Plant Cell Biology Research Group, Institute of General Botany, Justus-Liebig University, D–35390 Giessen, Germany
Sieve element (SE) protoplasts were liberated by exposing excisedphloem strands of Vicia faba to cell wall-degrading enzyme mixtures.Two types of SE protoplasts were found: simple protoplasts withforisome inclusions and composite twin protoplasts—twoprotoplasts intermitted by a sieve plate—of which oneprotoplast often includes a forisome. Forisomes are giant proteininclusions of SEs in Fabaceae. Membrane integrity of SE protoplastswas tested by application of CFDA, which was sequestered inthe form of carboxyfluorescein. Further evidence for membraneintactness was provided by swelling of SE protoplasts and forisomedispersion in reaction to abrupt lowering of medium osmolarity.The absence of cell wall remnants as demonstrated by negativeCalcofluor White staining allowed patch-clamp studies. At negativemembrane voltages, the current-voltage relations of the SE protoplastswere dominated by a weak inward-rectifying potassium channelthat was active at physiological membrane voltages of the SEplasma membrane. This channel had electrical properties thatare reminiscent of those of the AKT2/3 channel family, localizedin phloem cells of Arabidopsis (Arabidopsis thaliana). All inall, SE protoplasts promise to be a powerful tool in studyingthe membrane biology of SEs with inherent implications for theunderstanding of long-distance transport and signaling.
Use of fungal enzymes that degrade plant cell walls enablesthe isolation of plant cell protoplasts, which have become aninvaluable tool in plant biology. For example, protoplasts haveyielded considerable insight into plasma membrane-bound ionchannels and carbohydrate carriers in a variety of plant cellsranging from large parenchyma cells to tiny guard cells.
Due to technical barriers, sieve elements (SEs) are missingfrom other cell types that have been protoplasted successfully.A major problem is the tuning of the digesting mixture; theconventional enzyme mixes turn phloem tissues into a mash. Anotherproblem is the unequivocal identification of SE protoplasts.They easily fragment into smaller protoplasts during isolationand, therefore, can hardly be distinguished from those of other,smaller cell types.
Nevertheless, it remains tempting to isolate and identify SEprotoplasts for several reasons. The SE plasma membrane containsnumerous ion channels and carbohydrate carriers that are essentialfor sieve tube functioning (e.g. Patrick et al., 2001; van Bel,2003). Ion channels are not only meaningful for the ion householdof SEs, but also for the regulation of photoassimilate transportrates through sieve tubes (Fromm and Bauer, 1994; Ache et al.,2001; Deeken et al., 2002; van Bel and Hafke, 2005). They alsoplay a central role in long-distance signaling, such as thepropagation 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 havebeen 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 phloemby in situ hybridization techniques (Marten et al., 1999; Lacombeet al., 2000; Ache et al., 2001). However, the exact cellularlocalization, ion channel densities, ion channel types, anddistribution along the phloem path are unknown. SE protoplastsisolated from the respective phloem sections would provide aunique tool for unequivocal information about these issues.The same applies to calcium channels, which have been postulatedto occur in the SE plasma membrane (Volk and Franceschi, 2000).SE protoplasts from successive phloem sections would also enableto identify, characterize, and quantify carbohydrate carriersin the SE plasma membrane at various sites along the phloemtranslocation pathway. Differential deployment of sugar carriersis 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 membranebiophysics. The mass flow through the pressurized sieve tubesmakes high demands on the physical properties of the SE plasmamembrane. Knowledge of physical properties like elasticity andfluidity and their impact on the activity of transmembrane proteinssuch as mechanosensitive channels is necessary for a betterunderstanding of the pressure regulation of phloem transport.
For isolation of SE protoplasts, we employed Vicia faba phloemsince SEs in this species contain giant calcium-sensitive proteinbodies (forisomes; Knoblauch et al., 2001, 2003) meant to actas tools for SE identification. Here, we present a method forpreparation and identification of functional SE protoplastsof V. faba. The integrity of the SE protoplasts was tested byuse of fluorochromes and osmotic treatments. Furthermore, patch-clampexperiments were carried out to investigate the functionalityof SE protoplasts.
