methodology open access isolation of dimorphic ...methodology open access isolation of dimorphic...

METHODOLOGY Open Access

Isolation of dimorphic chloroplasts from thesingle-cell C4 species Bienertia sinuspersiciShiu-Cheung Lung, Makoto Yanagisawa and Simon DX Chuong*

Abstract

Three terrestrial plants are known to perform C4 photosynthesis without the dual-cell system by partitioning twodistinct types of chloroplasts in separate cytoplasmic compartments. We report herein a protocol for isolating thedimorphic chloroplasts from Bienertia sinuspersici. Hypo-osmotically lysed protoplasts under our defined conditionsreleased intact compartments containing the central chloroplasts and intact vacuoles with adhering peripheralchloroplasts. Following Percoll step gradient purification both chloroplast preparations demonstrated highhomogeneities as evaluated from the relative abundance of respective protein markers. This protocol will opennovel research directions toward understanding the mechanism of single-cell C4 photosynthesis.

Keywords: Bienertia sinuspersici, Chloroplast isolation, Dimorphic chloroplasts, Osmotic swelling, Photosynthesis,protoplast, Single-cell C4, Vacuole isolation

BackgroundThe majority of terrestrial plants house chloroplasts pri-marily in one major cell type of leaves (i.e. mesophyllcells), and perform C3 photosynthesis to assimilate atmo-spheric CO2 into a 3-carbon product, 3-phosphoglycericacid. In C4 species, on the other hand, a Kranz-type leafanatomy featuring the second type of chlorenchyma cellssurrounding the vascular bundles (i.e. bundle sheathcells) was reported as early as in the late 1800’s [1]. Inthese species, the initial carbon fixation into 4-carbonacids was first documented in the 1960’s [2,3]. The phy-siological relevance of the Kranz anatomy in relation tothe C4 photosynthetic pathways, however, had not beenelucidated until the successful separation of the twotypes of chlorenchyma cells and their respectivedimorphic chloroplasts. With the development of variousmechanical and enzymatic methods for separating themesophyll and bundle sheath cells, the biochemistry ofC4 cycles has been intensively studied over the past fewdecades focusing explicitly on characterizing the enzy-matic properties and determining their precise subcellu-lar locations in these cell types (for review, see [4]),leading to the current C4 models. In the C4 model, atmo-spheric CO2 is initially converted into C4 acids by

phosphoenolpyruvate carboxylase (PEPC) in mesophyllcells. The C4 acids are broken down by a C4 subtype-spe-cific decarboxylation enzyme in bundle sheath cells, andthe liberated CO2 is subsequently re-fixed by ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco). TheC4 pathway concentrates CO2 at the site of Rubisco andminimizes the photorespiration process, an unfavorableoxygenase activity of Rubisco with O2.The indispensable relationship between the Kranz anat-

omy and C4 photosynthesis has been an accepted featureuntil the discovery of three terrestrial single-cell C4 spe-cies, Suaeda aralocaspica (formerly called Borszczowiaaralocaspica) [5], Bienertia cycloptera [6,7], and B. sinus-persici [8] in the Chenopodiaceae family. In chlorench-yma cells of these succulent Chenopodiaceae species, theC4 cycles are operational in the absence of Kranz anat-omy due to the division of cytoplasm into two compart-ments. The cytoplasmic channels that connect the twocompartments not only allow metabolite exchange butalso limit inter-compartmental gas diffusion, resemblingthe function of plasmodesmata traversing the thickenedand sometimes suberized bundle sheath cell wall. Thetwo cytoplasmic compartments house two distinct chlor-oplast types and different subsets of enzymes, respec-tively. Accordingly, the peripheral compartment proximalto the CO2 entry point is specialized for carboxylationand regeneration of the initial carbon acceptor,

* Correspondence: [email protected] of Biology, University of Waterloo, 200 University Avenue West,Waterloo, Ontario N2L 3G1, Canada

Lung et al. Plant Methods 2012, 8:8http://www.plantmethods.com/content/8/1/8

PLANT METHODS

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phosphoenolpyruvate, whereas the central compartmentdistal to the CO2 entry point is responsible for decarbox-ylation of C4 acids and Rubisco-catalyzed re-fixation ofthe liberated CO2. In agreement with the immunolocali-zation patterns of the major enzymes involved [5,7,9], thetwo cytoplasmic compartments appear to be functionallyequivalent to the mesophyll and bundle sheath cells ofKranz-type C4 plants, respectively. Based on differentialspeed centrifugation for the enrichment of each chloro-plast type in subcellular fractions of B. sinuspersici leaves,Offermann et al. [10] have recently examined the proteindistribution patterns of dimorphic chloroplasts and con-firmed their functional similarities to the mesophyll andbundle sheath cells of Kranz-type C4 plants, respectively.Thorough studies of the enzymology of the single-cell C4

model, however, requires homogenous preparations ofthe dimorphic chloroplasts with specific techniquesbased on the unique cell anatomy, in analogy with theprevious efforts of separation and subcellular fractiona-tion of the dual cell types from Kranz-type C4 plants (forreview, see [4]).Here, we present our empirically optimized protocol

for separating the dimorphic chloroplasts from chlor-enchyma protoplasts of B. sinuspersici. By reducing theosmotic potential of culture medium to a suitable level,the isolated protoplasts were hypo-osmotically bursted,concomitantly extruding one type of chloroplasts encasedin the central cytoplasmic compartment and another typeof chloroplasts from the peripheral compartment adheredto the external surface of intact vacuoles. Following asubsequent centrifugation step, these two structures canbe separated into the sedimented and floating fractions,respectively. Finally, two homogenous populations ofdimorphic chloroplasts with minimal cross-contamina-tion can be further purified by using a Percoll gradient.Overall, this dimorphic chloroplast isolation protocol canbe applied in multiple areas of research toward furtherunderstanding the development of single-cell C4 systems.

