- © 2007 American Society of Plant Biologists
Abstract
Sieve element (SE) protoplasts were liberated by exposing excised phloem strands of Vicia faba to cell wall-degrading enzyme mixtures. Two types of SE protoplasts were found: simple protoplasts with forisome inclusions and composite twin protoplasts—two protoplasts intermitted by a sieve plate—of which one protoplast often includes a forisome. Forisomes are giant protein inclusions of SEs in Fabaceae. Membrane integrity of SE protoplasts was tested by application of CFDA, which was sequestered in the form of carboxyfluorescein. Further evidence for membrane intactness was provided by swelling of SE protoplasts and forisome dispersion in reaction to abrupt lowering of medium osmolarity. The absence of cell wall remnants as demonstrated by negative Calcofluor White staining allowed patch-clamp studies. At negative membrane voltages, the current-voltage relations of the SE protoplasts were dominated by a weak inward-rectifying potassium channel that was active at physiological membrane voltages of the SE plasma membrane. This channel had electrical properties that are reminiscent of those of the AKT2/3 channel family, localized in phloem cells of Arabidopsis (Arabidopsis thaliana). All in all, SE protoplasts promise to be a powerful tool in studying the membrane biology of SEs with inherent implications for the understanding of long-distance transport and signaling.
Use of fungal enzymes that degrade plant cell walls enables the isolation of plant cell protoplasts, which have become an invaluable tool in plant biology. For example, protoplasts have yielded considerable insight into plasma membrane-bound ion channels and carbohydrate carriers in a variety of plant cells ranging from large parenchyma cells to tiny guard cells.
Due to technical barriers, sieve elements (SEs) are missing from other cell types that have been protoplasted successfully. A major problem is the tuning of the digesting mixture; the conventional enzyme mixes turn phloem tissues into a mash. Another problem is the unequivocal identification of SE protoplasts. They easily fragment into smaller protoplasts during isolation and, therefore, can hardly be distinguished from those of other, smaller cell types.
Nevertheless, it remains tempting to isolate and identify SE protoplasts for several reasons. The SE plasma membrane contains numerous ion channels and carbohydrate carriers that are essential for sieve tube functioning (e.g. Patrick et al., 2001; van Bel, 2003). Ion channels are not only meaningful for the ion household of SEs, but also for the regulation of photoassimilate transport rates through sieve tubes (Fromm and Bauer, 1994; Ache et al., 2001; Deeken et al., 2002; van Bel and Hafke, 2005). They also play a central role in long-distance signaling, such as the propagation of electrical signals via the phloem (Fromm, 1991; Rhodes et al., 1996; Lautner et al., 2005; Furch et al., 2007).
The properties of phloem-associated potassium channels have been determined by heterologous expression in Xenopus oocytes (Marten et al., 1999; Bauer et al., 2000; Lacombe et al., 2000; Ache et al., 2001; Deeken et al., 2002; Phillipar et al., 2003). These potassium ion channels have been located in the phloem by in situ hybridization techniques (Marten et al., 1999; Lacombe et al., 2000; Ache et al., 2001). However, the exact cellular localization, ion channel densities, ion channel types, and distribution along the phloem path are unknown. SE protoplasts isolated from the respective phloem sections would provide a unique tool for unequivocal information about these issues. The same applies to calcium channels, which have been postulated to occur in the SE plasma membrane (Volk and Franceschi, 2000). SE protoplasts from successive phloem sections would also enable to identify, characterize, and quantify carbohydrate carriers in the SE plasma membrane at various sites along the phloem translocation pathway. Differential deployment of sugar carriers is likely essential for carbohydrate allocation in intact plants (e.g. Patrick et al., 2001; Kühn, 2003; Hafke et al., 2005).
Isolation of SE protoplasts may also allow the study of membrane biophysics. The mass flow through the pressurized sieve tubes makes high demands on the physical properties of the SE plasma membrane. Knowledge of physical properties like elasticity and fluidity and their impact on the activity of transmembrane proteins such as mechanosensitive channels is necessary for a better understanding of the pressure regulation of phloem transport.
For isolation of SE protoplasts, we employed Vicia faba phloem since SEs in this species contain giant calcium-sensitive protein bodies (forisomes; Knoblauch et al., 2001, 2003) meant to act as tools for SE identification. Here, we present a method for preparation and identification of functional SE protoplasts of V. faba. The integrity of the SE protoplasts was tested by use of fluorochromes and osmotic treatments. Furthermore, patch-clamp experiments were carried out to investigate the functionality of SE protoplasts.
