| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
Article |
Department of Genetics, Development and Cell Biology & Microscopy and NanoImaging Facility, Iowa State University, Ames, Iowa 50011-1020 USA; Department of Biochemistry, Biophysics and Molecular Biology, Iowa State University, Ames, Iowa 50011-3260 USA
Received for publication May 3, 2006. Accepted for publication November 7, 2006.
ABSTRACT
Tobacco floral nectaries undergo changes in form and function. As nectaries change from green to orange, a new pigment is expressed. Analysis demonstrated that it is ß-carotene. Plastids undergo dramatic changes. Early in nectary development, they divide and by stage 9 (S9) they are engorged with starch. About S9, nectaries shift from quiescent anabolism to active catabolism resulting in starch breakdown and production of nectar sugars. Starch is replaced by osmiophilic bodies, which contain needle-like carotenoid crystals. Between S9 and S12, amyloplasts are converted to chromoplasts. Changes in carotenoids and ascorbate were assayed and are expressed at low levels early in development; however, following S9 metabolic shift, syntheses of ß-carotene and ascorbate greatly increase in advance of expression of nectar redox cycle. Transcript analysis for carotenoid and ascorbate biosynthetic pathways showed that these genes are significantly expressed at S6, prior to the S9 metabolic shift. Thus, formation of antioxidants ß-carotene and ascorbate after the metabolic shift is independent of transcriptional regulation. We propose that biosynthesis of these antioxidants is governed by availability of substrate molecules that arise from starch breakdown. These processes and events may be amenable to molecular manipulation to provide a better system for insect attraction, cross pollination, and hybridization.
Key Words: amyloplast • ascorbate • ß-carotene • chromoplast • development • floral nectary • Solanaceae • starch • tobacco
The changes in plastid ultrastructure and function during development have been studied in a variety of organs and plants. These changes typically follow various pathways, which may lead to formation of amyloplasts (Bechtel and Wilson, 2003
), and in the cases of petal and fruit development, into chromoplasts (Pyke, 1999
; Camara et al., 1995
; Pyke and Page, 1998
; Weston and Pyke, 1999
; Pyke and Howells, 2002
). These changes may also occur in certain roots and underground stems without passing through the chloroplast stage. The switchover from plastids containing chlorophylls to those containing carotenoids (Cheung et al., 1993
; Fraser et al., 1994
; Ronen et al., 2000
; Busch et al., 2002
; Castillo et al., 2005
) is related to ripening and senescence. Even though this process has been documented many times, some systems may have unique aspects that warrant further study. This is the case for tobacco floral nectaries where large, engorged amyloplasts are initially converted into intermediate plastids—amylochromoplasts (plastids containing both starch and ß-carotene crystals)—and finally to chromoplasts. In addition, the catabolic and anabolic processes associated with these conversions define a complex and well-coordinated series of biochemical reactions that involve nectar production (Pacini et al., 2003
; Carter et al., 2006
) and nectar, nectary, and gynoecium protection.
Plastids are dynamic and versatile double-membrane-bounded organelles that are classified into at least six distinct categories—chloroplasts, leucoplasts, elaioplasts, amyloplasts, chromoplasts, and undifferentiated plastids or proplastids (Waters and Pyke, 2004
). They have a range of sizes, shapes, and numbers per type of cell. They are related to the age of a cell and its developmental state (Osteryoung and McAndrew, 2001
). Plastids seem to be well suited to adapting to the complexity and function of the changing cell in which they exist. Even though both extrafloral and floral nectaries have been variously studied (O'Brien et al., 1996
; Horner et al., 2003
; Pacini et al., 2003
; Stpiczynska et al., 2004
, 2005
), only cursory attention has been directed to the changes in the plastids. Therefore, the developing tobacco floral nectary provides an interesting system in which to study these changes as part of a gland whose dynamics are associated with starch synthesis and catabolism, nectar and hydrogen peroxide production, and antioxidant (ß-carotene and ascorbate) synthesis—a series of integrated processes reported together in this study for the first time. The purpose of this study is to follow the developmental changes in the plastids up to maturity and beyond, to relate both associated biochemical and molecular processes with these changes, and to understand the underlying mechanisms that control them. This combined information will serve as the basis for developing strategies to manipulate the components comprising the nectar, which in turn will have the potential for increasing insect visitation and ultimately cross-pollination for hybridization and crop improvement.
MATERIALS AND METHODS
Ornamental tobacco plants that overproduce nectar [from interspecific cross of Nicotiana langsdorffii x N. sanderae Hort. Var Sutton's Scarlet line LxS8 (Kornaga et al., 1997
)] were grown in a temperature-controlled glasshouse on the Iowa State University campus under a photoperiod regimen that allowed for continuous flowering. Most plants provided an array of flowers at different stages of development that can be identified by stage (stages 2, 4, 6, 9, 10, 11, and 12; Figs. 1--7) per Koltunow et al. (1990)
. The flowers were processed for both light and electron microscopy, and biochemically analyzed for starch, carotenoids, and ascorbic acid according to the following methods.
|
).