ResultsRationale of the isolation procedures in relation to cellanatomyFor a better clarification of the each isolation step asdescribed in the subsequent sections, we first summarizethe major changes in the complex subcellular organiza-tion of chlorenchyma cells throughout the process as illu-strated with schematic diagrams (Figure 1). In a maturechlorenchyma cell, a large central vacuole (as depicted ingrey) separates the cytoplasm (yellow) into the peripheral(PCC) and the central (CCC) cytoplasmic compartments,which are interconnected by cytoplasmic channels traver-sing the vacuole (Figure 1A). The rounding of protoplastsby enzymatic digestion of the cell wall causes no changesin the integrity of the two cytoplasmic compartments

and the cytoplasmic channels (Figure 1B). Reducing theosmotic potential of the cell-stabilizing buffer inducesthe swelling of the protoplast and its central vacuole, dis-rupting the cytoplasmic channels and pushing the CCCagainst the plasma membrane (Figure 1C). The hypo-osmotic shock causes the plasma membrane to stretchbeyond its extension limit that eventually breaks, releas-ing the intact CCC and vacuole with attached peripheralchloroplasts (P-Chls; Figure 1D). This is in agreementwith the previous observation showing that the two cyto-plasmic compartments are separated by a single vacuole[9,11]. Eventually, the central vacuole carrying P-Chls onthe external surface (Figure 1E) and the isolated CCCcontaining central chloroplasts (C-Chls; Figure 1F) canbe separated into two discrete entities.

Isolation of chlorenchyma protoplasts from B. SinuspersiciAs a first step, we isolated chlorenchyma cells fromB. sinuspersici leaves and enzymatically prepared a homo-genous population of healthy protoplasts. Interestedreaders should refer to our previous report for detailedtechnical considerations on plant growth conditions andchlorenchyma protoplast isolation [12]. Previously, pro-gressive developmental variation has been identified atdifferent stages of B. sinuspersici leaves in terms of theunique subcellular compartmentation and photosyntheticgene expression [11,13]. Similar developmental gradientswere also observed across the base-to-tip dimension ofleaves [13]. Thus, these major sources of variability wereinevitably taken into considerations in order to standar-dize the degree of C4 functionality of the starting materi-als for dimorphic chloroplast isolation. To this end, weroutinely propagated B. sinuspersici by vegetative cuttingsand collected entire leaves, 2 cm or longer in length,from healthy branches of 3- to 4-month-old plants(Figure 2A). At this stage, the single-cell C4 compartmen-tation has reached maturity as evident by the presence ofthe distinctive subcellular distribution of dimorphicchloroplasts in the majority of chlorenchyma cells (Figure2B). In a mature chlorenchyma cell, the C-Chls are den-sely packed into a large, spherical CCC structure,whereas the P-Chls are distributed throughout the thinlayer of cytoplasm, (PCC; Figure 2B). Following cell wallremoval by cellulase treatment under our optimized con-ditions, the resulting protoplasts exhibited no observablechanges in their unique chloroplast distribution, which isconsidered a prerequisite for subsequent preparations ofpure dimorphic chloroplasts (Figure 2C). During the pro-toplast isolation, we occasionally observed vesicles withexternally adhering P-Chls (Figure 2D). Given theirpotential of serving as an excellent source of pure P-Chls,we continued to optimize the conditions for inducing theformation of these P-Chl-containing vesicles fromisolated protoplasts. To guarantee intactness and full


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Figure 1 Illustration of the cellular and subcellular morphologies during the process of dimorphic chloroplast isolation. (A) A maturechlorenchyma cell. For clarity, only one cytoplasmic channel is illustrated despite the multiple occurrences in an actual cell. (B) An isolatedprotoplast with digested cell wall. (C) A swelling protoplast with increased osmotic pressure of the cell sap as indicated by arrows. (D) A brokenprotoplast with torn plasma membrane as indicated by arrows. (E) An intact vacuole carrying peripheral chloroplasts on the external surface. (F)An isolated central cytoplasmic compartment. Scale bars = 10 μm (A-E) or 5 μm (F).


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functionality of the purified dimorphic chloroplasts, atleast 90% viability of the isolated protoplasts should beachieved prior to chloroplast separation as evaluated byfluorescein diacetate staining (Figure 2E).

Hypo-osmotic shock treatment of isolated protoplastsinduced vesicle formationPreliminary trials suggested that the vesicles with adher-ing P-Chls could be derived from the swelling proto-plasts in an osmotic potential-dependent manner.Accordingly, we carried out time-lapse imaging of thevesicle formation from isolated protoplasts subjected tohypo-osmotic shock treatments. We diluted the sucroseconcentration of the protoplast culture medium from0.7 M to various concentrations. When the sucrose con-centration was substantial decreased to 0.1 M, the iso-lated protoplasts expanded rapidly and lysedapproximately 15 s after dilution, releasing the CCCsand vesicles with P-Chls (Figure 3A; Additional file 1).The dense CCC structures gradually loosened and theencased C-Chls eventually dissociated from the CCCstructures, whereas the vesicles continued to expandbeyond the stretching limit of their membrane andeventually bursted causing the P-Chls to adhere tostrings of collapsed membrane which were inseparable

from the CCC structures (Figure 3A; Additional file 1).On the other hand, when the sucrose concentration wasreduced to the ideal 0.2 M, the isolated protoplasts gra-dually swelled and lysed approximately 25 s after dilu-tion, releasing the CCCs and P-Chl-carrying vesicles(Figure 3B; Addition file 2). Under this condition, themajority of C-Chls remained tightly packed within thedense CCC structures and the vesicles continued toswell for a few seconds yet remained intact after celllysis (Figure 3B; Additional file 2). Subtle reduction ofthe sucrose concentration to 0.5 M did not burst theisolated protoplasts, although some vesicles were occa-sionally pinched off from the intact swelling protoplasts(Figure 3C; Additional file 3). The majority of these vesi-cles, however, did not carry any chloroplasts (Figure 3C;Additional file 3).