Isolation and Identification of V. faba SE Protoplasts
Following an incubation period of 10 h in the enzymatic digestionmedium, formation of phloem protoplasts released from disintegratingV. faba phloem strands was observed using light microscopy andconfocal laser scanning microscopy (CLSM). The enzymatic treatmentliberates protoplasts of various phloem cell types, which arethen tracked and identified by screening the phloem strandsunder light microscopy. Phloem protoplasts are always producedin low but sufficient numbers for patch-clamp studies, or generalbiophysical and physiological studies (see section on functionalityof SE protoplasts below).
Phloem protoplasts are highly variable in shape, size, and structure.Two types of SE protoplasts were found. Simple SE protoplastsarise from more central parts of disintegrating SEs and areidentified by inclusion of a large protein body named the forisome(Fig. 1A ). Forisomes are responsible for sieve plate occlusiondue to a turgor- and damage-induced, calcium-dependent conformationchange in intact SEs of legumes (Knoblauch et al., 2001, 2003).Thus, protoplasts containing a forisome must inevitably be SEprotoplasts.
Companion cell protoplasts (CCPs) were often found adhered toSEs (Fig. 1, C and D). CCPs typically contain chloroplasts aggregatedat one side; the cytoplasmic compartment occupies 20% to 25%of the total protoplast volume. A quick calculation shows thatthe spindle-shaped CCs with a diameter of 3 to 5 µm anda length of 200 to 300 µm would indeed produce sphericalCCPs 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 protoplastswith an average diameter of 40 to 60 µm. They can readilybe distinguished from CCPs by the equal distribution of thecytoplasm at the margin, matching about 1% of the total protoplastvolume (Fig. 1B).
Membrane Integrity of V. faba SE Protoplasts
As a test for membrane integrity, SE protoplasts were loadedwith the colorless CFDA-AM ester as described for intact phloemtissue (e.g. Knoblauch and van Bel, 1998). During incubation,the ester was cleaved by intracellular SE esterases (Oparkaand Read, 1994) into the membrane-impermeant and fluorescentform carboxyfluorescein (CF). Containment of CF inside the protoplastsafter washing demonstrates the integrity of the SE protoplastmembrane (Fig. 1E).
Information on the intactness of the connection between bothprotoplasts in a twin protoplast was obtained by labeling SEprotoplasts with RH-414, a membrane-soluble fluorescent probe(Fig. 1F). The CLSM pictures of RH-414 staining show labelingof a membrane system enveloping the large forisome-containingand the small protoplast of SE twin protoplasts. Continuingfluorescent striping in the sieve plate area indicates thatthe extensive plasma membrane system lining the sieve poreshad remained intact during the isolation procedure (Fig. 1F).
Mechanism of SE Protoplast Formation
The mechanism of SE protoplast formation is not yet understoodin detail. Precursors of the composite SE twin protoplast emergenear the sieve plates (Fig. 1, B and G). At either side of asieve plate, the SE membrane collapses in such a way that taperingends 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 appearsat either side of the sieve plate (Fig. 1, G and J). At thisstage, the twin protoplast precursor is already sealed as evidencedby CF accumulation in both protoplasts (Fig. 1I). After constrictionof the membranous tails, the composite SE protoplast startsrounding off before it gradually detaches from the phloem strand.
Formation of simple SE protoplasts (Fig. 1A) also depends onmembrane tail formation (Fig. 1H). Thus, both simple and compositeSE protoplasts rely on coalescence and constriction of membranoustails as a crucial step in protoplast formation. The amalgamationmechanism 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 aggregateresult of various bottlenecks in SE protoplast preparation asoutlined 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 uniformdiffusion of the enzyme mixture.
2. The sensitivity of phloem tissue to wounding.
In comparison to other cell types, the SE/CC complex is verysensitive to the slightest injury. Phloem slicing induces massivewound effects and turgor changes and triggers a physiologicaland structural collapse of most SEs in a tissue slice. Thus,the preparation method requires the use of thick phloem strandsto have a few intact SEs and imposes unavoidable artifacts thatminimize SE protoplast yield.
3. Composition of the enzyme mixture.
The composition of the enzyme mixture is a critical factor forthe success of SE protoplast formation in any plant species.Extensive concentration tests (not presented here) evidencedthat only enzyme mixtures in a narrow concentration window leadto successful cell wall degradation and a complete detachmentof SE protoplasts. Higher concentrations turn the phloem tissueinto mash; lower ones only liberate parenchyma protoplasts.
