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(American Journal of Botany. 2008;95:782-792.) doi: 10.3732/ajb.2007381 © 2008 Botanical Society of America, Inc. |
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Cell Biology |
2 Department of Botany, Miami University, Oxford, Ohio 45056 USA 3 Department of Cell Technology, Faculty of Engineering, Takushoku University, 815-1 Tate-machi, Hachioji, Tokyo, 193-0985, Japan 4 Department of Botany, Miami University, Hamilton, Ohio 45011 USA
Received for publication 21 November 2007. Accepted for publication 11 April 2008.
ABSTRACT
Cellular degeneration is essential for many developmental and stress acclimation processes. Undifferentiated parenchymatous cells in the central vascular cylinder of pea primary roots degenerate under hypoxic conditions created by flooding at temperatures >15°C, forming a long vascular cavity that seems to provide a conduit for longitudinal oxygen transport in the roots. We show that specific changes in the cell wall ultrastructure accompanied previously detected cytoplasmic and organellar degradation in the cavity-forming roots. The degenerating cells had thinner primary cell walls, less electron-dense middle lamellae, and less abundant cell wall homogalacturonans in altered patterns, compared to healthy cells of roots grown under cold, nonflooded conditions. Cellular breakdown and changes in wall ultrastructure, however, remained confined to cells within a 50-µm radius around the root center, even after full development of the cavity. Cells farther away maintained cellular integrity and had signs of wall synthesis, perhaps from tight regulation of wall metabolism over short distances. These observations suggest that the cell degeneration might involve programmed cell death. We also show that warm, nonflooded or cold, flooded conditions that typically do not induce vascular cavity formation can also induce variations in cell wall ultrastructure.
Key Words: cell wall ultrastructure • cellular breakdown • flooding and warm temperature response • homogalacturonan (HG) • Pisum sativum • primary root • programmed cell death (PCD) • vascular cavity
Cell death and degeneration are important parts of the plant life cycle and occur during many developmental processes such as aleurone degeneration, tracheary element differentiation, leaf sculpting, megaspore abortion, suspensor cell degeneration, abscission, and senescence of aged organs (Gray, 2004
). Cell death and degeneration also occur in response to stresses such as pathogen infection and changes in environmental conditions. Cell degeneration can be necrotic under severe stress and even lead to death of the entire plant, but it can also be part of adaptive responses to survive certain stresses. Such responses include rapid removal of infected cells to resist the spread of pathogen (the hypersensitive response), facultative aerenchyma formation in roots to increase oxygen transport during flooding, and some less pronounced changes in cellular ultrastructure to regulate metabolic and other adaptive cellular activities (Gray, 2004).
The anatomy of the vascular cylinder of the primary roots of pea (Pisum sativum L. cv. Alaska) varies depending on the developmental status and environmental conditions (Lu et al., 1991; Niki et al., 1995; Gladish and Niki, 2000; Niki and Gladish, 2001). Roots grown at optimum conditions (temperatures 
15°C, moist growth medium) have immature parenchymatous cells at the center of the vascular cylinder up to 15–20 cm away from the root tip (Fig. 1A). Beyond this point, the parenchymatous cells start differentiating into late-maturing metaxylem tracheary elements (LMTEs; Lu et al. 1991). In roots grown at warmer temperatures (>15°C) and well-watered growth medium, some or all of these immature parenchymatous cells degenerate instead of differentiating into LMTEs. Degeneration of the parenchymatous cells results in the formation of a long cavity in the central vascular region of the primary roots (Lu et al., 1991; Niki et al., 1995; Fig. 1B). To date, vascular cavities have been reported in 20 species of Fabaceae (Rost et al., 1991). Apparent restriction of this response to a closely related group of species might be of phylogenetic significance. At 25°C, 80–100% of roots in a population form cavities if plants are grown under very wet conditions, but the frequency of cavity formation becomes low to none if the growth medium is relatively dry. Sudden flooding at such warm temperatures induces rapid development of vascular cavities in the roots that otherwise do not have them. Vascular cavities never form at 10°C, irrespective of the level of water in the growth medium (Gladish and Niki, 2000; Niki and Gladish, 2001). Warm temperatures increase the rate of growth in pea roots. An increased growth rate probably increases the respiratory demand in cells of these roots (Gladish and Niki; 2000). Waterlogging is known to lower the rate of oxygen diffusion in soil (Ponnamperuma, 1972). If available oxygen at the center of the vascular cylinder becomes insufficient to meet the demand, the hypoxic stress induces vascular cavity formation (Gladish and Niki, 2000). Pea seedlings that form full-length vascular cavities continue to grow in the hypoxic environment, while those that fail to form full-length cavities, stop growing. Vascular cavities are thus thought to provide an internal, longitudinal pathway for oxygen, from the part of the root that is near the soil surface to the tissues closer to the tip (Gladish and Niki, 2000; Niki and Gladish, 2001). The formation of cortical aerenchyma under hypoxia has the same effect in other plant species. (Drew et al., 1979; Crawford, 1982), and so vascular cavities are also thought to be a type of aerenchyma (Gladish and Niki, 2000; Niki and Gladish, 2001). The cellular and molecular mechanisms involved in the cellular breakdown that leads to vascular cavity formation are not clear yet.
