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Physiology and Development |
2Laboratory of Anatomy and Morphology, V. L. Komarov Botanical Institute of Russian Academy of Sciences, Prof. Popov Street 2, 197376, St. Petersburg, Russia; 3School of Biological Sciences, Washington State University, Pullman, Washington 99164-4236 USA; 4Department of Baking Technology, Urals State Economic University, Ekaterinburg, Russia
Received for publication March 20, 2003. Accepted for publication July 11, 2003.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Key Words: anatomy • Borszczowia aralocaspica • C4 photosynthesis • C4 plants • Chenopodiaceae • development • immunolocalization • ultrastructure
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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; Kanai and Edwards, 1999
).
In a surprising challenge to this paradigm of C4 mechanism, it has recently been shown that Kranz anatomy, consisting of two photosynthetic cell types, is not required for function of C4 photosynthesis in terrestrial plants. Borszczowia aralocaspica Bunge and Bienertia cycloptera Bunge, two succulent species in the family Chenopodiaceae, have C4/crassulacean acid metabolism (CAM) type carbon isotope composition; they lack Kranz anatomy and have unusual chlorenchyma not previously reported in C4, C3, or CAM plants (Freitag and Stichler, 2000
, 2002
). Studies on biochemistry and immunolocalization of photosynthetic enzymes show that mature leaves of these species have dimorphic chloroplasts, one specialized for supporting fixation of atmospheric CO2 in the C4 cycle and the other specialized for donation of atmospheric CO2 from C4 acids to the C3 cycle (Voznesenskaya et al., 2001b
, 2002
). The two species have very different and fascinating solutions to the spatial compartmentation, with Borszczowia partitioning the functions in opposite ends of elongated cells and Bienertia partitioning these functions into peripheral and central cytoplasmic compartments of shorter and wider cells. These species are unprecedented examples of not only the occurrence of dimorphic chloroplasts within a single photosynthetic cell type in terrestrial plants, but also the ability of plant cells to create highly complex biochemical polarization.
The exquisite structural and biochemical polarization exhibited by these two species raises a number of questions about how this is developed. Taking only the chloroplasts as an example, a number of possibilities can be proposed, from initial presence of two types of chloroplasts (genetically) in young cells to differentiation of two biochemical forms followed by partitioning or partitioning followed by biochemical differentiation. Similar questions apply to the mitochondria. In this paper, we present our study on development of C4 features in leaf chlorenchyma of B. aralocaspica. It is interesting to note here that the spatial separation of dimorphic chloroplasts in mature, elongated chlorenchyma cells of B. aralocaspica is similar to that in species having Salsoloid and Suaedoid-type Kranz leaf anatomy (Freitag and Stichler, 2000
; Voznesenskaya et al., 2001b
). In theory, these elongated cells in mature leaves of B. aralocaspica could originate from Kranz type anatomy by degradation of adjoining cell walls between photosynthetic cells (i.e., conversion from Kranz to a single photosynthetic cell as leaves mature). Programmed fusion of cells by dissolution of end walls is known to occur among certain cell types such as articulated laticifers (Dickison, 2000
). In the present study, the development of the B. aralocaspica C4 cells is characterized with respect to cell size, the position of organelles in the cytoplasm, the timing of the occurrence of dimorphic chloroplasts, and expression of photosynthetic proteins and carbon fixation. We show that single-cell C4 in this remarkable species is achieved through complex differentiation during development of individual cells and probably not through genetically diverse organelles.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Light and electron microscopy
Samples were taken for microscopy, quantitative anatomy, immunolocalization, and starch analysis. For each developmental stage a minimum of four samples were taken from the middle part of 2–3 leaves. Samples for ultrastructural study were fixed at 4°C in 2% (v/v) paraformaldehyde and 2% (v/v) glutaraldehyde in 0.1 mol/L phosphate buffer (pH 7.2), postfixed in 4% (m/v) OsO4, and then, after a standard acetone dehydration procedure, embedded in Spurr's epoxy resin. Cross sections were made on a Reichert Ultracut R ultramicrotome (Reichert-Jung, Heidelberg, Germany). For light microscopy, semi-thin sections were stained with 1% (m/v) toluidine blue O in 1% (m/v) Na2B4O7. Ultra-thin sections were stained for transmission electron microscopy with 2% (m/v) lead citrate or 1 : 2 dilution of 1% (m/v) KMnO4 : 4% (m/v) uranyl acetate. Hitachi H-600 (Hitachi Scientific Instruments, Nissei Sangyo America, Mowatain View, California, USA) and JEM-1200 EX (JEOL USA, Peabody, Massachusetts, USA) transmission electron microscopes were used for observation and photography. The lengths of appressed and nonappressed thylakoid membranes (including both intergranal and end granal thylakoid membranes) were measured with a curvimeter on at least 10 median sections of chloroplasts. The granal index was calculated as the length of all appressed thylakoid membranes as a percentage of the total length of all thylakoid membranes in a chloroplast. The thylakoid density was calculated as the length of thylakoid membranes (in micrometers) per 1 µm2 of chloroplast stroma area (analyzed for the total area of the chloroplast excluding starch grains). The chloroplast area in sections was estimated with an image analysis program (UTHSCSA, Image Tool for Windows, version 3.00, University of Texas Health Science Center, San Antonio, Texas, USA) on the same micrographs.