4. Composition of the SE cell wall.
SEs of dicotyledonous plants develop cell walls thicker thanthose 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, thenacreous layer with a pearly appearance (Evert, 1990). The complexityof the SE cell wall is a potential ground for cumbersome enzymaticdigestion.
5. Constriction of the membranous tails.
Transformation of parenchyma cells into protoplasts "only" demandsthe removal of cell wall material and the breakage of plasmodesmata.The production of SE protoplasts is more complicated in thatthe large SEs fragment during the isolation process, which requiresconsiderable membrane reconstitution. Presumably, the SE plasmamembrane is collapsing during the enzyme treatment. In a fewcases, the membrane constricts and coalesces at one side (compositeprotoplasts) or at both sides (single protoplasts) of the emergingprotoplast body. Coalescence of free membrane ends to a closedtail (Fig. 1, G and H) is a critical step toward formation ofviable SE protoplasts. In most cases, a mismatch between themembrane ends or an incomplete coalescence of membrane tailsprevents the final formation of SE protoplasts. It should benoted that the emergence of a composite protoplast depends onsuccessful sealing at either side of the sieve plate. In protoplastswith a sieve plate, the creation of the twin configuration isnecessary since otherwise the protoplast is not sealed at bothsides.
Given the numerous handicaps, it may be some time before onecan expect to gain high yields of SE protoplasts.
Functionality of SE Protoplasts; Activity of Calcium Channels
Indicative of membrane integrity of SE protoplasts is theirreaction to osmotic shocks. SE protoplasts were shocked osmoticallyby an abrupt change from 600 to 50 mol m–3 mannitol inthe external medium by microperfusion, while the external calciumconcentration was maintained constant at 1 mol m–3 (Fig. 2, A and B ).The sudden decline in the external osmolarity induced a gradualswelling (Fig. 2, B–D) within 30 to 120 s. Further protoplastswelling resulted in a burst (not shown) of the SE protoplast.The forisome inside SE protoplasts dispersed instantaneouslyin response to the osmotic shock (Fig. 2, A–D) in keepingwith the forisome behavior in intact SEs (Knoblauch et al.,2001). The calcium-dependent forisome dispersion (Knoblauchet al., 2003) is ascribed to calcium influx due to activationof mechanosensitive calcium channels.
The functionality of SE protoplasts was further demonstratedby applying suction to the SE plasma membrane via a microcapillaryconnected to a pressure device (Fig. 3, A–D ). In reactionto suction, the forisome dispersed presumably as a result ofCa2+ influx through mechanosensitive channels (Fig. 3, A–D).Forisome dispersion failed to occur (Fig. 3, E–H) in thepresence of the Ca2+ channel blocker Gd3+ (2 mol m–3).These results (Fig. 3, A–H) again indicate functionalmechanosensitive calcium channels in the SE plasma membrane.
To test the suitability for patch-clamp studies, SE protoplastswere tested on cellulosic and callosic wall remnants by CalcofluorWhite (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 withCW. In intact phloem tissue, cell walls, sieve plates, as wellas the pore plasmodesmata units were stained intensively (Fig. 4A).The CW staining gradually disappeared with incubation time inthe digesting medium (Fig. 4B), indicative of dissolution ofthe cell wall. After an incubation time of 10 h, several SEprotoplasts solely showed CW staining of the sieve plate (Fig. 4, C and D).
Simple patch-clamp measurements were executed using SE protoplaststo merely test whether their membranes were functional. Underprevailing conditions, inward and outward currents were observed.
At negative membrane voltages, instantaneous and time-dependentcurrents were observed in response to a series of test voltagesbetween –172 and +53 mV. The corresponding steady-statecurrent-voltage relationship recorded in asymmetrical potassium-gluconatesolutions showed that the currents only weakly rectified atmembrane voltages between +53 mV and –172 mV (Fig. 5, B and D ).The I-V plot of the steady-state currents (Fig. 5B) revealeda reversal voltage at –43 ± 2 mV (n = 4), closeto the predicted equilibrium voltage for a 10-fold K+ gradient(EK+ = –58 mV) across the membrane, but was differentfrom that of Mg2+ (–9 mV), Cl– (0 mV), or gluconate(+58 mV). Since the resting potential of the SE plasma membranein V. faba is around –130 mV (Hafke et al., 2005) andthus more negative than EK+, the observed channel may contributeto K+ loading/retrieval into the SE.