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observed anatomical features of the degenerating cells of vascular cavity-forming roots. These include condensation of nuclear material, degradation of cell organelles, dilution of cytoplasm as a result of tonoplast failure, and thinning of cell walls. Alteration in cell wall ultrastructure is an essential and complex step involved in many developmental processes (Rose, 2003).
Cell walls are composed of cellulose microfibrils embedded in a matrix of hemicelluloses, pectins, and proteins. Pectins play distinct roles in cell expansion and development, wall thickening, porosity, pH, and ionic status (Willats et al., 2001). A pectin network in the cell junctions adheres to adjacent cells, and breakage of these links is required for formation of intercellular spaces (Knox, 1992), separation of root border cells (Stephenson and Hawes, 1994), and for many other developmental processes, such as fruit ripening, leaf and fruit abscission, and pod dehiscence (Roberts et al., 2000). Pectins are also known to elicit defense responses against pathogens by signal transduction (Darvill et al., 1992).
Pectins are complex polysachcharides rich in 1,4-linked 
-d-galacturonic acid (GalA) residues that form approximately 20–35% of the cell wall material in dicotyledons. The three most ubiquitous pectins are homogalacturonan (HG), rhamnogalacturonan I (RGI), and rhamnogalacturonan II (RGII). HGs are linear chains of GalA residues. The carboxyl group at the C-6 position of each GalA residue can be methyl esterified. HGs are synthesized in dictyosomes (Golgi bodies) and secreted by vesicle traffic into the cell walls in a highly methyl-esterified form. Some or all of the methyl-esterified GalA residues are later de-esterified by pectin methylesterase in the cell walls (Mohnen, 1999; Willats et al., 2001, 2006; Rose, 2003). Such de-esterification of HGs facilitates inter- and intramolecular networking through Ca2+ bonds, thus strengthening the cell walls (Jarvis, 1984). Depending on the degree of esterification, the HGs are classified as high-ester HGs, low-ester HGs, or unesterified HGs. The degree and pattern of methyl esterification vary among plants, tissues and even within a single cell wall based on the functional requirements for HGs (Rose, 2003; Willats et al., 2006). The quantity and/or distribution patterns of HGs in cell walls have been observed to change during many of the developmental processes mentioned (Stephenson and Hawes, 1994; Roberts et al., 2000), in defense against pathogens (Salerno and Gianinazzi, 2004) and in response to abiotic stress such as Al-induced root inhibition and hypoxia-induced cortical aerenchyma formation (Le Van et al., 1994; Gunawardena et al., 2001b). The complex changes in cell wall ultrastructure during warm temperature and flooding-induced vascular cavity formation have never been characterized.