RESULTS AND DISCUSSION
Isolation and Identification of V. faba SE Protoplasts
Following an incubation period of 10 h in the enzymatic digestion medium, formation of phloem protoplasts released from disintegrating V. faba phloem strands was observed using light microscopy and confocal laser scanning microscopy (CLSM). The enzymatic treatment liberates protoplasts of various phloem cell types, which are then tracked and identified by screening the phloem strands under light microscopy. Phloem protoplasts are always produced in low but sufficient numbers for patch-clamp studies, or general biophysical and physiological studies (see section on functionality of SE protoplasts below).
Phloem protoplasts are highly variable in shape, size, and structure. Two types of SE protoplasts were found. Simple SE protoplasts arise from more central parts of disintegrating SEs and are identified by inclusion of a large protein body named the forisome (Fig. 1A ). Forisomes are responsible for sieve plate occlusion due to a turgor- and damage-induced, calcium-dependent conformation change in intact SEs of legumes (Knoblauch et al., 2001, 2003). Thus, protoplasts containing a forisome must inevitably be SE protoplasts.
A to D, Isolation and identification of V. faba SE protoplasts. A, Light-microscopic image of a simple SE protoplast. Simple SE protoplasts can readily be identified by inclusion of a forisome (asterisk), which is typical of fabaceaen SEs. B, An emerging composite SE protoplast (twin protoplast), partially detached from the phloem strand, composed of two small cylindrical protoplast precursors intermitted by a sieve plate (arrow). Both protoplasts enclose a forisome (white asterisks) and tiny P-plastids (arrowheads) near the protoplast membrane. Next to the SE protoplast, a large parenchyma cell protoplast (PPCP) is visible. C, An isolated CCP adhered to a SE during isolation. The forisome of the adjacent intact SE is marked with an asterisk. D, An enzymatically isolated one-layer phloem strand. A CCP is adhered to a collapsed SE (sieve plate marked with an arrow). At the right, the precursor of a large vacuolar parenchyma cell (PPC) is visible. E and F, Membrane integrity of V. faba SE protoplasts. E, CLSM image of a composite SE protoplast loaded with CFDA-AM ester. The twin protoplast consists of a large (large arrowhead) and a small protoplast (small arrowhead) separated by a sieve plate (arrow). Both protoplasts have accumulated fluorescent CF arising from the esterase-mediated cleavage of CFDA-AM. CF is not removed by washing, indicative of an intact membrane system. F, CLSM image of a twin protoplast adhered to a phloem strand. The protoplast was stained using the membrane-soluble fluorescent dye RH-414. Staining shows two protoplasts enclosed by a plasma membrane intermitted by a sieve plate (arrow) in which each sieve pore is lined by a plasma membrane (orange striping). The larger protoplast (large arrowhead) contains a forisome (light-transmission picture not shown). G to J, Mechanism of SE protoplast formation. G, Formation of a SE protoplast precursor in disintegrating phloem tissue. Due to distal collapse of the SE plasma membranes at either side of the sieve plate (arrow), two adjoining SE protoplasts emerge. Note the formation of a membranous tail (arrowhead), composed of appending membranes. Forisomes are marked with asterisks. H, Formation of a simple SE protoplast as a result of constriction from the sieve plate (arrow). Membranous tails (arrowheads) occur at both sides of the SE protoplast precursor. I, CLSM picture of an emerging SE twin protoplast loaded with CFDA. At this stage of formation, the protoplast is already sealed as demonstrated by the accumulation of CF. The large protoplast encloses a forisome (asterisk); the sieve plate is marked with an arrow. J, Transmission picture of I. Clearly visible are SE plastids (arrowheads) in the small protoplast precursor. The protoplast is being formed at a branching point of a sieve tube with two sieve plates. The lower part of the sieve plate separates two large SE protoplast precursors. The small upper sieve plate part may give access to a third tiny flat protoplast sealed with a SE plasma membrane (small arrows). The pictures were taken after 3 h of enzyme incubation with the exception of A and H (taken after 10 h of enzyme incubation).
In addition, composite SE protoplasts, which arise from the sieve plate area (the joint between the SEs), showed up. Hence, composite SE protoplasts include a sieve plate, which provides an unequivocal means of identification (Fig. 1, B, E, and F). Composite protoplasts possess a protoplast at either side of the sieve plate (“twin protoplasts”; Fig. 1, B and E). One of them usually includes a forisome. SE protoplasts with forisomes in both compartments were seldom observed (Fig. 1B).