Clamping the SE plasma membrane from a holding potential of–22 mV to test voltages between –172 mV and +153mV results in activation of time-dependent outward currents(Fig. 5, C and D) positive to +70 mV. In addition, instantaneouscurrents were observed. For tail-current analysis (Fig. 5E),the plasma membrane of the SE protoplast was clamped from theholding voltage of –78 mV to a conditioning voltage of+153 mV to activate the time-dependent component. In a subsequentstep, the plasma membrane was clamped to a series of test voltagesbetween –172 mV and +53 mV. During the test pulse, themacroscopic tail currents gradually deactivated (Fig. 5E). Aplot of tail-current amplitude revealed a reversal voltage of–50 mV (Fig. 5F), close to the equilibrium voltage forK+ (–58 mV). Under prevailing artificial conditions withpotassium-gluconate on both sides of the membrane, this channelis not active at physiological membrane voltages.
SE Protoplasts from Other Plant Species
Modifications of the preparative steps with respect to enzymeconcentrations, preparation temperatures, and incubation timesyielded 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 intermittedby a sieve plate. After countless tests, typical isodiametricround protoplasts separated from the sieve plate have only beenobtained for Nicotiana. Despite a broad range of digestion conditionstested with respect to enzyme composition, osmolarity, durationof incubation, and temperature of the enzyme mixture, solelythe longish precursor of Cucurbita SE protoplasts was produced.This form is ascribed to cell wall remnants residing on theplasma membrane. In view of additional experience acquired withother species, preparation of SE protoplasts seems to be highlyspecies specific and the yield will probably always remain low.
SE protoplasts are a promising tool for studying phloem biophysics.In the near future, V. faba SE protoplasts may be adopted asa model system for transporter deployment in the SE plasma membranein view of the relatively easy mode of isolation and the resultsobtained with intact Vicia plants by other groups.
Vicia faba ‘Witkiem’, Cucurbita pepo ‘GelberZentner’, and Nicotiana tabacum plants were grown in potsin a greenhouse at temperatures varying between 20°C and30°C at 60% to 70% humidity and a 14-h/10-h light/dark periodwith supplementary lamp light (model SONT Agro 400 W; Phillips).The irradiance level was 200 to 250 µmol m–2 s–1at the plant apex. Plants were all taken in the vegetative periodjust before flowering.
Internodes were excised from 3- to 4-week-old V. faba plants.Then, tangential cuts were made to split the internodes. Forcoarse mechanical isolation of stem phloem strands, tangentialtissue sheets with a thickness of approximately 300 µmwere sliced with a razor blade from the fracture face of thesplit internode. After preincubation of the slices for 15 minin a standard medium (WM) containing 600 mol m–3 mannitol,1 mol m–3DL-dithiotreitol (DTT), and 25 mol m–3MES/NaOH, pH 5.7, the tissue was transferred into an enzymemixture containing 400 mol m–3 mannitol, 100 mol m–3KCl, 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 phloemstrands were filtered through a 80-µm nylon mesh and washedtwo times with the appropriate experimentation solution. Forpatch-clamp experiments, protoplasts were washed with standardbath solution and collected by centrifugation (Pico Fuge; Stratagene)twice.
The mechanism of protoplast formation and detachment were observedunder microscopic surveillance (Leica DM-LB, fluorescence microscope,equipped with a special water immersion objective, HCX APO L40x/0.80W U-V-I objective; Leica). The protoplasts were transferredinto a small volume of WM in a bathing chamber equipped witha microperfusion system. Here, SE protoplasts were treated withvarious solutions and permanent microscopic surveillance.
Light micrographs were taken with a digital camera (Canon PowerShot S40) connected to a computer (Canon Digital Camera SolutionDisk v8.0 software package).