The primary objective of this study was to detect qualitative as well as quantitative changes in structure and composition of cell walls in response to sudden flooding at warm temperature. The study is part of a series of investigations designed to detect morphological and biochemical changes at the subcellular and molecular levels during the process of stress-induced vascular cavity formation. We examined the dimension and integrity of the primary cell walls and middle lamellae at the triangular cell junctions, using transmission electron microscopy (TEM). The quantity and distribution of two forms of HGs in the cell walls were estimated by in situ immunogold labeling using monoclonal antibodies JIM5 and JIM7. JIM5 optimally binds HGs that have more than four contiguous unesterified GalA residues adjacent to or flanked by methyl-esterified residues, while JIM7 binds to HGs with esterified residues at every second position, and the intervening residue may or may not be esterified (Clausen et al., 2003).
We compared vascular cavity-forming roots grown under warm, flooded conditions and noncavity-forming roots grown under cold, nonflooded conditions. Our results provide information toward understanding how cell wall metabolism might be regulated during vascular cavity formation. We also examined roots grown under warm, nonflooded or cold, flooded conditions to determine if cell wall ultrastructure and metabolism are affected by less stressful growth conditions that do not typically induce visible cell breakdown and cavity formation. The ultrastructural changes detected in this study, in addition to those detected by Niki et al. (1995), suggest that the process of vascular cavity formation might involve programmed cell death (PCD).
MATERIALS AND METHODS
Plant material and growth conditions
Beakers (2 L) were filled with vermiculite, moistened with 750 ml distilled water, covered with aluminum foil, and sterilized by autoclaving at 121°C for 1 h. Pea seeds (Pisum sativum L. cv. Alaska) were surface sterilized in 10% (v/v) bleach with 10 drops/L Tween-20 for 5 min, and washed in sterile double-distilled water four times, 5 min each. Seeds (20–30) were sown in the sterilized vermiculite and covered with 2–3 cm of sterilized moist vermiculite. Some beakers were placed in a growth chamber in dark at 25°C. After 5 d, when the roots were 8–10 cm in length, half of the beakers were flooded (treatment 25F) with sterile distilled water to just below the planting level to induce vascular cavity formation. Samples were then harvested at different times to follow the development of cavity formation. Samples of roots without cavities were collected from the remaining nonflooded beakers (25NF). As noncavity-forming cold controls, some plants were grown at 10°C and flooded after 15 d when the roots were 8–10 cm in length (10F), and some plants were likewise grown at 10°C but not flooded (10NF).
Scanning electron microscopy
Root tissue segments (5-mm) from different locations within 1–-3 cm of the root tip were excised. Tissues were fixed overnight in 2.4% glutaraldehyde (w/v) and 0.3% paraformaldehyde (w/v) in 0.03 M phosphate buffer (7.2 pH), then postfixed with 0.25% osmium tetroxide fixation for 1 h. After fixation, samples were dehydrated in ethanol series, flash-frozen, and freeze-fractured longitudinally in liquid nitrogen, thawed in absolute ethanol, critical-point dried, mounted on SEM stubs, coated with gold, and examined with a JEOL (Tokyo, Japan) JSM-840A scanning electron microscope.
Transmission electron microscopy
For identifying cells at different stages of cellular degeneration, 2-mm root tissue segments from different locations within 1–3 cm of the root tip were excised at different times. Tissues were immediately fixed and processed for TEM using the protocol described by Niki et al. (1995) with minor modifications. Root segments were fixed overnight in 2.4% glutaraldehyde (w/v) and 0.3% paraformaldehyde (w/v) in 0.03 M phosphate buffer (7.2 pH), postfixed with 0.025% tannic acid (w/v) for 3 h followed by 0.25% osmium tetroxide fixation for 1 h at 4°C. After fixation, the segments were stained with uranyl acetate, dehydrated with an ethanol series, embedded in Spurr’s resin (Spurr, 1969), sectioned, and mounted on copper grids.