In situ immunolocalization samples and reagents
Leaf samples were fixed at 4°C in 2% (v/v) paraformaldehyde and 1.25% (v/v) glutaraldehyde in 0.05 mol/L phosphate buffer, pH 7.2. The samples were dehydrated with a graded ethanol series and embedded in London Resin White (LR White, Electron Microscopy Sciences, Fort Washington, Pennsylvania, USA) acrylic resin. Antibodies used (all raised in rabbit) were anti-spinach rubisco (LSU) IgG (courtesy of B. McFadden, Washington State University), commercially available anti-maize phosphoenolpyruvate carboxylase (PEPC) IgG (Chemicon, Temecula, California, USA), anti-Amaranthus hypochondriacus L. var 1023 mitochondrial NAD- malic enzyme (NAD-ME) IgG, which was prepared against the 65 KD 
subunit (courtesy of J. Berry, SUNY Buffalo: Long, Wang, and Berry, 1994
), anti-Zea mays L. 62 KD NADP-malic enzyme (NADP-ME) IgG (courtesy of C. Andreo, University of Rosario, Argentina: Maurino, Drincovich, and Andreo, 1996
), anti-Zea mays pyruvate,Pi dikinase (PPDK) IgG (courtesy of T. Sugiyama, Plant Science Center, RIKEN, Japan), anti-Pisum sativum L. glycine-decarboxylase (courtesy of D. Oliver, Iowa State University), and anti-Spinacea oleracea L. ADPG pyrophosphorylase (AGPase) (courtesy of T. Okita, Washington State University).
Light microscopy immunolocalization observations
Cross sections, 0.8–1 µm thick, were dried from a drop of water onto gelatin-coated slides and blocked for 1 h with TBST+BSA (10 mmol/L Tris-HCl, 150 mmol/L NaCl, 0.1% v/v Tween 20, 1% m/v bovine serum albumin, pH 7.2). They were then incubated for 3 h with either the preimmune serum diluted in TBST+BSA (1 : 100), anti-rubisco (1 : 500 dilution), anti-PEPC (1 : 200 dilution), anti-NAD-ME (1 : 100), anti-NADP-ME (1 : 20), anti-PPDK (1 : 100), anti-AGPase (1 : 20) or anti-glycine decarboxylase (GDC)(1 : 600) antibodies. The slides were washed with TBST+BSA and then treated for 1 h with protein A-gold (10 nm particles diluted 1 : 100 with TBST + BSA). After washing, the sections were exposed to a silver enhancement reagent for 20 min according to the manufacturer's directions (Amersham, Arlington Heights, Illinois, USA), stained with 0.5% (m/v) Safranin O, and imaged in a reflected/transmitted mode using a BioRad 1024 confocal system with Nikon Eclipse TE 300 inverted microscope (BioRad, Hercules, California, USA) and Lasergraph image program 3.10. The background labeling with pre-immune serum was very low although some infrequent labeling occurs in areas where the sections were wrinkled due to trapping of antibodies/label (results not shown).
Transmission electron microscopy immunolocalization
Thin sections on coated nickel grids were incubated for 1 h in TBST+BSA to block nonspecific protein binding on the sections. They were then incubated for 3 h with the preimmune serum diluted in TBST + BSA, or anti-PEPC (1 : 50 dilution), anti-NAD-ME (1 : 50), anti-NADP-ME (1 : 20), or anti-GDC (1 : 400) antibodies. After washing with TBST plus BSA, the sections were incubated for 1 h with Protein A-gold (10 nm) diluted 1 : 100 with TBST/BSA. The sections were washed sequentially with TBST plus BSA, TBST, and distilled water, and then post-stained with a 1 : 4 dilution of 1% (m/v) potassium permanganate and 2% (m/v) uranyl acetate. Images were collected using a JEM-1200 EX transmission electron microscope.