Companion cell protoplasts (CCPs) were often found adhered to SEs (Fig. 1, C and D). CCPs typically contain chloroplasts aggregated at one side; the cytoplasmic compartment occupies 20% to 25% of the total protoplast volume. A quick calculation shows that the spindle-shaped CCs with a diameter of 3 to 5 μm and a length of 200 to 300 μm would indeed produce spherical CCPs with a diameter of 10 to 20 μm.
In contrast to CCPs, which have a diameter of 10 to 20 μm, prosenchymatic phloem parenchyma cells form large protoplasts with an average diameter of 40 to 60 μm. They can readily be distinguished from CCPs by the equal distribution of the cytoplasm at the margin, matching about 1% of the total protoplast volume (Fig. 1B).
Membrane Integrity of V. faba SE Protoplasts
As a test for membrane integrity, SE protoplasts were loaded with the colorless CFDA-AM ester as described for intact phloem tissue (e.g. Knoblauch and van Bel, 1998). During incubation, the ester was cleaved by intracellular SE esterases (Oparka and Read, 1994) into the membrane-impermeant and fluorescent form carboxyfluorescein (CF). Containment of CF inside the protoplasts after washing demonstrates the integrity of the SE protoplast membrane (Fig. 1E).
Information on the intactness of the connection between both protoplasts in a twin protoplast was obtained by labeling SE protoplasts with RH-414, a membrane-soluble fluorescent probe (Fig. 1F). The CLSM pictures of RH-414 staining show labeling of a membrane system enveloping the large forisome-containing and the small protoplast of SE twin protoplasts. Continuing fluorescent striping in the sieve plate area indicates that the extensive plasma membrane system lining the sieve pores had remained intact during the isolation procedure (Fig. 1F).
Mechanism of SE Protoplast Formation
The mechanism of SE protoplast formation is not yet understood in detail. Precursors of the composite SE twin protoplast emerge near the sieve plates (Fig. 1, B and G). At either side of a sieve plate, the SE membrane collapses in such a way that tapering ends of SE membranes, situated at both sides of the sieve plate, seem to coalesce and form filamentous plasma membrane tail ends (Fig. 1G). As a result, a longish membrane compartment appears at either side of the sieve plate (Fig. 1, G and J). At this stage, the twin protoplast precursor is already sealed as evidenced by CF accumulation in both protoplasts (Fig. 1I). After constriction of the membranous tails, the composite SE protoplast starts rounding off before it gradually detaches from the phloem strand.
Formation of simple SE protoplasts (Fig. 1A) also depends on membrane tail formation (Fig. 1H). Thus, both simple and composite SE protoplasts rely on coalescence and constriction of membranous tails as a crucial step in protoplast formation. The amalgamation mechanism is obscure but deserves further (electron microscopic) studies.
Yield of V. faba SE Protoplasts and Formation Mechanism
The minute yield of SE protoplasts is regarded to be the aggregate result of various bottlenecks in SE protoplast preparation as outlined here.
1. The tight packing of the phloem tissue.
In contrast to the loose packing of parenchymatous tissues, the tight packing of the phloem tissue impedes a quick and uniform diffusion of the enzyme mixture.
2. The sensitivity of phloem tissue to wounding.
In comparison to other cell types, the SE/CC complex is very sensitive to the slightest injury. Phloem slicing induces massive wound effects and turgor changes and triggers a physiological and structural collapse of most SEs in a tissue slice. Thus, the preparation method requires the use of thick phloem strands to have a few intact SEs and imposes unavoidable artifacts that minimize SE protoplast yield.
3. Composition of the enzyme mixture.
The composition of the enzyme mixture is a critical factor for the success of SE protoplast formation in any plant species. Extensive concentration tests (not presented here) evidenced that only enzyme mixtures in a narrow concentration window lead to successful cell wall degradation and a complete detachment of SE protoplasts. Higher concentrations turn the phloem tissue into mash; lower ones only liberate parenchyma protoplasts.
4. Composition of the SE cell wall.
SEs of dicotyledonous plants develop cell walls thicker than those of adjacent parenchyma cell walls. In several species, SE cell walls consist of two morphologically distinct layers, a relatively thin outer layer and a thicker inner layer, the nacreous layer with a pearly appearance (Evert, 1990). The complexity of the SE cell wall is a potential ground for cumbersome enzymatic digestion.