The vacuolar origin of vesicles from lysed protoplastsThe time-lapse images clearly showed that the releasedvesicles from the hypo-osmotically lysed protoplasts ori-ginally made up the bulk of cell volume (Figure 3),implying that these vesicles derived from the large cen-tral vacuoles. Due to the unusual subcellular organiza-tion of B. sinuspersici chlorenchyma cells, we sought toconfirm the vacuolar origin of the vesicles by staining

Figure 2 Isolation of chlorenchyma protoplasts from B. sinuspersici. (A) A healthy branch of vegetatively propagated B. sinuspersici suitablefor protoplast isolation. (B) A cross section of B. sinuspersici leaf showing chlorenchyma cells with two types of chloroplasts, i.e. the peripheralchloroplasts (P-Chls) in the peripheral cytoplasmic compartments and the central chloroplasts enclosed in the central cytoplasmic compartments(CCC). (C) A bright field image of chlorenchyma protoplasts indicating the integrity of the two cytoplasmic compartments for subsequentisolation of the dimorphic chloroplasts. (D) A bright field image of a protoplast and a vesicle with externally associated P-Chls. (E) Vital stainingof isolated protoplasts imaged under epifluorescence microscopy. Fluorescein diacetate staining (left panel), chlorophyll autofluorescence (middlepanel) and a merged image (right panel). Scale bars = 10 mm (A) or 20 μm (B-E).


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with 5-(and-6)-carboxy-2’,7’-dichloro-fluorescein diace-tate (CDCFDA), a pH-sensing vital probe for thevacuole lumen [14]. In a CDCFDA-stained protoplast,prominent fluorescent signals were typically found inthe vacuole which occupies the massive cell contentseparating the CCC and PCC (Figure 4A). Followinghypo-osmotic shock treatment of CDCFDA-stained pro-toplasts, the fluorescent signals persisted in the resultingvesicles indicating their vacuolar origin (Figure 4B and4C). The occurrence of some patchy CDCFDA signalsin other subcellular locations (Figure 4), similar to theobservations from previous studies [9,11], might beattributed to the presence of reactive oxygen species[15]. In addition to the CDCFDA staining, the observa-tion of prismatic and raphide crystals in the vesiclesprovided further evidence of their vacuolar origin (Fig-ure 5). The occurrence of these crystals is a widespreadphenomenon in plant vacuoles due to the accumulationof calcium oxalate as crystalline deposits [16].

Subfractionation of dimorphic chloroplasts from lysedprotoplastsGiven a promising strategy to separate CCCs and intactvacuoles with adhering P-Chls by osmotic treatment ofisolated protoplasts, we further optimized the conditionsfor the best yields of homogenous populations of thesetwo suborganellar structures. Since preliminary trials

suggested that the isolated protoplasts under an over-crowded condition responded heterogeneously to theosmotic shock, we routinely adjusted the cell density to105 protoplasts mL-1 before the treatment to ensure thehighest percentage of protoplast lysis. Pre-conditioningof the isolated protoplasts at 0°C potentially renderedtheir plasma membrane more brittle due to the reducedfluidity and facilitated the subsequent cell lysis. Rapiddilution of the isolated protoplasts at room temperaturein an EDTA-containing buffer without the sucrose osmo-ticum resulted in stretching and fragmentation of theplasma membrane. This observation is considered a com-bined effect of protoplast swelling due to the reducedosmotic strength of the buffer and the sudden alternationin membrane fluidity due to the thermal change and thechelation of divalent ions. At the optimum 0.2 M concen-tration of sucrose, 98% of isolated protoplasts were bur-sted and ca. 80% of the released vacuoles remained intact(Figure 6). On the other hand, relatively low percentagesof protoplasts were lysed at higher sucrose concentra-tions (i.e. 0.3-0.7 M), whereas the released vacuoles atunfavorably low sucrose concentration (i.e. 0.1 M) wereconcomitantly bursted (Figure 6). Taking together, weroutinely lysed the isolated protoplasts by rapid dilutionof the protoplast culture buffer to 0.2 M sucrose whichprovided the best yields of intact CCCs and intact P-Chl-containing vacuoles. Due to the substantial difference in

Figure 3 Time-lapse imaging of protoplasts subjected to osmotic swelling. (A) Diluting the cell stabilizing buffer to a sucrose concentrationof 0.1 M led to rapid swelling and bursting of both the protoplasts and their vacuoles. (B) Lowering the osmotic potential of the cell stabilizingbuffer to the optimum 0.2 M sucrose resulted in protoplast lysis and the release of intact central cytoplasmic compartments and vacuoles withattached peripheral chloroplasts. (C) Subtle dilution of the cell stabilizing buffer to 0.5 M sucrose did not lyse the protoplasts while vesicles wereslowly pinching off from the vacuoles of protoplasts as indicated by arrowheads. Scale bars = 20 μm.


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Figure 4 Vacuolar staining of chlorenchyma protoplasts and osmotically-derived vesicles. Isolated protoplasts were stained with 5-(and-6)-carboxy-2’,7’-dichloro-fluorescein diacetate (CDCFDA) and lysed by osmotic swelling. The isolated protoplasts and induced vesicles wereobserved under light and epifluorescence microscopy. CDCFDA staining, chlorophyll autofluorescence, merged images of the two channels, andbright field images are shown. (A) An isolated protoplast. (B) A swelling protoplast before cell lysis. (C) A vacuole-derived vesicle. Scale bars = 10μm.