For the study of cell wall degeneration, all samples were taken at 2 cm from the tip after 24 h of flooding. Root segments were fixed in 4% paraformaldehyde (w/v) in 0.05 M sodium cacodylate buffer (pH 7.2) at room temperature for 4 h, washed four times in the buffer for 15 min each, then fixed with 1% osmium tetroxide (w/v) in the same buffer at room temperature for 2 h, rinsed in double-distilled water four times for 15 min, and dehydrated in an ethanol series: 25% (30 min), 50% (30 min), 75% (45 min), 95% (90 min) and 100% (overnight). Tissue segments were passed through an ethanol–LR White (medium grade, Electron Microscopy Sciences, Hatfield, Pennsylvania, USA) resin series (3:1, 1:1, 1:3) for 2 h each and infiltrated in 100% LR White resin with at least two changes of resin and overnight infiltration at room temperature with constant rotation with a Pelco-R2 rotary mixer (Ted Pella, Redding, California, USA). Samples were finally embedded in LR White resin and polymerized at 56–58°C for 24 h. At least five representative roots were selected randomly for each growth condition. Ultrathin cross sections (90 nm thick, c. 0.3 mm2 area) of the central vascular region were obtained using Reichert Ultracut-S ultra-microtome(Reichert, Leica Microsystems, Bannockburn, Ilinois, USA), mounted on nickel grids (Electron Microscopy Sciences) for indirect immunogold labeling.
Indirect immunogoldlabeling
The samples on TEM grids were treated with 1% bovine serum albumin (BSA) (w/v) in phosphate-buffered saline (PBS) for 30 min to block nonspecific binding sites. Sections were then incubated with the primary antibodies JIM5 and JIM7 (Plant Probes, UK, 1:10 dilution in PBS) at room temperature for 3 h. Some sections were incubated with only PBS instead of primary antibody as a control. All sections were then washed three times in PBS and incubated with 10 nm gold labeled antirat IgG secondary antibody (Sigma, St. Louis, Missouri, USA; 1:20 dilution in PBS) for 1 h at room temperature. Finally sections were rinsed three times with PBS, then poststained with uranyl acetate and lead citrate to increase contrast. The sections were observed and photographed using a JEOL JEM-100S transmission electron microscope at 80 kV. At least three different sections from different locations of each cavity-forming roots were examined. Triangular junctions of cells from three different locations within each section were imaged: (1) c. 50 µm or more from the center of cavity (zone A), (2) c. 30–50 µm away from the center of cavity (zone B), and (3) immediately next to the cavity up to c. 30 µm from the center of cavity (zone C). Corresponding locations in noncavity-forming roots were analyzed and imaged at 40000 or 50000x for comparison.
Image analysis and statistics
Cell wall area and number of gold-labeled HG molecules present in different parts of the cell junction were determined for each image using NIH Scion Image (Scion, Frederick, Maryland, USA). The mean relative cross-sectional areas covered by the entire cell wall, primary cell wall, and middle lamella (wall area per total cell area imaged x 100) were calculated for each sample type. The mean HG density (number of labeled HGs per µm2) in each part of the cell wall at the junctions was calculated for both JIM5- and JIM7-labeled samples. The mean density of each of the HGs was compared across the different samples to determine if the differences are significant. HG densities were compared by Tukey’s studentized range test for multiple comparisons (P 0.05) using the program SAS, version 9.1.3 (SAS Institute, Cary, North Carolina, USA). The ratio of the density of each type of HG in primary cell walls to each type in the middle lamellae (P:M ratio) was determined to study the changes in the distribution patterns of HGs under different conditions.
RESULTS
Cells at different stages of cellular degeneration
Cells of noncavity-forming roots (treatments 10NF, 10F, and 25NF) appeared to be healthy and lacked signs of organellar degradation (Fig. 2A). Compared to the cell walls of 10NF roots, the walls of 25NF roots were thinner, whereas those of 10F roots were thicker (detailed later). In cavity-forming 25F roots, the vascular cavity began forming within 3 h of flooding, but only a few cells in the center of the vascular cylinder had characteristics of degeneration such as distortion of nuclei, condensation of chromatin, initial signs of tonoplast failure, and cell wall breakdown (Fig. 2B). Cavities were fully developed by 24 h after flooding. At this late stage, cells at different locations in the section had different degrees of breakdown depending on their radial distance from the center (Fig. 2C, D). Cells at the center of the section were completely degenerated, leaving a cavity. Cells immediately next to a cavity (zone C) had prominent signs of degradation, such as failure of the tonoplast, severe or complete degradation of cell organelles, and extreme thinning or breakdown of cell walls. Cells slightly away from the cavity (zone B) had partial degeneration with cytoplasm and cell organelles leaking into the vacuole due to failure of the tonoplast, but cell walls and organelles were still intact. At many cell junctions in this region, cells on one side had characteristics of degradation, whereas neighboring cells looked healthy. Cells at the periphery of the section (zone A) looked healthy with normal cell organelles, intact membranes, and thick cell walls.