5. Constriction of the membranous tails.
Transformation of parenchyma cells into protoplasts “only” demands the removal of cell wall material and the breakage of plasmodesmata. The production of SE protoplasts is more complicated in that the large SEs fragment during the isolation process, which requires considerable membrane reconstitution. Presumably, the SE plasma membrane is collapsing during the enzyme treatment. In a few cases, the membrane constricts and coalesces at one side (composite protoplasts) or at both sides (single protoplasts) of the emerging protoplast body. Coalescence of free membrane ends to a closed tail (Fig. 1, G and H) is a critical step toward formation of viable SE protoplasts. In most cases, a mismatch between the membrane ends or an incomplete coalescence of membrane tails prevents the final formation of SE protoplasts. It should be noted that the emergence of a composite protoplast depends on successful sealing at either side of the sieve plate. In protoplasts with a sieve plate, the creation of the twin configuration is necessary since otherwise the protoplast is not sealed at both sides.
Given the numerous handicaps, it may be some time before one can expect to gain high yields of SE protoplasts.
Functionality of SE Protoplasts; Activity of Calcium Channels
Indicative of membrane integrity of SE protoplasts is their reaction to osmotic shocks. SE protoplasts were shocked osmotically by an abrupt change from 600 to 50 mol m−3 mannitol in the external medium by microperfusion, while the external calcium concentration was maintained constant at 1 mol m−3 (Fig. 2, A and B ). The sudden decline in the external osmolarity induced a gradual swelling (Fig. 2, B–D) within 30 to 120 s. Further protoplast swelling resulted in a burst (not shown) of the SE protoplast. The forisome inside SE protoplasts dispersed instantaneously in response to the osmotic shock (Fig. 2, A–D) in keeping with the forisome behavior in intact SEs (Knoblauch et al., 2001). The calcium-dependent forisome dispersion (Knoblauch et al., 2003) is ascribed to calcium influx due to activation of mechanosensitive calcium channels.
Dispersion of a forisome in an intact SE protoplast of V. faba in response to a hypo-osmotic shock. A, The composite SE protoplast, stored in a medium containing 600 mol m−3 mannitol and 1 mol m−3 CaCl2, is composed of a larger protoplast (large arrowhead) and a smaller protoplast (small arrowhead, dotted outline) separated by a sieve plate (arrow). The outline of the forisome is delineated by a dotted line, those of the protoplasts by a dashed line. B, After a rapid bath perfusion with a solution containing 50 mol m−3 mannitol and 1 mol m−3 CaCl2, the protoplast swells. C and D, The ready dispersion of the forisome within seconds indicates calcium influx into the protoplast. E to H, A simple SE protoplast, including a forisome (asterisk), in the standard medium containing 600 mol m−3 mannitol and 1 mol m−3 CaCl2 was osmotically shocked by a solution containing 50 mol m−3 mannitol and 4 mol m−3 EGTA (calcium chelator). The calcium-dependent forisome fails to disperse in the absence of free Ca2+ ions. Following medium change, the protoplast swells (E and F) and collapses after exceeding a critical expansion level within a few seconds (G and H). Note that the oval form of the SE protoplast is dictated by the shape of the forisome.
The same osmotic shock (600–50 mol m−3) in a calcium-free medium containing 4 mol m−3 of the calcium chelator EGTA only induced protoplast swelling without forisome dispersion (Fig. 2, E and F). The expansion resulted in rupture and collapse of the SE protoplasts (Fig. 2, G and H), while the forisome stayed in the condensed conformation due to the absence of calcium. Thus, the forisome dispersion in 1 mol m−3 Ca2+ suggests (Fig. 2, A–D) the presence of functional mechanosensitive calcium channels in the SE plasma membrane.
The functionality of SE protoplasts was further demonstrated by applying suction to the SE plasma membrane via a microcapillary connected to a pressure device (Fig. 3, A–D ). In reaction to suction, the forisome dispersed presumably as a result of Ca2+ influx through mechanosensitive channels (Fig. 3, A–D). Forisome dispersion failed to occur (Fig. 3, E–H) in the presence of the Ca2+ channel blocker Gd3+ (2 mol m−3). These results (Fig. 3, A–H) again indicate functional mechanosensitive calcium channels in the SE plasma membrane.