Figure 5 Formation of calcium oxalate crystals in the vacuole-derived vesicles. The vesicles were obtained from lysis of protoplasts byosmotic swelling at 0.2 M sucrose and isolated on the floating layer after centrifugation at 100g, 4 min. Formation of calcium oxalate crystalswas induced by resuspension of the vesicles in cell stabilizing buffer with 0.7 M sucrose. (A) A light micrograph of prismatic crystals. (B) A lightmicrograph of raphide crystals. (C) A light micrograph of both prismatic and raphide crystals. Scale bars = 20 μm.


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densities of these two suborganellar structures, subse-quent incubation on ice and low-speed centrifugation(100g, 4 min) led to their separation into the sedimentedand floating fractions, respectively. The recovered pelletsand floating layers contained a homogeneous populationof CCCs (Figure 7A) and intact vacuoles with adheringP-Chls (Figure 7B), respectively. The CCCs could beeasily dispersed mechanically by pipetting up and downfollowing the cold treatment and the C-Chls releasedfrom the CCCs were further purified from other impuri-ties on a typical two-step Percoll gradient. By centrifuga-tion (2,500g, 10 min) of the P-Chl-containing vacuoles ona similar Percoll gradient, the loosely adhering P-Chlswere easily dissociated from the vacuoles and entered the

Percoll solutions. In both cases of C-Chls and P-Chls, thelower green band at the 40%/85% interface contained atleast 50% of the loaded chloroplasts. The purity andintactness of the aspirated chloroplasts from this inter-face were confirmed by phase contrast microscopy (datanot shown).

Evaluating cross-contamination of the isolated dimorphicchloroplastsThe banding patterns of the resolved polypeptides fromthe isolated dimorphic chloroplasts were visualized andcompared by silver staining (Figure 8A). Various differ-ential band intensities were identified by comparing thetwo protein profiles, with the most striking difference

Figure 6 The release of central cytoplasmic compartments and vesicles from protoplasts under different osmotic conditions. Thesucrose concentration of the cell stabilizing buffer was either undiluted (0.7 M) or diluted to 0.1, 0.2, 0.3 or 0.5 M, and the protoplasts wereincubated on ice for 10 min. The number of intact protoplasts, central cytoplasmic compartments (CCC) and vacuole-derived vesicles werecounted using a haemocytometer under light microscopy. The occurrence (%) is calculated from the number of protoplasts, CCC or vesicles afterosmotic treatment divided by the number of protoplasts used. Each bar represents the mean value from three independent experiments ± 1standard deviation.


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found at the electropheretic mobility regions equivalentto pyruvate orthophosphate dikinase (PPDK; 97 kDa)and the large-subunit (LSU; 53 kDa) and small-subunit(SSU; 14 kDa) of Rubisco (Figure 8A). These proteindistribution patterns are in agreement with the recentobservations following the subcellular fractionation of P-Chls and C-Chls [10]. In fact, previous immunolocaliza-tion experiments also revealed that the labelling ofPPDK was mainly associated with the P-Chls, whereasthe LSU was predominantly detected in the C-Chls [9].Western blot analyses further confirmed that the immu-noreactive band of PPDK from P-Chls was two-fold

more intense than that from C-Chls, whereas LSU wasalmost exclusively found in C-Chls but barely detectablein P-Chls (Figures 8B, C). The relative distribution ofthe photosystem I and II proteins in the dimorphicchloroplasts (Figures 8B, C) also correlated with the pre-vious ultrastructural studies which indicated that C-Chlshad a higher granal index and generally greater sizes ofgrana stacks than P-Chls [7]. The Western blot analysesrevealed that the C-Chls had slightly more photosystemII manganese-stabilizing proteins (PsbO; Figures 8B, C),which are located predominantly in the stacked regionsof thylakoid membrane. The P-Chls, on the other hand,

Figure 7 Bright field images of the two isolated dimorphic chloroplast populations. (A) A homogenous population of central cytoplasmiccompartments. (B) A homogenous population of vacuole-derived vesicles carrying peripheral chloroplasts on the external surfaces. Scale bars =40 μm.

Figure 8 Silver staining and Western blot analyses of protein extracts from the isolated dimorphic chloroplasts. Two micrograms ofproteins from total chloroplasts (lane 1), peripheral chloroplasts (lane 2) and central chloroplasts (lane 3) were separated on SDS/12% PAGE.Molecular weights are shown on the left in kilodaltons. (A) Total proteins were visualized by silver staining. Polypeptides of the highly abundantpyruvate orthophosphate dikinase (PPDK), Rubisco large-subunit (LSU) and small-subunit (SSU) are indicated. Other bands of significantly differentintensities between the dimorphic chloroplasts are indicated by arrowheads. (B) Resolved proteins were electroblotted onto nitrocellulosemembranes and probed with antibodies against PPDK, LSU, Photosystem II manganese-stabilizing protein (PsbO), cytochrome f (Cyt-f), orPhotosystem I subunit II (PsaD). (C) The immunoblots were analyzed by densitometric quantification. The abundance of each protein inperipheral (P-Chls) and central (C-Chls) chloroplasts was calculated relative to that of total chloroplasts and shown with standard errors, afterthree (PsbO) or four (others) independent experiments.


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contained more photosystem I subunit II proteins(PsaD; Figures 8B, C), which are commonly found in theunstacked regions. The cytochrome f subunit of thecytochrome b6f complex, which is evenly distributedthroughout the thylakoid membrane, was equally abun-dant in both types of chloroplasts (Figures 8B, C). Over-all, the electrophoretic analyses indicated minimal cross-contamination of the two isolated populations ofdimorphic chloroplasts.

