margin. Gently replace peel against the mesophyll, keeping a gentle tension to avoid folds, then cut at the end of the peel near the leaf margin using a fresh (sharp) razor blade. NOTE: It is often easier to peel away the epidermis some minutes after excision when the leaf is less turgid, and to work from the petiole to the apex of the leaf. Ideally, epidermal peels should be free from wrinkles, folds, dirt and, once mounted, air bubbles. Successful impalements are best obtained from open stomata with young guard cells (arrows, Figure 1 B), as judged by the thickness of the stomatal lip and squat shape of the guard cells.
Press the abaxial side of the leaf with the excised epidermal peel gently onto the prosthetic adhesive coating of the measuring chamber glass. Remove the remaining leaf tissue and cover the epidermal peel immediately with OB to prevent it drying.
Key steps for pulling microelectrodes
Pull microelectrodes to give tip resistances of 300–500 MΩ when filled with 200 mM K+-Ac.
For double-barrelled microelectrodes with the higher input resistances (and correspondingly lower electrolyte leakage rates), pull double-barrelled microelectrodes, after twisting 360o , using settings to give a pull time around 25 s. NOTE: We use settings similar to those used for Vicia and tobacco guard cells  , but with the coil heat elevated to give pull times roughly 25% less than used for Vicia guard cells. The resulting microelectrodes have 1.8-2.0 cm-long shanks and tips that tapered with a 1–1.5 o angle (Figure 2 A).
Key steps for impaling Arabidopsis guard cells
Electrical recordings using double-barrelled microelectrodes are carried out largely as described previously [12, 35] with some modifications. For K+ currents, microelectrode barrels are filled with 200 mM K+-Acetate, pH 7.5, to minimise interference from the anion current and recordings are carried out in continuously-flowing RB1; for measurements of anion current, both electrode barrels are filled with 200 mM CsCl and the cells bathed in flowing RB2. Currents recorded under voltage clamp are normalised to the surface area of the impaled guard cells and, for K+ channel analysis, are corrected for background (instantaneous) currents as described previously [12, 35] using Henry’s EP suite software (Y-Science, University of Glasgow, UK). NOTE: The typical length and radius of Arabidopsis guard cells are 20 and 5 μm, respectively. For the data summarised in the Tables, these parameters were 22 ± 0.6 μm 2 and 4 ± 0.1 μm, respectively. Assuming a spheroid geometry, the mean guard cell surface area and volume were 468 ± 12 μm 2 and 783 ± 21 μm 3 , respectively.
Carry out impalement by first positioning the microelectrode to rest over one guard cell and present the tip across the stomatal pore before advancing along the axis of the microelectrode to impale the second guard cell. NOTE: The initial movement of the microelectrode towards the guard cell requires very gentle manipulation. A ‘snapping’ of the tip through the cell wall and into the guard cell should occur together with an increase in input resistance to approximately 1 GΩ and decrease (more negative) in membrane potential (see Additional file 1: Table S1).
Wait 2–3 min for a seal to stabilize after impalement. NOTE: As with Vicia guard cells  , successful impalements show an increase in input resistance and membrane voltage over 2–3 minutes. Impalements carried out in RB1 buffer, but with 0.1 mM KCl, will give much larger changes in voltage as the microelectrode seals into the cell. For purposes of the comparisons below, we allowed recordings to stabilise under free-running conditions for 10 minutes before collecting voltage clamp data.
Switch to the RB1 or RB2 for K+ and anion currents measurements, respectively, using a gravity-fed system.
Three key factors are essential for successful, two-electrode, voltage clamp recordings with Arabidopsis guard cells. First, the preparation and handling of the plants is important, incorporating a pretreatment regime with a stomatal opening buffer prior to the start of experiments; second, microelectrode design must meet the demands for impalements of very small cells, notably in the use of fine tips with input resistances roughly 5-fold higher than typically used for Vicia and tobacco guard cells; finally, a modified clamp and brace to carry the amplifier headstages and construction of light-weight, but rigid halfcells are essential prerequisites to provide stability without mechanical relaxation for long-term recordings. Overall, this combination of factors is sufficient to achieve measurements comparable to those with the much larger guard cells of Vicia and tobacco. These methods should now greatly speed the analysis of many mutants of Arabidopsis by simplifying electrophysiological studies of the guard cells.
We thank Amparo Ruiz-Prado for growth room support. This work was funded by grants BB/F001673/1, BB/F001630/1 and BB/I024496/1 to MRB from the UK Biotechnology and Biological Sciences Research Council. CE was supported initially by a Glasgow University Open PhD Scholarship and by a grant from Plant Biosciences Ltd, Norwich. ZHC was supported additionally by a 2011 UWS Research Travel Fellowship. Seed of the nia1nia2 and pyr1/pyl1/pyr2/pyl4 mutants were gifts from Prof. S. Neill in University of the West England and Prof. S. Cutler of University of California in Riverside, respectively.
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