Staining for polysaccharides
Sections, 0.8–1 µm thick, were made from the same samples used for immunolocalization, dried onto gelatin coated slides, incubated in periodic acid (1% m/v) for 30 min, washed, and then incubated with Schiff's reagent (Sigma, St. Louis, Missouri, USA) for 1 h. After rinsing, the sections were analyzed by light microscopy.
Enzyme extraction and assay
Enzymes were extracted from illuminated leaves harvested during the photoperiod. Leaves were frozen in liquid N2 until analyzed. The material was ground in a Ten-Broeck grinder (Willmad-Labglass, Buena, New Jersey, USA) with cold extraction buffer (1 mL per 50 mg fresh mass) for about 30 s. The extraction buffer contained 100 mmol/L HEPES-KOH (N-[2-Hydroxyethyl]piperazine-N'-[2-ethanesulfonic acid/ potassium hydroxide) (pH 7.5), 10 mmol/L MgCl2, 5 mmol/L DTT, 2 mmol/L EDTA and 2% PVPP. The extract was then centrifuged in an Eppendorf microcentrifuge (Brinkman Instruments, Westbury, New York, USA) (2 min, 1466 rad/s). For assay of rubisco the enzyme was preincubated at room temperature for 12 min in the presence of 10 mmol/L bicarbonate and 10 mmol/L MgCl2 to obtain maximum activation. For rubisco and PEPC assay, the reaction was started by adding 50 µL of extract to 450 µL of media with 10 mmol/L NaH14CO3 containing 1 mmol/L RuBP (ribulose-1,5-bisphosphate) for rubisco assay or 5 mmol/L PEP in case of PEPC assay. The reaction was stopped after 30 s by addition of 200 µL 1 N HCl + 4 N Formic acid, and acid-stable radioactivity was determined with a scintillation counter. The PPDK activity was measured spectrophotometrically in a 1-mL reaction mixture. The assay medium for PPDK contained 50 mmol/L HEPES/KOH (pH 8.0), 10 mmol/L MgCl2, 3 mmol/L DTT, 0.1 mmol/L EDTA, 10 mmol/L NaHCO3, 1.25 mmol/L pyruvate, 0.2 mmol/L NADH, 2.5 mmol/L KH2PO4, 1.25 mmol/L ATP, 10 units malate dehydrogenase, and 1 unit PEPC. The reaction was started by addition of 50 µL of extract.
Exposure of leaves to 14CO2
Rates of 14CO2 fixation with excised leaves
An intact plant of Borszczowia aralocaspica was incubated in a growth chamber at 800 photosynthetic photon flux density (PPFD). For determining rates of CO2 fixation, different aged leaves were excised and after measurement of fresh mass they were placed in a glass vial with a thin layer of water to provide high humidity. The leaves were preincubated for 5 min at 1500 PPFD (combination of white light from a Schott lamp with a heat filter and and a red, light-emitting diode lamp). The plant tissue was then exposed to 1.8% 14CO2 gas (specific activity, 13 mCi/mol) for 5 min at 1500 PPFD. The reaction was terminated by plunging the tissue into hot ethanol, and disintegrations per minute incorporated per unit chlorophyll were determined.
Determination of photosynthetic products
Leaves were excised from the plant 4–6 h into the light period. They were placed immediately into a glass vial (final volume 30 mL), with the base of the leaves submerged in distilled water, and they were preilluminated for 2 min with 950 µmol photosynthetic quanta · m–2 · s–1 from two sides (two 150-W halogen lamps, type ELD, Atlanta Light Bulbs, Tucker, Georgia, USA) at ca. 30°C. The lights were filtered through 5 cm of water in a glass container to avoid excess heat. Prior to introduction of 14CO2 into the vial, the plant material was flushed quickly with a stream of humidified, CO2-free air. Before initiating the experiment, 14CO2 was generated in a separate vial by addition of NaH14CO3 to HCl. Immediately after flushing the plant cuvette with CO2-free air, 2 mL of 14CO2 gas, containing 4 µC of 14CO2, was injected, resulting in a final CO2 concentration of 356 µL/L.