DiscussionPotential feasibility of isolating dimorphic chloroplastsfrom B. SinuspersiciIn view of the chlorenchyma cell anatomies of the threecurrently known terrestrial single-cell C4 species, isola-tion of dimorphic chloroplasts from the two Bienertiaspecies is technically more feasible than that fromS. aralocaspica. Firstly, chlorenchyma cells of the twoBienertia species (i.e. B. sinuspersici and B. cycloptera)enclose one type of chloroplasts within a distinctivespherical structure (CCC) at the centre of the cell andrandomly distribute another type of chloroplasts in athin layer of cytoplasm at the cell periphery (PCC).Separation of the dimorphic chloroplasts from chlor-enchyma cells of these species is therefore technicallyfeasible provided that the integrity of CCC is preservedduring the process. Secondly, the mature chlorenchymacells of Bienertia species effectively limit gas diffusionby the formation of CCCs in the cell interior, as ameans to reduce CO2 leakage at the site of C4 acid dec-arboxylation and exposure of Rubisco to O2, At the cellperiphery, the entire chlorenchyma cells are surroundedby extensive intercellular space to facilitate fixation ofatmospheric CO2 (Figure 2B) [7]. Therefore, the looselypacked chlorenchyma cells can be readily isolated bygently squeezing the leaves in a mortar and pestle [12].Technically, this rapid release of chlorenchyma cellsdevoid of contaminations such as epidermis, water sto-rage and vacuolar tissues facilitates the subsequent iso-lation of pure and functional dimorphic chloroplasts. Incontrast, mature leaves of S. aralocaspica partitiondimorphic chloroplasts by localization of the first chlor-oplast type at one pole of the elongated chlorenchymacells proximal to the atmosphere and the second chlor-oplast type at the opposite pole of the cells proximal tothe vascular tissues [5]. Thus, the operation of the sin-gle-cell C4 cycle in this species depends primarily onthe polarization of the dimorphic chloroplasts and theradial elongation of the cells for effective limitation ofCO2 and O2 diffusion [5,17]. Isolation of the two polar-ized chloroplast types solely based on the difference intheir subcellular locations might be technically challen-ging, if not impossible. Perhaps, we speculate that any

change in cell shape due to cell wall removal, as thefirst step in routine chloroplast isolation procedures,might distort the polarization of the dimorphic chloro-plasts, although the isolation of protoplasts from S. ara-locaspica chlorenchyma cells has yet been attempted.The leaf anatomy of S. aralocaspica also reveals abso-lute exclusion of intercellular air space toward the innerpoles of chlorenchyma cells due to the tight cell-celladhesion [5,17], implicating the difficulty in chlorench-yma cell isolation. Moreover, our successful plant pro-pagation by vegetative cuttings added another technicaladvantage of using B. sinuspersici leaves for large-scaleisolation of dimorphic chloroplasts with synchronizedC4 development.

Technical considerations for isolation of centralchloroplastsA reliable protocol for separating the dimorphic chloro-plasts is primarily based on the criteria of no cross-con-tamination. The isolation of C-Chls is relativelyeffortless due to the confinement of these organelleswithin the large, dense CCC structures, which can beeasily recovered from the cell lysates by low-speed cen-trifugation (Figure 7A). It is worthy of note, however,that the C-Chl preparation might be potentially con-taminated with P-Chls due to incomplete protoplastlysis. Previously, we reported a low-speed centrifuga-tion-based method for floatation of healthy, intact chlor-enchyma protoplasts on a sucrose medium andsedimentation of the stressed protoplasts [12]. In thepresent study, any protoplasts surviving from the osmo-tically mediated lytic treatment might therefore be co-sedimented with the CCC fraction upon low-speed cen-trifugation leading to contamination of this fraction withP-Chls. Accordingly, we optimized our procedures forosmotic shock treatment and showed that the isolatedprotoplasts at a suitable cell density were almost exclu-sively bursted under the defined chemical, thermal andosmotic conditions (Figure 6), leading to a homogenouspreparation (Figure 7A) and satisfactory purity (Figure8) of C-Chls. In addition to P-Chls, the potential con-tamination of C-Chl preparation with chloroplasts fromimmature chlorenchyma cells should also be taken intoaccount. The expression patterns of photosyntheticgenes strongly suggested that these chloroplasts fromimmature chlorenchyma cells, as evident in the youngleaf samples, might function in a C3 default mode ofphotosynthetic and photorespiratory pathways [13].Since no chloroplast clumping was observed at thisearly stage of leaf development [11], contamination dueto immature chlorenchyma cells is not considered amajor concern in the preparation of C-Chls using ourlow-speed centrifugation-based method.


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Technical considerations for isolation of peripheralchloroplastsOn the other hand, purification of P-Chls from the pro-toplast lysates is deemed technical more challenging.Recently, Offermann et al. [10] reported the first proto-col for dimorphic chloroplast isolation from B. sinusper-sici based on differential speed centrifugation. Accordingto our preliminary study, this centrifugation techniquedid not provide satisfactory purity of P-Chls due to twomajor problems. First, residual CCCs remained in thesupernatant after the low-speed centrifugation leadingto contamination of the P-Chl fraction. Attempts toincrease the centrifugal force or duration of centrifuga-tion, on the other hand, resulted in a considerable lossof P-Chls from the supernatant. More remarkably, thesupernatant fraction of P-Chls was contaminated withC-Chls due to their dissociation from CCCs, whichremains to be an unavoidable problem given the factthat the CCC is not confined by a membrane asrevealed at the electron microscopic level (data notshown). In fact, rapid dispersal of C-Chls was commonlyobserved when the chlorenchyma protoplasts were sub-jected to unfavorable culture conditions [12] or mechan-ical disruption such as a gentle press on a microscopicslide [10]. The integrity of CCC structures is primarilymaintained by the association of C-Chls in the outerregions of CCCs with the surrounding cytoskeleton net-works, of which microtubules have been found to berelatively important [9]. During the dimorphic chloro-plast isolation, the instability of the CCCs might beattributed to a combined effect of osmotic swelling ofC-Chls in the hypotonic medium and depolymerizationof microtubules by low temperature. Despite the conten-tious issue of cold treatment, chloroplast isolation athigher temperature is not recommended due to thepotential risk of protein degradation and denaturationafter cell lysis, particularly if chloroplasts are isolated forproteomics or enzymology studies. Similarly, chloro-plasts for protein import studies should be isolated atlow temperature due to the high susceptibility of thechloroplast protein import receptors to proteolysis [18].Of relevance to preventing proteolytic degradation, ourchloroplast isolation method has an added advantage byconfining the majority of proteolytic enzymes in theintact vacuoles.Due to the considerable contamination of the P-Chls