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The area as well as the integrity of the cell walls in the cell junctions of cavity-forming 25F roots varied, based on the location in the section. Cell junctions in zone A had negligible signs of degeneration. The primary cell walls and middle lamellae were intact, and the middle lamellae were electron-dense and seemed to be tightly packed with wall material similar to cells of noncavity-forming roots (data not shown). Cells in zone B were at various stages of degradation, but most cell walls were intact, and the middle lamellae were electron-dense (Fig. 3B). At some cell junctions, the primary cell walls were degraded on the side toward the degenerating cells, whereas the walls toward the healthy neighboring cells appeared to be unaffected (Fig. 3C, D), which indicates that wall degradation was specifically targeted to certain cells and not others. Wall areas of cell junctions in both zone A and B of 25NF roots were similar to those of 10NF roots (Fig. 4). Cell junctions in zone C had extremely thin or no primary cell walls (Fig. 3E, F). Average relative cross-sectional area of these primary walls was low (4.9%) compared to those in all other samples (Fig. 4). Middle lamellae occupied two-thirds of the total cross-sectional wall area in cell junctions of zone C (Fig. 4), but were almost transparent (Fig. 3E) or sometimes even completely degraded (Fig. 3F).
HG density
The density of HG tended to differ among the different samples for both JIM5- and JIM7-labeled HGs even though the variations were not always statistically significant at P 0.05 (Fig. 5A, B).
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Variations among cells in cavity-forming roots
Sudden flooding at 25°C triggers rapid vascular cavity formation, but in this study we observed that not all the parenchymatous cells in the central vascular cylinder responded in a similar fashion. After 24 h of flooding, cells at the center degenerated completely to form the actual cavity. Cells adjacent to the cavity (zone C) also had severe signs of breakdown, possibly to enlarge the cavity later (Fig. 2C). As reported previously by Niki et al. (1995), these cells had distorted nuclei, condensed chromatin, degraded cell organelles, failed tonoplast, dilute cytoplasm, and highly degraded cell walls. At the triangular cell junctions, the middle lamellae were nearly electron-transparent and occupied a larger area compared to those of nondegrading cell junctions. The changes in cell junction structure could be the result of loosening and breakdown of wall components in these regions. Densities of both HGs in the middle lamellae were significantly lower than in the cells of zone A, suggesting breakdown of structural components in the middle lamella. Primary cell walls in zone C occupied less cross-sectional area than in cells of zone A and B, also indicating breakdown and loss of cell wall material. Surprisingly, the densities of both HGs in the primary cell walls did not differ significantly among the three different zones. Both HGs appear to be more concentrated in the primary cell wall than in the middle lamella. Because organelles degenerate in cells of zone C, it is unlikely that new HGs are synthesized or deposited in the primary cell wall. The higher concentration of HGs in the primary cell walls than in the middle lamellae could mean that HG molecules do not breakdown significantly in the primary cell wall area, even at very late stages of cellular breakdown. Loss of other wall material like cellulose or hemicelluloses might be responsible for the thin primary walls. Pectins (HGs and RGs together) are known to maintain structural integrity of cell walls, even when much of the cellulose fibers are removed (Shedletzky et al., 1990, 1992). Alternatively, breakdown of primary cell walls might be accompanied by HG breakdown, but at the same time hidden HG motifs might be exposed to immunolabeling. This situation could result in no apparent change in labeling.