After giving a short pulse of 14CO2, the plant material was killed by plunging into boiling 80% ethanol (v/v). Tissue was boiled an additional 2–3 min, ground thoroughly with a mortar and pestle with the addition of small amount of acid-washed sand, and extracted again with 96, 80, 60, 40% ethanol and twice with water. All extracts were pooled and concentrated to approximately 1 mL. The leaf extract was partitioned with CHCl3. Separation and identification of the labeled photosynthetic products were accomplished using two-dimensional thin-layer electrophoresis and chromatography methods (Shurmann, 1969
; Moore and Seemann, 1990
). Recovery of radioactivity from the plates was >90%.
13C carbon isotope determination
Carbon isotope fractionation values were determined on leaf samples taken from plants grown in growth chambers, using a standard procedure relative to Pee Dee Belemnite (PDB) limestone as the carbon isotope standard (Bender et al., 1973
).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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As shown in Fig. 2C, in mature leaves (ca. 3 cm long) there is only one layer of chlorenchyma cells. These large, elongated cells have a large central vacuole and have an obvious difference in thickness of the cytoplasmic layer at opposite ends of the cells (Fig. 2C). In the distal part of the cell (away from the center of the leaf), the layer of cytoplasm is rather thin and contains narrow, elongated chloroplasts. The chloroplasts of the distal end are preferentially distributed along cell walls exposed to intercellular air spaces. The proximal ends of these cells (next to the water storage cells and peripheral vascular bundles) are tightly packed. They are devoid of intercellular air spaces along their radial walls; however, there are some intercellular air spaces between the ends of the cells and the water-storage or vascular tissues (Fig. 2C). The layer of cytoplasm at the proximal end of these cells is much thicker than in the distal end and contains abundant organelles distributed along cell walls without any relationship to the position of air spaces. In these mature cells, starch is abundant in chloroplasts in the proximal part of the cell and practically absent in the chloroplasts in the distal part (Fig. 2F). An interesting feature of the mature cells is the absence of any organelles in the very thin layer of cytoplasm at the interface between proximal and distal parts (Fig. 2C). Usually, the nuclei are located towards the proximal part of the cell at the border of the zone lacking organelles.
Electron microscopy
In chlorenchyma cells of young leaves, a few small chloroplasts (see Table 1 for sizes) are evenly distributed in the rather thick cytoplasmic layer (Fig. 3A); they have a poorly developed internal structure consisting of small grana of 2–5 thylakoids interconnected with short intergranal thylakoids (Fig. 3B–C). Chloroplasts throughout the cell have nearly the same structure and size (Table 1), and they contain protein crystals and/or starch grains. Mitochondria, which are rather small in young tissue (Table 1), are also evenly distributed throughout the cell cytoplasm, and they have an electron-translucent matrix in the center with a few small cristae around their periphery.
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In mature leaves, chlorenchyma cells are very large (Table 1) and are differentiated clearly into two different compartments, separated by the thin cytoplasmic area without organelles (Fig. 3G). The distal parts of these cells have a rather thin layer of peripheral cytoplasm and contain elongated, nearly agranal chloroplasts (Fig. 3H), while the cytoplasmic layer at the proximal end is thicker and contains granal chloroplasts (Fig. 3I). The granal index was 22% for distal chloroplasts and 49% for proximal chloroplasts with a ratio of appressed to nonappressed thylakoids membranes of 0.29 and 0.95, respectively. The two chloroplast types have nearly the same total density of thylakoids (30.4 for distal and 27.2 for proximal), but in the distal chloroplasts the density of appressed thylakoids is about half that of the proximal chloroplasts, while the density of nonappressed thylakoids in distal chloroplasts is 1.7 times higher. Interestingly, chloroplasts at the medial region of the cell at the transition point where the thin cytoplasm interfaces with the distal region usually have a level of grana development intermediate between those characteristic for distal and proximal chloroplasts; the granal index of these chloroplasts was 35%, with a ratio of appressed to nonappressed thylakoids membranes of 0.55. Both distal and proximal chloroplasts in mature chlorenchyma cells have nearly the same length but proximal chloroplasts had a greater width (length of their short axis) (Table 1). Starch grains are much more abundant in proximal chloroplasts (Fig. 3I), while protein crystals occur in both types of chloroplasts (not shown). Numerous, rather large mitochondria (see Table 1 for sizes) having intensively developed tubular cristae in a dense matrix, are located in the proximal part of the cell. The mitochondria are positioned to the outside of the cytoplasmic layer close to the cell wall, between the chloroplasts that are adjacent to the vacuole (Fig. 3I). The structure and thickness (about 0.1 µm) of cell walls in different parts of the chlorenchyma cell is uniform.