SE protoplasts of N. tabacum and C. pepo were isolated as describedfor V. faba with slight modifications of the enzyme mixture,incubation time, and isolation temperature. SE protoplasts ofN. tabacum were isolated over a period of 4 h and a temperatureof 31°C in the above-mentioned enzyme mixture containing0.55% cellulase and 0.035% pectolyase. SE precursors of Cucurbitawere isolated over a period of 14 h and a temperature of 4°Cin the above-mentioned enzyme mixture containing 0.6% cellulaseand 0.04% pectolyase.
Staining of Protoplasts with CFDA, RH-414, and CW
To test their membrane integrity, SE protoplasts were loadedwith CFDA-AM ester (Molecular Probes) as described for intactphloem tissue (Knoblauch and van Bel, 1998). The solution wasprepared from a CFDA-AM ester stock solution dissolved in WMto give a final concentration of 2.1 µM CFDA. After applicationof the CFDA-AM, protoplasts were incubated for 45 min at roomtemperature. During this period, the ester was cleaved by endogenousSE esterases into the membrane-impermeant and fluorescent formCF (Oparka and Read, 1994). Following thorough washing withWM, protoplast fluorescence was examined using CLSM (Leica TCS4D) with a Krypton-Argon laser (Omnichrome) at 488 nm as describedbefore (Knoblauch and van Bel, 1998).
The protoplast plasma membrane was stained using the membrane-solublefluorochrome RH-414 (Molecular Probes). RH-414 was diluted froma stock solution in WM to give a final concentration of 4.3µM in a manner described before for the membrane-solublefluorochrome RH-160 (Knoblauch and van Bel, 1998). Protoplastswere incubated for 5 min in RH-414 before washing with WM andscanning 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 mannitolfor 15 min. After thorough washing with WM, cells were observedunder an epifluorescence microscope (Leica DMLB) using a BP340-380 excitation filter and an LP 425 barrier filter combination(Leica Microsystems).
SE protoplasts were bathed in a hyperosmotic solution containing600 mol m–3 mannitol, 1 mol m–3 DTT, 1 mol m–3CaCl2, and 25 mol m–3 MES/NaOH, pH 5.7. An abrupt bathchange to a hypo-osmotic medium containing 50 mol m–3mannitol, 1 mol m–3 DTT, 1 mol m–3 CaCl2, and 25mol m–3 MES/NaOH, pH 5.7, by a homemade microperfusionsystem imposed an osmotic shock. As a control experiment forSE protoplast swelling in a calcium-free solution, protoplastswere preincubated in the hyperosmotic solution and osmoticallyshocked with a solution containing 50 mol m–3 mannitol,1 mol m–3 DTT, 4 mol m–3 EGTA, and 25 mol m–3MES/NaOH, pH 5.7.
SE protoplasts were bathed in the hyperosmotic standard solution(see above) containing 600 mol m–3 mannitol, 1 mol m–3DTT, 1 mol m–3 CaCl2, and 25 mol m–3 MES/NaOH, pH5.7. Mechanical stress (suction) was exerted on SE protoplastsvia patch-clamp microcapillaries connected to a pressure device(Cell Tram Oil microinjector; Eppendorf).
A microcapillary filled with the respective bathing medium wasmaneuvered to the protoplast by means of an LN SM-1 micromanipulator(Luigs & Neumann). Contact between protoplast and microcapillarywas made by suction with the aid of a Cell Tram Oil microinjector(Eppendorf). As a control, protoplasts were incubated in theabove-mentioned hyperosmotic solution supplied with 2 mol m–3of the calcium channel blocker Gd3+ (as GdCl3). Protoplastswere observed using an epifluorescence microscope (Leica DM-LB,fluorescence microscope, equipped with a special water immersionobjective, HCX APO L40x/0.80 W U-V-I objective; Leica). Micrographswere taken with a digital camera (Canon Power Shot S40).