in the cell lysate with dissociated C-Chls, we sought analternative method for P-Chl purification by isolation offloating vacuoles as P-Chl carriers. The separation ofplant vacuoles from isolated protoplasts is a commonphenomenon in hypotonic solutions and was documen-ted as early as when protoplasts were isolated for thefirst time from a plant source [19]. Since then, the pro-toplast-based techniques for plant vacuole isolation have

been well established [20,21]. These isolated vacuolesare commonly surrounded by their tonoplasts withloosely adhering protoplasmic materials [19], includingthe P-Chls in the present study (Figure 2D). Accord-ingly, we further optimized the conditions for isolatingthe vacuoles with adhering P-Chls since the floatation ofP-Chls on a sucrose medium might effectively preventtheir contamination with CCCs in the pellet or disso-ciated C-Chls in the supernatant. As expected, our P-Chl preparation using this method barely containedLSU, which was found almost exclusively in the C-Chlfraction as revealed by silver staining and Western blotanalysis (Figure 8). The striking 10-fold difference inLSU level between the dimorphic chloroplasts is inagreement with our immunogold quantification analysisat the electron microscopic level showing that the LSUsignals were distributed between P-Chls and C-Chls at aratio of 1:20 (unpublished data). This observation sub-stantially contrasted the previous results obtained usingthe differential centrifugation method [10]. In theirstudy, Western blot analysis revealed the immunoreac-tive band intensities of LSU for the P-Chl and C-Chlfractions at a ratio of 1: 3.5 [10], suggesting an impurepreparation of P-Chls. Two possible sources of contami-nation in their P-Chl preparation with LSU might be, asdiscussed earlier, that the P-Chl fraction was easily con-taminated with unpelleted CCCs or dissociated C-Chlsusing the differential centrifugation method. Also, theC3-like chloroplasts from immature chlorenchyma cellswere inseparable from the P-Chls because neither typeof chloroplasts forms intracellular clumping. On theother hand, since vacuoles were not prominently foundin young chlorenchyma cells [11], our preparation of P-Chls using the vacuole-based method was not prone tocontamination with C3-like chloroplasts, as indicated bythe low level of LSU (Figure 8). Moreover, the unequaldistribution of proteins associated with the photosys-tems (PsbO and PsaD) in the two chloroplast fractionspresented here also supports earlier ultrastructuralresults showing that chloroplasts in the PCC have alower granal index than those in the CCC [7].

Potential applicationsWith the development of the current method to isolatetwo homogenous populations of dimorphic chloroplastsfrom B. sinuspersici, research can be carried out towarda thorough understanding of the single-cell C4 mechan-ism. Enzymology of the C4 photosynthetic and photore-piratory pathways can be explicitly characterized. Giventhe different subsets of enzymes in the two types ofchloroplasts, the possibility of differential protein importcan be evaluated by in vitro import assays and biochem-ical characterization of the protein import receptors atthe chloroplast envelope using the isolated dimorphic


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chloroplasts. Recently, high-throughput proteomic ana-lyses of the stroma [22] and membrane fractions [23] ofthe purified mesophyll and bundle sheath cells frommaize leaves have provided interesting insights into thedifferent roles of dimorphic chloroplasts in C4 photo-synthesis as well as other plastid functions. Similar pro-teomic analysis of the isolated dimorphic chloroplastsfrom B. sinuspersici might allow an informative compar-ison of their respective roles in single-cell C4 photo-synthesis and other physiological metabolism. Onanother perspective, the concomitant isolation of intactvacuoles using the current method might also be useful.For instances, one might further rule out the possibilityof crassulacean acid metabolism (CAM) in the single-cell C4 species by characterization of the metabolitecontents in the vacuoles. Alternatively, the regulation ofphotosynthetic enzymes in this single-cell C4 speciesmight be studied in relation to their turnovers invacuoles through the autophagic pathway.

ConclusionIn conclusion, we have established a reliable method forisolating and purifying the dimorphic chloroplasts from asingle-cell C4 species, B. sinuspersici. Following the hypo-osmotic lysis of isolated protoplasts under our definedconditions, one type of chloroplasts can be easily obtainedfrom the discrete subcellular structures in the pellet frac-tion after low-speed centrifugation, whereas another typeof chloroplasts along with vacuoles can be purified by floa-tation on a sucrose medium using the intact vacuoles ascarriers. The purity and homogeneity of the dimorphicchloroplast preparations were evident from the electro-pheretic analyses of their respective protein markers. Thistechnique can be applied in various experiments to unra-vel the novel photosynthetic pathways and other physiolo-gical aspects in the single-cell C4 model.