Expansion of a forisome in an intact SE protoplast of V. faba in response to suction using a microcapillary connected to a pressure device. A, A simple oval SE protoplast including a forisome in a medium containing 600 mol m−3 mannitol and 1 mol m−3 CaCl2. A microcapillary (arrow) filled with bathing medium and connected to a pressure device is visible in the vicinity of the protoplast. The outline of the forisome is delineated by a dashed line. The oval form of the SE protoplast is dictated by the shape of the forisome. B and C, In response to suction exerted on the plasma membrane of the SE protoplast, the forisome starts to disperse at both ends (marked with arrowheads). As shown in C, the oval form of the protoplast disappears before the midsection of the forisome has dispersed completely. D, The full dispersal is accompanied by the rounding-off of the SE protoplast. E to H, A simple SE protoplast including a forisome (asterisk) stored in the standard medium containing 600 mol m−3 mannitol, 1 mol m−3 CaCl2, and 2 mol m−3 GdCl3 (calcium channel blocker; E) was attached to a microcapillary (arrow) containing the bath solution (F). Strong suction exerted on the protoplast (G and H) demonstrates the absence of the calcium-dependent forisome dispersion, presumably due to inhibition of Ca2+ influx by blocking the mechanosensitive calcium channels localized in the SE plasma membrane.
Detection of Cell Wall Remnants by Calcofluor White Staining
To test the suitability for patch-clamp studies, SE protoplasts were tested on cellulosic and callosic wall remnants by Calcofluor White (CW) staining (Choi and O'Day, 1984; Nakamura et al., 1984). As control experiments, intact phloem tissue (Fig. 4A ) and a disintegrating phloem strand (Fig. 4B) were stained with CW. In intact phloem tissue, cell walls, sieve plates, as well as the pore plasmodesmata units were stained intensively (Fig. 4A). The CW staining gradually disappeared with incubation time in the digesting medium (Fig. 4B), indicative of dissolution of the cell wall. After an incubation time of 10 h, several SE protoplasts solely showed CW staining of the sieve plate (Fig. 4, C and D).
Detection of cell wall cellulose and callose using CW staining on intact tissue, disintegrating phloem tissue, and isolated SE protoplasts. A, CW staining of an intact SE in the main vein of a V. faba leaf. The sieve plate (arrow), the pore plasmodesmata units (arrowheads), as well as the cell wall of the SE show distinct staining. B, CW staining of a disintegrating phloem strand. Cell wall staining is less intent than under A. The sieve plates are marked with arrows. C, Solely the sieve plate (arrow) shows CW staining in an intact twin protoplast. Cell wall staining has disappeared completely. D, Transmission picture of C. In one protoplast of the twin protoplast, a dispersed forisome is visible (asterisk). The sieve plate is marked with an arrow.
Functionality of SE Protoplasts; Membrane Currents across the Plasma Membrane
Simple patch-clamp measurements were executed using SE protoplasts to merely test whether their membranes were functional. Under prevailing conditions, inward and outward currents were observed.
Inward Currents
At negative membrane voltages, instantaneous and time-dependent currents were observed in response to a series of test voltages between −172 and +53 mV. The corresponding steady-state current-voltage relationship recorded in asymmetrical potassium-gluconate solutions showed that the currents only weakly rectified at membrane voltages between +53 mV and −172 mV (Fig. 5, B and D ). The I-V plot of the steady-state currents (Fig. 5B) revealed a reversal voltage at −43 ± 2 mV (n = 4), close to the predicted equilibrium voltage for a 10-fold K+ gradient (EK+ = −58 mV) across the membrane, but was different from that of Mg2+ (−9 mV), Cl− (0 mV), or gluconate (+58 mV). Since the resting potential of the SE plasma membrane in V. faba is around −130 mV (Hafke et al., 2005) and thus more negative than EK+, the observed channel may contribute to K+ loading/retrieval into the SE.
Membrane currents across the plasma membrane measured in the whole-cell configuration. A, Current responses to voltage steps from the holding potential of −22 mV to a series of test voltages of 3.5-s duration in steps of 25 mV between −172 mV and +53 mV recorded in standard bath (6 mol m−3 potassium-gluconate, pH 5.5) and pipette (60 mol m−3 potassium-gluconate, pH 7.5) solutions. B, Current-voltage (I-V) relationship of the steady-state current densities from A. Data points were fitted with a polynomial function of the third order. The black arrow marks the Nernst potential for K+ (EK+). C, Time- and voltage-dependent as well as instantaneous outward currents elicited by applying depolarizing voltages in the whole-cell configuration. Current responses to voltage steps from the holding potential (−22 mV) to a series of test voltages of 3.5-s duration in steps of 25 mV between −172 mV and +153 mV recorded in standard solutions as mentioned in A. D, Current-voltage (I-V) relationship of the steady-state currents (Iss) from C. The black arrow marks the Nernst potential for K+ (EK+). E, Tail-current analysis of the time-dependent outward currents. Deactivation current densities in response to a double-pulse protocol starting from a holding potential of −72 mV to a prepulse voltage of +153 mV for 5 s. Tail currents were obtained by following current deactivation at test voltages between −172 mV and +53 mV. Erev, Reversal potential. F, I-V plot of tail currents from E. Values were calculated as the difference between the instantaneous and the stationary currents 1 s after stepping to each deactivating voltage. The predicted equilibrium voltage (EK+) is indicated for K+ with an arrow.