Membrane currents were recorded using standard patch-clamp techniquesaccording to Hamill et al. (1981). Micropipettes were pulledfrom borosilicate glass microcapillaries (GC150F-10; Clark ElectromedicalInstruments) using an L/M-3P-A puller (List-Medical). The standardexperimental solutions contained 60 mol m–3 potassium-gluconate,2 MgCl2 mol m–3, 300 mol m–3 mannitol, 10 mol m–3Bis-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–3MES, titrated with Bis-Tris propane to pH 5.5 in the bath. TheAg/AgCl electrode was connected to the bath by a 3% (w/v) agarbridge filled with 100 mol m–3 KCl. While applying a positivepressure to the pipette, the pipette tip was dipped into thebath and brought into contact with a SE protoplast with theaid of an LN-SM-1 micromanipulator. After compensation of theoffset potential of the pipette, contact was made between thepipette tip and the protoplast and gentle suction was appliedto obtain a gigaseal. The membrane under the patch was brokenby 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-clampamplifier (HEKA Elektronik), filtered with an eight-pole Besselfilter with a cutoff frequency of 200 Hz, and sampled five timesthe filter frequency at 1 kHz on a personal computer. Data weredigitized (ITC-16; Instrutech) and analyzed using PULSE andPULSFIT software (HEKA Elektronik). Voltages and currents weregiven with reference to extracellular side of the membrane asground (Bertl et al., 1992). All membrane voltages were correctedoff-line for liquid junction potential (Neher, 1992) using anLJP-calculator (Ng and Barry, 1995). All measurements were carriedout at ambient temperatures between 20°C and 22°C.
We thank Prof. Dr. Hubert Felle for his permanent willingnessto constructive discussions and critical reading of the manuscript,Dr. Martin Fronius for the supply of the patch-clamp equipmentand expert technical advice, Prof. Dr. Wolfgang Clauss (Instituteof Animal Physiology, JLU Giessen) for the hospitality in hisinstitute, Kai Konrad and Prof. Dr. Rainer Hedrich (Julius-von-SachsInstitute for Biosciences, University of Würzburg) forhelpful comments on patch-clamp studies, and Tina Henrich fordedicated technical assistance.
Received July 24, 2007; accepted September 15, 2007; published September 20, 2007.
The author responsible for distribution of materials integralto the findings presented in this article in accordance withthe policy described in the Instructions for Authors (www.plantphysiol.org)is: Jens B. Hafke ([email protected] ).
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* Corresponding author; e-mail [email protected] .
Bauer CS, Hoth S, Haga K, Phillipar K, Aoki N, Hedrich R (2000) Differential expression and regulation of K+ channels in the maize coleoptile: molecular and biophysical analysis of cells isolated from cortex and vasculature. Plant J 24: 139–145
Deeken R, Geiger D, Fromm J, Koroleva O, Ache P, Langenfeld-Heyser R, Sauer N, May ST, Hedrich R (2002) Loss of AKT2/3 potassium channel affects sugar loading into the phloem of Arabidopsis. Planta 216: 334–344
Hafke JB, van Amerongen JK, Kelling F, Furch ACU, Gaupels F, van Bel AJE (2005) Thermodynamic battle for photosynthate acquisition between sieve tubes and adjoining parenchyma in transport phloem. Plant Physiol 138: 1527–1537
Lacombe B, Pilot G, Michard E, Gaymard F, Sentenac H, Thibaud JB (2000) A shaker-like K+ channel with weak rectification is expressed in both source and sink phloem tissues of Arabidopsis. Plant Cell 12: 837–851
Ng B, Barry H (1995) The measurement of ionic conductivities and mobilities of certain less common organic ions needed for junction potential corrections in electrophysiology. J Neurosci Methods 56: 37–41
Oparka KJ, Read ND (1994) The use of fluorescent probes for studies on living plant cells. In N Harris, KJ Oparka, eds, Plant Cell Biology: A Practical Approach. Oxford University Press, Oxford, pp 27–50
Phillipar K, Büchsenschütz K, Abshagen M, Fuchs I, Geiger D, Lacombe B, Hedrich R (2003) The K+ channel KZM1 mediates potassium uptake into the phloem and guard cells of the C4 grass Zea mays. J Biol Chem 278: 16973–16981
van Bel AJE, Hafke JB (2005) Physicochemical determinants of phloem transport. In NM Holbrook, M Zwieniecki, eds, Long-Distance Transport Systems in Plants: Integration and Coordination. Elsevier Science, New York, pp 19–43
Figure 1. 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).
Figure 2. 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.
Figure 3. 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.
Figure 4. 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.
Figure 5. 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.
Figure 6. 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.