Immunocytochemistry
In young leaves, which have evenly distributed chloroplasts in the chlorenchyma cells, labeling for rubisco is present in all chloroplasts including those in chlorenchyma, hypodermal, epidermal, and water-storage tissues (Fig. 4A). The labeling for PEPC is distributed throughout the cytoplasm of chlorenchyma cell (Fig. 4B). There is low labeling for PPDK (Fig. 4C), and little or no labeling for AGPase or GDC (results not shown) in young leaves.
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In the photosynthetically mature leaves, practically all labeling for rubisco (Fig. 4G) is concentrated in the chloroplasts of the proximal part of the chlorenchyma cell, with some labeling in the few chloroplasts in water-storage cells, but no labeling for rubisco in chloroplasts of the distal part of the cell. The PEPC in mature leaves occurred throughout the whole chlorenchyma cell cytoplasm (Fig. 4H). Electron microscopy observations show that labeling for this enzyme is located in the cytosol, both in the distal and proximal ends of the cell (Fig. 5A–B). In mature leaves, PPDK is strongly compartmentalized to chloroplasts in the distal part of the chlorenchyma cells with minor labeling in chloroplasts of the proximal part (Fig. 4I). The study of mature leaves showed that GDC is localized in the proximal part of chlorenchyma cells (Fig. 4J), and electron microscopy observations revealed that it is confined to the mitochondria (Fig. 5C). Labeling for NAD-ME is also located proximally in chlorenchyma cells (result not shown) and confined to the mitochondria (Fig. 5D). There was no labeling with NADP-ME (results not shown). The labeling for AGPase is also mostly expressed in the proximal part of chlorenchyma cells, with some labeling in the hypodermal and water-storage tissues and occasionally in phloem tissues, where it is obviously confined to the chloroplasts (Fig. 4K).
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13C carbon isotope determination13C value that range from –10.31 to –12.37
. Also, two other samples of leaves ca. 0.8 cm to ca. 1 cm in length had an average carbon isotope value of –11.15 (not shown in table). Thus, all values are in the range typical of C4 plants.
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Rates of CO2 fixation in excised leaves measured under high CO2 and high light progressively increased from young to intermediate to mature (Table 2). Analysis of photosynthetic products in a 14CO2 pulse-12CO2 chase experiment on mature leaves of B. aralocaspica showed that the primary initial products after 8 s pulse with 14CO2 were malate and aspartate (Table 3). During a 15-s pulse followed by chase for 30 s and 120 s there was progressive decrease of label in C4 acids and increase in label in other products. Comparative results are shown for two other species in subfamily Salsoloideae, a Kranz NAD-ME type C4 plant Salsola laricina Pall., and a C3 plant, Suaeda heterophylla (Kar. et Kir.) Bunge et Boiss, which had labeling patterns characteristic of C4 and C3 photosynthesis, respectively.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Ultrastructure
In developing chlorenchyma of young leaves, there is no selective differentiation, compartmentation or partitioning of organelles as seen in the mature chlorenchyma cells. All the chloroplasts in these young cells are structurally identical and have small grana with short intergranal thylakoids. At our intermediate sampling stage of leaf development, there were clear indications of the early stages of organelle partitioning. In some cells there was an obvious increase of the chloroplasts in the proximal end of the cell while in adjacent cells this process had not yet occurred. This is a further indication that chloroplasts are initially uniformly distributed in the cell and then partitioned as the cells mature. Also, in some cells there was evidence for increased partitioning of mitochondria towards the proximal end of the cell. At this intermediate sampling stage, the thin organelle-deficient cytoplasmic zone seen at the medial region between proximal and distal cytoplasmic compartments in the mature cells has not developed. In mature cells, there is a clear compartmentation of two types of chloroplasts, nearly agranal in the distal part of the cell and granal in the proximal part, with a distinct organelle-free zone between (also see Voznesenskaya et al., 2001b
). Thus, the developmental sequence analysis demonstrates that the two types of chloroplasts seen in the mature cells originate from one type of plastid.