The observed weak rectifying potassium-selective channel active at negative membrane voltages shares electric properties with the AKT2/3 K+-channel family found in Arabidopsis (Arabidopsis thaliana) phloem cells (Marten et al., 1999; Lacombe et al., 2000; Deeken et al., 2002). This channel family may be involved in K+ transport accompanying phloem loading (Deeken et al., 2000, 2002) and unloading processes (Lacombe et al., 2000), control of membrane potential (Marten et al., 1999; van Bel and Hafke, 2005), and reestablishment of the membrane potential that depolarizes during a phloem-propagating action potential (Marten et al., 1999).
Outward Currents
Clamping the SE plasma membrane from a holding potential of −22 mV to test voltages between −172 mV and +153 mV results in activation of time-dependent outward currents (Fig. 5, C and D) positive to +70 mV. In addition, instantaneous currents were observed. For tail-current analysis (Fig. 5E), the plasma membrane of the SE protoplast was clamped from the holding voltage of −78 mV to a conditioning voltage of +153 mV to activate the time-dependent component. In a subsequent step, the plasma membrane was clamped to a series of test voltages between −172 mV and +53 mV. During the test pulse, the macroscopic tail currents gradually deactivated (Fig. 5E). A plot of tail-current amplitude revealed a reversal voltage of −50 mV (Fig. 5F), close to the equilibrium voltage for K+ (−58 mV). Under prevailing artificial conditions with potassium-gluconate on both sides of the membrane, this channel is not active at physiological membrane voltages.
SE Protoplasts from Other Plant Species
Modifications of the preparative steps with respect to enzyme concentrations, preparation temperatures, and incubation times yielded SE protoplasts from Nicotiana tabacum (Fig. 6, A and B ) and SE protoplast precursors from Cucurbita pepo (Fig. 6C). These SE protoplasts were composed of two protoplasts intermitted by a sieve plate. After countless tests, typical isodiametric round protoplasts separated from the sieve plate have only been obtained for Nicotiana. Despite a broad range of digestion conditions tested with respect to enzyme composition, osmolarity, duration of incubation, and temperature of the enzyme mixture, solely the longish precursor of Cucurbita SE protoplasts was produced. This form is ascribed to cell wall remnants residing on the plasma membrane. In view of additional experience acquired with other species, preparation of SE protoplasts seems to be highly species specific and the yield will probably always remain low.
SE protoplasts from other plant species. A and B, Composite SE protoplasts from N. tabacum as a twin protoplast precursor with two longish compartments (A) and in a nearly round configuration as known for V. faba protoplasts (B). Note that N. tabacum protoplasts contain numerous SE plastids. C, Longish precursor of a composite SE protoplast of C. pepo; the longish shape hints at the presence of cell wall remnants. A to C, Sieve plates are indicated by arrows.
Outlook
SE protoplasts are a promising tool for studying phloem biophysics. In the near future, V. faba SE protoplasts may be adopted as a model system for transporter deployment in the SE plasma membrane in view of the relatively easy mode of isolation and the results obtained with intact Vicia plants by other groups.
MATERIALS AND METHODS
Plant Material
Vicia faba ‘Witkiem’, Cucurbita pepo ‘Gelber Zentner’, and Nicotiana tabacum plants were grown in pots in a greenhouse at temperatures varying between 20°C and 30°C at 60% to 70% humidity and a 14-h/10-h light/dark period with supplementary lamp light (model SONT Agro 400 W; Phillips). The irradiance level was 200 to 250 μmol m−2 s−1 at the plant apex. Plants were all taken in the vegetative period just before flowering.