MethodsIsolation of chlorenchyma protoplasts from B. SinuspersicileavesB. sinuspersici plants were propagated asexually by vege-tative cuttings essentially as described [12]. Isolation ofleaf-derived chlorenchyma protoplasts from B. sinusper-sici has been optimized previously [12]. Briefly, 20mature leaves (> 2 cm in length) were freshly harvestedfrom 3- to 4-month-old vegetatively propagated plantsand chlorenchyma cells were isolated from the leaves incell-stabilizing (CS) buffer [0.7 M sucrose, 25 mMHEPES-KOH (pH 6.5), 5 mM KCl and 1 mM CaCl2] bygentle pressing using a mortar and pestle. The chlor-enchyma cells were collected on a piece of 40-μm nylonmesh filter (Sefar America Inc., USA) with a stack ofabsorbent paper underneath for removal of the buffersolution. The isolated cells were gently shaken off from

the nylon mesh filter into 5 mL of enzyme solution [CSbuffer with 1.5% (w/v) cellulase Onozuka R10 (YakultHonsha Co. Ltd., Tokyo, Japan) and 0.1% (w/v) bovineserum albumin (Sigma-Aldrich, cat. no. A7030)] andincubated at 25°C in the dark without shaking. The pro-gress of cellulase treatment was monitored under lightmicroscopy such that the appearance of round-shapedprotoplasts indicates complete cell wall digestion, whichwas normally achieved within 4 h. The protoplast-wash-ing steps were performed by floating the cells on top ofthe supernatant after centrifugation at 100g for 2 minand gently aspirating them using a micropipette. Ahomogenous population of chlorenchyma protoplastswas obtained after washing in 5 mL of CS buffer twice.The viability of the isolated protoplasts was assessed byvital staining and the protoplast yield was determinedusing a Neubauer-Levy haemocytometer (Hausser Scien-tific, Horsham, PA, USA).

Separation of dimorphic chloroplasts from Osmoticallylysed protoplastsThe isolated protoplasts were washed once in wash buf-fer [0.7 M sucrose, 25 mM HEPES-KOH (pH 6.5),5 mM KCl] and resuspended in an appropriate volumeof wash buffer to obtain a cell density of 105 protoplastsmL-1. All subsequent steps were carried out at 0~4°Cunless otherwise specified. To separate the P-Chls andC-Chls by the hypo-osmotic shock method, 500 μL ofprotoplast suspension were prechilled on ice for 5 minand then diluted with 1.25 mL of dilution buffer [25mM HEPES-KOH (pH 6.5), 5 mM KCl, 2 mM EDTA],which had been equilibrated at room temperature, toobtain a final sucrose concentration of 0.2 M. Thediluted protoplasts were gently mixed by inverting thetube several times and incubated at room temperaturefor 2 min. Following the osmotic lysis of protoplasts, thesamples were immediately transferred on ice for 10 minwith minimal agitation. During this 10-min incubationon ice, the released CCCs were settled to the bottom ofthe tube while the intact vacuoles carrying P-Chlsfloated to the top of the medium. The sedimentation ofCCCs and the floating of vacuoles with P-Chls wereseparated by centrifugation at 100g for 4 min in aswinging-bucket rotor (A-8-11 for Eppendorf centrifuge5417R; Eppendorf Canada Ltd., Mississauga, Canada).The floating layer of vacuoles with P-Chls was carefullytransferred to a fresh microfuge tube and the aqueousmedium was aspirated off with a micropipette withoutdisturbing the pellet containing CCCs. The CCC pelletwas washed once in 2 mL of CS buffer by gently tappingwith finger, centrifuged at 100g for 2 min, and resus-pended in 300 μL of dilution buffer. The CCCs werethen dissociated by pipetting 10 times up and downusing a narrow bore 200-μL micropipette tip. To further


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purify C-Chls from the other organelles in CCCs, thedissociated CCCs were subjected to Percoll gradientcentrifugation as described below. Similar Percoll gradi-ents were used to isolate the loosely associated P-Chlsfrom the vacuole surfaces.

Percoll gradient purification of chloroplastsThe chloroplast purification procedures were modifiedaccording to Smith et al. [24]. Briefly, up to 1 mL of chlor-oplast suspension was layered on a 40%/85% Percoll stepgradient consisting of an upper 500-μL Percoll solution[40% (v/v) Percoll, 50 mM HEPES-KOH (pH 7.3),330 mM sorbitol, 1 mM MgCl2, 1 mM MnCl2 and 2 mMEDTA] and a lower 500-μL Percoll solution [85% (v/v)Percoll, 50 mM HEPES-KOH (pH 7.3) and 330 mM sorbi-tol]. The gradient was centrifuged at 2,500g, 4°C for10 min in a swinging-bucket rotor (A-8-11 for Eppendorfcentrifuge 5417R; Eppendorf Canada Ltd., Mississauga,Canada). The intact chloroplasts at the 40%/85% interfaceof the Percoll gradient were aspirated and diluted with 6volumes of ice-cold HS buffer [50 mM HEPES-KOH (pH8.0) and 330 mM sorbitol]. The chloroplasts were pelletedby centrifugation at 750g, 4°C for 5 min and resuspendedin 50 μL of ice-cold HS buffer.

Isolation of total chloroplasts from chlorenchymaprotoplastsThe procedures for assembling a protoplast-rupturingdevice were modified according to the method describedby Smith et al. [24]. Briefly, the needle-fitting end of a 1-mL syringe barrel and the top part of a 500-μL microtubewere cut off to form a hollow tube and a slightly wideradaptor ring, respectively. A piece of 10-μm nylon meshfilter (Spectrum Lab Inc., Rancho Dominguez, CA, USA)was fitted against the cut end of the hollow tube and heldin place using the adaptor ring. All subsequent stepswere carried out at ~4°C. The isolated protoplasts wereresuspended in an appropriate volume of protoplastbreakage buffer [20 mM tricine-KOH (pH 8.4), 330 mMsorbitol, 5 mM EDTA, 5 mM EGTA, 10 mM NaHCO3]to obtain a cell density of 4 × 105 protoplasts mL-1. Fivehundred microliters of protoplast suspension were trans-ferred into the protoplast rupturing device and the plun-ger was inserted to force the suspension through themesh. An equal volume of protoplast breakage buffer wasthen added to the protoplast-rupturing device for com-pleting the breakage of residual protoplasts and CCCs onthe mesh. To isolate total chloroplasts from the celllysate, the pooled filtrates were subjected to Percoll gra-dient centrifugation as described above.