Borszczowia aralocaspica is an NAD-ME type C4 species. In mature chlorenchyma cells of B. aralocaspica, the chloroplasts in the distal and proximal parts have structural characteristics like those in mesophyll (distal part) and bundle sheath cells (proximal part), respectively, of Kranz type C4 NAD-ME chenopods. Previous studies suggest that chloroplasts whose biochemistry demands low levels of reductant (NADPH) relative to ATP have lower grana development and lower PSII activity (Edwards and Walker, 1983
; Voznesenskaya et al., 1999
; Pyankov et al., 2000
). The proposed energetics of C4 photosynthesis in NAD-ME type species results in a higher demand for reductant in the bundle sheath cells, which have granal chloroplasts, than in mesophyll cells with grana deficient chloroplasts. The ultrastructural features of chloroplasts in the two regions of the NAD-ME single-cell C4 B. aralocaspica are consistent with what is known for the classical anatomical NAD-ME C4 systems and gives further evidence for the operation of this mechanism in a single cell.
Developmentally, B. aralocaspica shows a similar pattern of chloroplast differentiation in a single cell that has been shown for both cells of Kranz C4 systems. The transition from having one chloroplast type in young cells to developing dimorphic chloroplasts during cell maturation in B. aralocaspica mimics the process seen in the two cells of Kranz type C4 (Laetsch, 1969
; Brangeon, 1973
; Appiano et al., 1979
; Voznesenskaya, 1981
; Liu and Dengler, 1994
; Voznesenskaya et al., 2003
). It appears that the single-cell C4 system is following the same developmental programming as the dual cell system but within a single cell. This holds some important implications with respect to the evolution of the organelle differentiation and partitioning process.
An interesting feature of B. aralocaspica is that the chloroplasts that are located in an intermediate position between the proximal and distal ends of the cell have an intermediate level of grana development. The same character was found in Bienertia cycloptera, where some chloroplasts lying in an intermediate position inside cytoplasmic strands between the central and peripheral compartments have an intermediate level of grana stacking (Voznesenskaya et al., 2002
). A diffusible signal or gradient is evidently at work with respect to regulating organelle differentiation within the single-cell system. The absence of a distinct permeability barrier such as the wall in the Kranz system possibly results in "leaky" regulation. For the C4 mechanism to be successful in Kranz type C4 plants, the increased CO2 in the bundle sheath cell must be inhibited from diffusing back to the airspace or mesophyll. There is some evidence that the bundle sheath wall in Kranz systems may help limit CO2 diffusion back out of the bundle sheath cells. For example, it is well known that in many Kranz type C4 species, the bundle sheath cell walls are thickened relative to mesophyll cells and have a suberin layer in some C4 grasses (Carolin et al., 1973
; Hattersley and Browning, 1981
) and C4 sedges (Carolin et al., 1977
). Bundle sheath cell walls are also thick in some dicotyledonous plants, especially in some NADP-ME species, while in NAD-ME species these walls are usually thinner (Voznesenskaya and Gamaley, 1986
). Our study of chlorenchyma cells in developing B. aralocaspica leaves revealed neither major differences in cell wall structure in different parts of these cells nor the presence of any kind of suberin inclusions. However, several features of this cell could limit leakage of CO2 from sites of C4 acid donation to rubisco, including lack of intercellular air space between the chlorenchyma cells at the proximal ends, chloroplasts surrounding mitochondria where CO2 is released via NAD-ME and the long liquid phase diffusion path towards the distal part of the cell. The following calculation using the equation of Nobel (1991)
shows that development of elongated cells can provide significant diffusive resistance of CO2 from sites of C4 acid decarboxylation at the proximal ends.
x = mean diffusion path in cell, D = diffusion coefficient for CO2 in the cytosol (1 x 10–9 m2/s; Nobel, 1991
), and K = total inorganic carbon/inorganic carbon as CO2. The area of chlorenchyma exposed to intercellular air space (Ames) (Fig. 2C, F) is estimated to be 11-fold higher than Aleaf (exposed cell surface area was estimated assuming cylindrical cells with rounded ends). The mean length of the cells is 83 µm, and the liquid phase diffusion path from the proximal end to intercellular air space at the distal end is ca. 50 µm (x). Decarboxylation through NAD-ME releases CO2, and assuming this is the main form of inorganic carbon at the distal end of the cell, the value of K = 1 at 20°C. With these inputs the calculated diffusive resistance to CO2 in the liquid phase is 44.6 s/cm which is the same order of magnitude of the diffusive resistance in Amaranthus edulis Speg., a Kranz NAD-ME type C4 species (Kiirats et al., 2002
). By comparison, in C3 leaves diffusive resistance between the intercellular space and rubisco is ca. 1.4 s/cm (Nobel, 1991
).