Protoplast Isolation
Internodes were excised from 3- to 4-week-old V. faba plants. Then, tangential cuts were made to split the internodes. For coarse mechanical isolation of stem phloem strands, tangential tissue sheets with a thickness of approximately 300 μm were sliced with a razor blade from the fracture face of the split internode. After preincubation of the slices for 15 min in a standard medium (WM) containing 600 mol m−3 mannitol, 1 mol m−3 dl-dithiotreitol (DTT), and 25 mol m−3 MES/NaOH, pH 5.7, the tissue was transferred into an enzyme mixture containing 400 mol m−3 mannitol, 100 mol m−3 KCl, 5 mol m−3 MgCl2, 1 mol m−3 DTT, 0.2% (w/v) polyvinylpyrrolidone-25, 0.5% (w/v) bovine serum albumin, 0.5% (w/v) cellulase ‘Onuzuka’ RS (Yakult Honsha), 0.03% (w/v) pectolyase Y-23 (Seishin), and 25 mol m−3 MES/NaOH, pH 5.7 (compare with Hafke et al., 2003).
After incubation for 10 h at 28°C, disintegrating phloem strands were filtered through a 80-μm nylon mesh and washed two times with the appropriate experimentation solution. For patch-clamp experiments, protoplasts were washed with standard bath solution and collected by centrifugation (Pico Fuge; Stratagene) twice.
The mechanism of protoplast formation and detachment were observed under microscopic surveillance (Leica DM-LB, fluorescence microscope, equipped with a special water immersion objective, HCX APO L40×/0.80 W U-V-I objective; Leica). The protoplasts were transferred into a small volume of WM in a bathing chamber equipped with a microperfusion system. Here, SE protoplasts were treated with various solutions and permanent microscopic surveillance.
Light micrographs were taken with a digital camera (Canon Power Shot S40) connected to a computer (Canon Digital Camera Solution Disk v8.0 software package).
SE protoplasts of N. tabacum and C. pepo were isolated as described for V. faba with slight modifications of the enzyme mixture, incubation time, and isolation temperature. SE protoplasts of N. tabacum were isolated over a period of 4 h and a temperature of 31°C in the above-mentioned enzyme mixture containing 0.55% cellulase and 0.035% pectolyase. SE precursors of Cucurbita were isolated over a period of 14 h and a temperature of 4°C in the above-mentioned enzyme mixture containing 0.6% cellulase and 0.04% pectolyase.
Staining of Protoplasts with CFDA, RH-414, and CW
CFDA
To test their membrane integrity, SE protoplasts were loaded with CFDA-AM ester (Molecular Probes) as described for intact phloem tissue (Knoblauch and van Bel, 1998). The solution was prepared from a CFDA-AM ester stock solution dissolved in WM to give a final concentration of 2.1 μm CFDA. After application of the CFDA-AM, protoplasts were incubated for 45 min at room temperature. During this period, the ester was cleaved by endogenous SE esterases into the membrane-impermeant and fluorescent form CF (Oparka and Read, 1994). Following thorough washing with WM, protoplast fluorescence was examined using CLSM (Leica TCS 4D) with a Krypton-Argon laser (Omnichrome) at 488 nm as described before (Knoblauch and van Bel, 1998).
RH-414
The protoplast plasma membrane was stained using the membrane-soluble fluorochrome RH-414 (Molecular Probes). RH-414 was diluted from a stock solution in WM to give a final concentration of 4.3 μm in a manner described before for the membrane-soluble fluorochrome RH-160 (Knoblauch and van Bel, 1998). Protoplasts were incubated for 5 min in RH-414 before washing with WM and scanning with CLSM (excitation 564 nm).
CW Staining for Detection of Cellulose and Callose
Isolated protoplasts or intact tissues were stained with 0.1% (w/v) CW (Choi and O'Day, 1984) dissolved in 400 mm mannitol for 15 min. After thorough washing with WM, cells were observed under an epifluorescence microscope (Leica DMLB) using a BP 340-380 excitation filter and an LP 425 barrier filter combination (Leica Microsystems).
Osmotic Experiments
SE protoplasts were bathed in a hyperosmotic solution containing 600 mol m−3 mannitol, 1 mol m−3 DTT, 1 mol m−3 CaCl2, and 25 mol m−3 MES/NaOH, pH 5.7. An abrupt bath change to a hypo-osmotic medium containing 50 mol m−3 mannitol, 1 mol m−3 DTT, 1 mol m−3 CaCl2, and 25 mol m−3 MES/NaOH, pH 5.7, by a homemade microperfusion system imposed an osmotic shock. As a control experiment for SE protoplast swelling in a calcium-free solution, protoplasts were preincubated in the hyperosmotic solution and osmotically shocked with a solution containing 50 mol m−3 mannitol, 1 mol m−3 DTT, 4 mol m−3 EGTA, and 25 mol m−3 MES/NaOH, pH 5.7.