SDS-PAGE and western blot analysisThe isolated chloroplasts were resuspended in solubili-zation buffer [50 mM phosphate buffer (pH 7.2), 1% (w/

v) SDS]. The solubilized proteins were quantified usingthe bicinchoninic acid protein assay kit (Pierce, Rock-ford, IL, USA) according to the manufacturer’s instruc-tions. Two micrograms of proteins were separated onSDS/12% PAGE. The resolved proteins were visualizedby silver staining as described previously [25] or electro-blotted onto nitrocellulose membranes, which wereprobed with primary antibodies against PPDK (1:4,000;courtesy of Dr. Chris Chastain), LSU (1:5,000; Agrisera,Vannas, Sweden, cat. no. AS03-037), PsbO (1:5,000;courtesy of Dr. Marilyn Griffith), cytochrome f (1:1,000;Agrisera, Vannas, Sweden, cat. no. AS08-306) or PsaD(1:500; Agrisera, Vannas, Sweden, cat. no. AS09-461),followed by a horseradish peroxidase-conjugated anti-rabbit secondary antibody (1:10,000; Sigma-Aldrich, cat.no. A6154). The signals were visualized by incubatingthe probed membranes with Amersham ECL-Plus solu-tion (GE Healthcare, cat. no. RPN2132) and exposingthe membranes to Amersham Hyperfilm ECL films (GEHealthcare, cat. no. 28-9068-39), which were developedusing a CP1000 Agfa photodeveloper (AGFA, Ontario,Canada), digitized and processed using Adobe Photo-shop CS (Adobe Systems Inc., Seattle, WA, USA). Theintensities of immunoreactive bands were densitometri-cally quantified using the gel-analyzer function of Ima-geJ software v.1.46 (National Institutes of Health, USA).

Cytochemical stainingFor vital staining, 200 μL of protoplasts were incubatedwith 4 μL of 0.2% (w/v) fluorescein diacetate [Sigma-Aldrich, cat. no. F5502; prepared in 100% (v/v) acetone]at room temperature for 15 min and washed twice with200 μL of CS buffer. For vacuolar staining, 200 μL ofprotoplasts were centrifuged at 100g for 2 min and thefloating protoplasts were collected and resuspended in200 μL of acidic staining buffer [0.7 M sucrose, 50 mMsodium citrate (pH 4), 5 mM KCl and 1 mM CaCl2].The protoplasts were stained with 1 μL of 10 mMCDCFDA [Invitrogen, cat. no. C-369] at room tempera-ture for 15 min and washed twice with 200 μL of acidicstaining buffer. The stained protoplasts were observedunder epifluorescence microscopy.

Epifluorescence and light microscopyFifty microliters of cytochemically stained protoplastswere examined and imaged in flat-bottomed depressionslides under an epifluorescence microscope (Zeiss AxioImager D1) equipped with a Zeiss AxioCam MRm cam-era. Leaf samples were fixed, dehydrated and embeddedin London Resin White (Electron Microscopy Sciences)acrylic resin as previously described [9]. Semi-thin (1μm) sections were stained with 1% (w/v) toluidine blueO and observed under light microscopy. Bright fieldmicrographs were acquired using a Zeiss Axiophot


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microscope (Carl Zeiss Inc., Germany) equipped with aQ-Imaging digital camera (Quorum Technologies Inc.,Guelph, Canada). All images were processed and com-posed using Adobe Photoshop CS (Adobe Systems Inc.,Seattle, WA, USA). Representative images were pre-sented after similar results were obtained from at least 3independent experiments.

Additional material

Additional file 1: An AVI movie showing bursting of protoplastsand their vacuoles at 0.1 M sucrose. This movie was made from thetime-lapse images of protoplasts subjected to osmotic swelling at 0.1 Msucrose. Representative images are shown in Figure 3A.

Additional file 2: An AVI movie showing protoplast lysis andformation of vacuole-derived vesicles at 0.2 M sucrose. This moviewas made from the time-lapse images of protoplasts subjected toosmotic swelling at 0.2 M sucrose. Representative images are shown inFigure 3B.

Additional file 3: An AVI movie showing osmotic swelling ofprotoplasts and pinching-off of vacuoles at 0.5 M sucrose. Thismovie was made from the time-lapse images of protoplasts subjected toosmotic swelling at 0.5 M sucrose. Representative images are shown inFigure 3C.

AcknowledgementsThis research was supported by the Natural Sciences and EngineeringResearch Council (NSERC) of Canada Research Grants and the University ofWaterloo Start-Up Fund to S.D.X.C. S.-C.L. received additional financialsupport from Ontario Graduate Scholarship (Government of Ontario, Canada)and President’s Graduate Scholarship (University of Waterloo, Canada). Theauthors gratefully acknowledge Dr. Chris Chastain (Minnesota StateUniversity Moorhead, MN, USA) for providing the anti-maize PPDK antibody.

Authors’ contributionsSCL designed and performed the experiments, analyzed the data, anddrafted the manuscript. MY performed the electrophoretic analyses. SDXCconceived the study and provided supervision and critical revision of themanuscript. All authors read and approved the final manuscript.

Competing interestsThe authors declare that they have no competing interests.

Received: 16 January 2012 Accepted: 6 March 2012Published: 6 March 2012

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doi:10.1186/1746-4811-8-8Cite this article as: Lung et al.: Isolation of dimorphic chloroplasts fromthe single-cell C4 species Bienertia sinuspersici. Plant Methods 2012 8:8.


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