In young and intermediate leaves, mitochondria in chlorenchyma cells are smaller than in mature leaves (see Table 1; usually the length of the short axis is the most reliable parameter, since in some cases a rather long mitochondrion profile was observed). In young leaves, these organelles have a translucent matrix in the center with only a few cristae on their periphery. Interestingly, in mature leaves of B. aralocaspica, large specialized mitochondria mostly have tubular cristae. In NAD-ME Kranz type C4 plants, a similar structure was observed for mitochondria in bundle sheath cells of Amaranthus, Portulaca oleracea L., and Mollugo cerviana (L.) Ser. (Laetsch, 1968
, 1971
), while some species of Atriplex, Suaeda, and Salsola have mitochondria with both tubular and lamellated types of cristae (Laetsch, 1968
; Downton, 1971
; Vosnesenskaja, 1976
; Voznesenskaya, 1976
; Shomer-Ilan et al. 1979
; Voznesenskaya and Gamaley, 1986
; Fisher et al., 1997
). In Bienertia cycloptera, another C4 species without Kranz anatomy, mitochondria also contained numerous well-developed lamellated cristae (Voznesenskaya et al., 2002
) as in C4 Suaeda species. Changes in mitochondria structure in B. aralocaspica during development may be related to increased mitochondrial transport and increased NAD-ME and glycine decarboxylase associated with function of the C4 cycle and photorespiration.
Immunolocalization
In young leaves of B. aralocaspica there was no selective compartmentation of rubisco. It was found in chloroplasts in various tissues including epidermal, hypodermal, water storage cells, and palisade-like chlorenchyma, with the latter having the largest number of chloroplasts. Development of selectivity in rubisco labeling in chloroplasts of chlorenchyma cells was correlated with structural differentiation of plastids and their positioning towards opposite ends of the cell. Evidence for increased expression of rubisco in proximal chloroplasts could be seen at an intermediate stage of leaf development. This occurred prior to observing any clear structural difference between chloroplasts located in different parts of the cell. Full biochemical compartmentation of rubisco, NAD-ME, and GDC was completed only together with structural differentiation of chloroplasts and mitochondria and their positioning in the chlorenchyma cell. Thus there is coupling between organelle partitioning and their biochemical differentiation.
The transition from rubisco being localized in all chloroplasts to selective localization of rubisco in one type of chloroplast in a single photosynthetic cell of B. aralocaspica is analogous to the C4 species Salsola richteri (Moq.) Kar. ex Litw., which has Salsoloid type Kranz anatomy. In the latter case, the enzyme was expressed in both bundle sheath and mesophyll chloroplasts in young leaves, with increasing preferential labeling of bundle sheath chloroplasts during development (5- and 8-d-old leaves) followed by complete compartmentation in bundle sheath chloroplasts of mature leaves (Voznesenskaya et al., 2003
). In mature leaves of B. aralocaspica, localization of rubisco in chloroplasts in the proximal part of the cell correlated with localization of starch and an enzyme required for starch biosynthesis (AGPase). This is consistent with studies on C4 chenopod species with Kranz anatomy, where there is co-occurrence of rubisco and starch in bundle sheath chloroplasts (Voznesenskaya et al., 1999
, 2003
).
The single-cell C4 plant B. aralocaspica and the Kranz type C4 Salsola richteri (both in family Chenopodiaceae, subfamily Salsoloideae), appear to be following a similar pattern of C4 development, since expression of certain photosynthetic enzymes in chloroplasts and mitochondria during development correlates with the differentiation in chloroplast and mitochondrial structure. In comparison, in Atriplex rosea L., with Atriplicoid type C4 Kranz anatomy (family Chenopodiaceae subfamily Chenopodioideae), bundle-sheath specific accumulation of rubisco occurs early in leaf development, just after the delimiting of bundle sheath tissue from the ground meristem and well before the final anatomical differentiation of both types of photosynthetic cells, and follows the order of vein formation (Dengler et al., 1995
). In developing amaranth leaves with Atriplicoid type of anatomy (family Amaranthaceae), it was shown that the C4 type expression of rubisco does not correlate with development of Kranz type anatomy, which occurs much earlier, but with the maturation of the smaller veins and sink-to-source transition. In the early stages of development rubisco protein is present in both mesophyll and bundle sheath cells in a C3-like pattern and becomes more localized to bundle sheath cells in a C4 pattern only at a later stage of development (Wang et al., 1992
; Ramsperger et al., 1996
). The transition from C3 to C4 mode of rubisco localization takes place in amaranth at the stage of sink-to-source transition that correlates with the beginning of photosynthesis and maturation of the peripheral veins (Wang et al., 1993
). Thus, during C4 development, chloroplasts undergo a C3 to C4 transition from a single chloroplast type containing rubisco to dimorphic chloroplasts; however, the timing of expression of certain photosynthetic enzymes during development of C4 photosynthesis in dicots does not follow a fixed pattern but shows some species-dependent variation.