Some cells in the next outer tiers (zone B) had initial signs of breakdown after 24 h of flooding, possibly they would undergo complete breakdown later. HG densities and localization patterns were between those of cells in zones A and C. In maize roots, oxygen concentration decreases radially in a gradient toward the center of the root, with the lowest concentration at the center of the stele (Armstrong et al., 1994). A similar gradient is likely to develop in pea roots, so that cellular breakdown is more severe toward the center where hypoxic stress is highest and less prominent in cells away from the center. At many cell junctions in this region, cells toward the central cavity had characteristics of degradation, whereas the outer neighboring cells looked healthy (Fig. 2D, 3C, D). The localization of cell degeneration could mean that the cell death process is tightly regulated spatially and does not proceed randomly.
Cells far away from the cavity had characters similar to the nondegrading cells of 10NF roots. They had normal nuclei and organelles, cell walls of normal thickness, and electron-dense middle lamellae tightly packed with wall material. The middle lamellae of these cells had a significantly higher concentration of both HGs than did the degrading cells of zone C, suggesting that a hypoxic environment does not induce cellular breakdown in the cells of zone A and that normal synthesis of wall material most likely continues in these cells. These cells might be at the higher end of the oxygen availability gradient than the cells closer to the central cavity and have enough oxygen to maintain normal function even after 24 h of flooding. Alternatively, the cell death process might be tightly regulated; therefore, cells beyond a certain radial distance are not affected, and there is sufficient xylem tissue to maintain water transport for survival and growth of the plant. JIM5 labeling was predominant in the middle lamella of these cells, which indicates the presence of a high concentration of contiguous unesterified GalA residues. High degrees of unesterified residues can mean high numbers of inter- and intramolecular cross-linking of HG chains, which also suggests that the middle lamella in these cells are strong and healthy. Because JIM7 labeling was most prominent in the primary cell walls at these cell junctions and was more abundant than JIM5 labeling, the primary cell walls have the highest concentration of highly esterified HGs, again supporting the idea of continued normal cellular metabolism because newly formed HGs are deposited in primary cell walls in a highly esterified form (Mohnen, 1999; Willats et al., 2001; Rose, 2003). High numbers of esterified HGs have been reported to significantly reduce the activity of certain cell-wall-degrading enzymes produced by plant pathogens (Vidhyasekaran, 2002; Taylor et al., 2006; Vicente et al., 2007). It might be possible that flooding at relatively warmer temperatures induce similar defense responses against wall-degrading enzymes in the cells away from the center of vascular cylinder.
Variability among noncavity-forming roots
In combination, the relatively high temperature and high water content in the growth medium are essential to the induction of vascular cavities in pea primary roots. Degeneration of cellular organelles, cell wall breakdown, and vascular cavity formation typically do not take place at 25°C without very wet or flooded conditions and never in response to flooding at 10°C (Gladish and Niki, 2000). In this study, we observed that slight changes in the cell wall ultrastructure do take place even in noncavity-forming roots grown at 25°C without flooding (25NF) or at 10°C with flooding (10F) in comparison to normal roots grown at 10°C without flooding (10NF).
Primary cell walls of roots grown at 25°C without flooding (25NF) had slightly less cross-sectional area than those of 10NF roots. Cell junctions of 25 NF roots had significantly lower densities of JIM5-labeled HGs than 10NF roots. These features suggest that high temperature by itself can cause some degradation of wall material, but it is probably not sufficient to break down the walls and lead to cavity formation. These structural differences might account for the occasional cavity observed by Gladish and Niki (2000) in roots grown at high temperature in relatively dry conditions. The density of JIM7-labeled HGs was slightly higher in walls of 25NF roots compared to 10NF roots. A high concentration of highly esterified GalA residues suggests that normal cellular functions such as the biosynthesis and deposition of HGs continue at high temperature. This increased synthesis of HGs might again be a defense response induced by warm temperature to prevent cell wall degradation.