Gd3+ Experiments
SE protoplasts were bathed in the hyperosmotic standard solution (see above) containing 600 mol m−3 mannitol, 1 mol m−3 DTT, 1 mol m−3 CaCl2, and 25 mol m−3 MES/NaOH, pH 5.7. Mechanical stress (suction) was exerted on SE protoplasts via patch-clamp microcapillaries connected to a pressure device (Cell Tram Oil microinjector; Eppendorf).
A microcapillary filled with the respective bathing medium was maneuvered to the protoplast by means of an LN SM-1 micromanipulator (Luigs & Neumann). Contact between protoplast and microcapillary was made by suction with the aid of a Cell Tram Oil microinjector (Eppendorf). As a control, protoplasts were incubated in the above-mentioned hyperosmotic solution supplied with 2 mol m−3 of the calcium channel blocker Gd3+ (as GdCl3). Protoplasts were observed using an epifluorescence microscope (Leica DM-LB, fluorescence microscope, equipped with a special water immersion objective, HCX APO L40×/0.80 W U-V-I objective; Leica). Micrographs were taken with a digital camera (Canon Power Shot S40).
Patch-Clamp Experiments
Membrane currents were recorded using standard patch-clamp techniques according to Hamill et al. (1981). Micropipettes were pulled from borosilicate glass microcapillaries (GC150F-10; Clark Electromedical Instruments) using an L/M-3P-A puller (List-Medical). The standard experimental solutions contained 60 mol m−3 potassium-gluconate, 2 MgCl2 mol m−3, 300 mol m−3 mannitol, 10 mol m−3 Bis-Tris propane, titrated with MES to pH 7.5 in the pipette, and 6 mol m−3 potassium-gluconate, 1 mol m−3 MgCl2, 1 mol m−3 CaCl2, 400 mol m−3 mannitol, 5 mol m−3 MES, titrated with Bis-Tris propane to pH 5.5 in the bath. The Ag/AgCl electrode was connected to the bath by a 3% (w/v) agar bridge filled with 100 mol m−3 KCl. While applying a positive pressure to the pipette, the pipette tip was dipped into the bath and brought into contact with a SE protoplast with the aid of an LN-SM-1 micromanipulator. After compensation of the offset potential of the pipette, contact was made between the pipette tip and the protoplast and gentle suction was applied to obtain a gigaseal. The membrane under the patch was broken by a short bipolar voltage pulse (±720 mV, 2.5 ms each) and simultaneous suction to obtain the whole-cell configuration. Whole-cell currents were recorded with an EPC-9 patch-clamp amplifier (HEKA Elektronik), filtered with an eight-pole Bessel filter with a cutoff frequency of 200 Hz, and sampled five times the filter frequency at 1 kHz on a personal computer. Data were digitized (ITC-16; Instrutech) and analyzed using PULSE and PULSFIT software (HEKA Elektronik). Voltages and currents were given with reference to extracellular side of the membrane as ground (Bertl et al., 1992). All membrane voltages were corrected off-line for liquid junction potential (Neher, 1992) using an LJP-calculator (Ng and Barry, 1995). All measurements were carried out at ambient temperatures between 20°C and 22°C.
Acknowledgments
We thank Prof. Dr. Hubert Felle for his permanent willingness to constructive discussions and critical reading of the manuscript, Dr. Martin Fronius for the supply of the patch-clamp equipment and expert technical advice, Prof. Dr. Wolfgang Clauss (Institute of Animal Physiology, JLU Giessen) for the hospitality in his institute, Kai Konrad and Prof. Dr. Rainer Hedrich (Julius-von-Sachs Institute for Biosciences, University of Würzburg) for helpful comments on patch-clamp studies, and Tina Henrich for dedicated technical assistance.
Footnotes
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The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Jens B. Hafke (jens.hafke{at}bot1.bio.uni-giessen.de).
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↵1 This work was supported by the Deutsche Forschungsgemeinschaft in the frame of Schwerpunktprogramm 1108 (BE 1925/8–2).
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↵[OA] Open Access articles can be viewed online without a subscription.
- Received July 24, 2007.
- Accepted September 15, 2007.
- Published September 20, 2007.
LITERATURE CITED