In B. aralocaspica, PEPC is localized in the cytosol throughout chlorenchyma cell development. Even in the youngest leaves at the shoot apex (around 1 mm in length) it is clearly present (results not shown). Occasional labeling with PEPC antibody occurred in the vacuole; as previously discussed in studies with Salsola arbusculiformis Drob., this may occur due to association of the antibody with phenolic compounds (Voznesenskaya et al., 2001a
). The PPDK is specifically localized in distal chloroplasts in mature leaves and there was little reaction with the antibody in young leaves. Reactions with anti-GDC and anti-NAD-ME revealed their localization in the proximal part of the cell where specialized mitochondria are found in the mature leaf. In C3 plants, the expression of GDC, a photorespiratory enzyme, during development also coincides with development of photosynthesis. In etiolated cotyledons of sunflower (C3), GDC is light induced (Aaron and Edwards, 1980
) and in some C3 grasses like wheat, the activity of GDC increases fivefold during development (Tobin et al., 1988
; Rogers et al., 1991
; Bowsher and Tobin, 2001
). In general, the developmental expression of these enzymes in the single-cell C4 system tracks what has been seen for Kranz type systems.
Enzyme activity, 14CO2 fixation, products, and carbon isotope composition
The biochemical and physiological data support the developmental, structural, and immunological results that this species has a robust mechanism for partitioning organelles and enzymes in a manner that allows C4 carbon fixation to occur in a single cell. The activity of photosynthetic enzymes examined increased during development including PEPC and PPDK of the C4 cycle and rubisco of the C3 cycle. The carbon isotope values for young, intermediate, and mature leaves all indicate the carbon is derived from C4 photosynthesis, and these values are consistent with previous results on mature leaves from field and growth chamber experiments (Voznesenskaya et al., 2001b
). The products of CO2 fixation in a pulse-chase experiment with mature leaves indicate the occurrence of C4 photosynthesis, and tests for diurnal leaf acidity indicate no crassulacean acid metabolism. Based on these analyses, B. aralocaspica is essentially acquiring all its carbon through C4 photosynthesis. Very early in leaf development, when young leaves have a length about 10% of that of mature leaves, chloroplast differentiation and polarization has not occurred, so carbon fixed during photosynthesis may be less enriched in 13CO2 due to discrimination by rubisco. However, the carbon isotope composition of the very young leaves is C4 type; thus their initial development likely occurs mainly from carbon imported from mature leaves. An analogous situation may exist in Kranz type C4 plants during a C3 to C4 type transition as photosynthesis develops in very young tissue.
Borszczowia aralocaspica represents an unusual discovery of a mechanism that allows true C4 photosynthesis to occur without Kranz anatomy. This study demonstrates that the mature cell is not derived from degeneration of a wall between two initially separate cells and also that the dimorphic organelles originate from one common pool and require developmentally regulated biochemical differentiation. The structural, developmental, and biochemical regulation and coordination that must exist to accomplish formation of this amazing single-cell C4 system must be complex as it is occurring all in the same cell. Considering the absolute requirement for organelle partitioning in the single-cell C4 syndrome, the control of organelle movement and establishment of subcellular domains must utilize a precise and stable cellular process. Along with the partitioning of organelles, selective expression of organelle-encoded enzymes such as the large subunit of rubisco in one, but not both, compartments of the cell has to occur in order for the C4 pathway to operate properly. This is further complicated by the necessity for nuclear encoded enzymes to be accumulated in a specific subset of organelles, for example, rubisco small subunit vs. PPDK in the dimorphic chloroplasts. Further research is required to determine the mechanism of polar localization of organelles and enzyme expression and C4 function within this single photosynthetic cell.
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FOOTNOTES |
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5 E-mail: vfrances{at}mail.wsu.edu
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LITERATURE CITED |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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