In contrast to root responses at 25°C in nonflooded condition, roots grown at 10°C in flooded conditions (10F) had a completely opposite response. Primary cell walls and middle lamellae at the cell junctions had higher cross-sectional areas, and both had significantly higher densities of JIM5-labeled HGs in comparison to roots grown in any of the other conditions. This means an increase in HGs with contiguous unesterified residues. The density of JIM7-labeled highly esterified HGs was the same as that in the 10NF roots, perhaps meaning that flooding at cold temperature induces a significant increase in HG deposition in the cell walls, and the HGs are esterified quickly, strengthening the cell walls. Thickening of xylem vessel secondary walls has been observed in rice in response to bacterial attack (Hilaire et al., 2001), and the strengthening of cell wall junctions has been observed in response to fungal attack in several cereals (Sherwood and Vance, 1980) and has been correlated with resistance to Pseudocercosporella sp. in winter wheat (Murray and Bruehl, 1983). Thus, at cold temperatures, when demand for oxygen is low, flooding may induce a different defense response in pea roots. Thickening of the cell wall possibly increases resistance against mechanical damage due to sudden flooding.
Characters of programmed cell death (PCD)
Cortical aerenchyma formation in roots of Sagittaria lancifolia and maize has been reported to involve programmed cell death (PCD) (Schussler and Longstreth, 2000; Gunawardena et al. 2001a). The cell death mechanism involved in vascular cavity formation is not known yet. Findings of this study provide new clues toward identifying the type of cell death.
After sudden flooding, pea primary roots that form full-length cavities continue to grow, whereas those that do not form the full-length cavities stop growing (Niki and Gladish, 2001), suggesting that the plants benefit from this cell degeneration in the vascular cylinder and subsequent cavity formation. Benefiting the plant is typical of PCD responses (Gray, 2004; Potten and Wilson, 2004). PCD is also known to be limited spatially to specific cells, a relatively small group of cells, or to specific tissues (Gray, 2004; Potten and Wilson, 2004). During vascular cavity formation in pea roots, cell death remains confined to the central undifferentiated parenchymatous cells of the central vascular cylinder. Cells away from the center appear to remain healthy and functional even after cavities are formed (Niki et al., 1995; current study). We also saw this apparent segregation of cell death; at many cell junctions near the cavity, cells on one side of a junction had characteristics of degradation, while the neighboring cells looked healthy (Figs. 2D, 3C, D). The degenerating cells near the center of vascular cylinders were observed to have distorted nuclei, condensed chromatin, failed tonoplast, and thinning and breakdown of primary cell walls and middle lamellae (Niki et al., 1995; current study). Most of these characters have been associated with PCD in plant cells (Gray, 2004). Changes in HG localization that we observed in the current study were also seen during PCD-induced aerenchyma formation in maize (Gunawardena et al., 2001b). In pea seedlings grown at 25°C, cells in the central vascular cylinder of the primary roots had signs of cellular degeneration within 3 h of flooding (Fig. 2B). Cellular breakdown due to PCD also occurs within a short time after an inductive stimulus. Mittler et al. (1997) detected fragmentation of DNA into 150-kb fragments as early as 6 h in pathogen-induced PCD in tomato. DNA laddering, which indicates nucleosomal DNA fragmentation was detected within 6 h in PCD-induced curling of pea root tips (Gladish et al., 2006). Breakdown of cell organelles, nuclear breakdown, and changes in cell wall polysaccharide distribution were observed during PCD-induced cortical aerenchyma formation in maize roots in cells that were less than 1 d old (Gunawardena et al., 2001a, b).
All these data suggest that PCD might be the mechanism of cell death involved in vascular cavity formation in primary roots of pea. Further investigations are needed, however, to detect other characteristics of PCD in this system. While high temperatures without flooding or flooding at colder temperatures typically do not induce characteristics of PCD or the formation of a vascular cavity, such conditions do induce some ultrastructural variations that might also be defense responses.
FOOTNOTES
1 The authors thank Dr. R. Edelman, M. Duley, and L. Strittmatter of the Electron Microscope Facility for guidance with the TEM work, Dr. R. Noble for advice on statistical analysis, and the Department of Botany, Miami University, Oxford, OH for funding. 
5 Author for correspondence (e-mail: sarkarp{at}muohio.edu)
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