Vibratome sections
Fresh floral gynoecia at S9 through S12 were glued in two orientations (trans- and longi-planes through gynoecia and nectaries) to the base reservoir with a fast-drying epoxy glue. Deionized water was added just above the top of each specimen, and 30-µm-thick sections were cut and placed in drops of deionized water on slides. Cover slips were added and sections were viewed either in bright-field (white light [WL]) mode or between crossed polarizers (CP), and digitally captured using a Zeiss MRc Axiocam camera (http://www.zeiss.de) mounted on an Olympus BH10 microscope (http://www.olympus-global.com).
Electron microscopy
For general ultrastructure, small portions of fresh, isolated nectaries were initially fixed in 2% glutaraldehyde and 2% paraformaldehyde in a 0.1 M sodium cacodylate buffer, pH 7.2, at 4°C overnight, washed in cold buffer, postfixed in 1% osmium tetroxide in same buffer at 4°C for 1.5 h, washed with buffer and deionized water, and en bloc stained with 2% aqueous uranyl acetate in the dark. The specimens were washed in deionized water. Then the specimens were divided into two batches for further processing into either Quetol or Spurr's resin mixtures. For Quetol resin embedding (manufacturer's instructions; http://www.polysciences.com) after deionized water washes, specimens were processed through increasing concentrations of Quetol and decreasing ratios of deionized water to pure Quetol monomer, and then into Quetol resin mixture (Quetol 651, 15 g; methyl nadic anhydride [MNA], 10 g; nonyl succinic anhydride [NSA], 20 g; and tridimethylaminomethyl phenol [DMP-30], 1 g), followed by casting in aluminum trays. Polymerization was done at 75°C for 1 d. For Spurr resin embedding (Spurr, 1969
) following deionized water washes, specimens were dehydrated in an ethanol series, transferred to pure acetone, and infiltrated in acetone and Spurr's hard resin mixture. Specimens were then transferred into pure resin mixture, cast in aluminum trays, and polymerized at 70°C. Specimens embedded in both resin mixtures were sectioned using either glass or diamond knives. Thin 60-nm-thick sections were mounted on Formvar-coated slotted grids and stained with 2% aqueous uranyl acetate and lead citrate (Reynolds, 1963
). Observations were made on a JEOL 1200EX TEM (www.jeol.com, and images were captured photographically on Kodak SO-163 film (www.kodak.com), which was later digitized for further processing using an Epson Perfection 3200 PhotoScanner (http://www.epson.com).
Biochemical analyses
Carotenoids analysis
Nectaries were harvested from flowers at different stages (Koltunow et al., 1990
) as previously described (Carter and Thornburg, 2000
). Approximately 10 nectaries were combined in a glass tissue homogenizer with a total volume of 300 µl acetone. After homogenization, the organic layer was removed, and the process was repeated, first with 300 µl acetone, then with 300 µl hexane. The three organic layers were combined, dehydrated with sodium sulfate, and evaporated to dryness, taken up in 50 µl hexane and used for analysis. Thin layer chromatography (TLC) was performed on silica gel plates using a 9 : 1 hexane : acetone solvent. HPLC was performed exactly as described (Barua and Olson, 1998
) on a Waters HPLC system (http://www.waters.com) using a 3-mm Microsorb-MV column (http://www.rainin-global.com). Carotenoids were estimated at 450 nm.
Ascorbic acid analysis
Nectaries were harvested from staged flowers as previously described. Dissected nectaries from approximately 30 flowers were homogenized in an equal volume of 1% oxalic acid in a Con-Torque tissue homogenizer (http://www.eberbachlabtools.com). Following homogenization, the samples were centrifuged (13 000 x g, 5 min) to remove debris. The pellet was re-extracted with an additional volume of 1% oxalic acid and reprocessed. The supernatants were combined and a 50 µl sample in a total volume of 2 ml of 1% oxalic acid was titrated to a pink endpoint with 0.05% 2,6-dichlorophenolindophenol (DCIP) in 0.1 M phosphate buffer, pH 7.0. A standard curve of the sample containing up to 20 µg of ascorbate was prepared to quantitate levels of ascorbate.
Labeling of carotenoids with [14C]-sucrose
Flower buds at S9 were harvested and placed with pedicels in water at room temperature. Subsequently 0.8 µCi [14C]-sucrose (per flower) was added to the water. Incubation continued for 48 h with additional water added when necessary. Nectaries (three per sample, duplicate samples) were extracted 2x with acetone and once with hexane. The combined organic phases were dried in a rotary evaporator, resuspended in 2 : 1 : 1 acetone : hexane : acetonitrile, and a 10 µL sample was counted by scintillation. The thin layer chromatograms with 10 µL per sample (Kodak 13181 Silica Gel; http://www.kodak.com) were developed in 9 : 1 hexane : acetone. Autoradiograms were prepared by exposing each chromatogram to a phosphorimager plate overnight.
Oligonucleotides for RT-PCR analysis
For synthesis of the oligonucleotides for RT-PCR analysis, the ornamental tobacco cDNA clones were mapped onto the Arabidopsis thaliana genomic clones encoding the same genes. This permitted the localization of the Arabidopsis introns in the tobacco cDNA sequences. Oligonucleotides were then synthesized to cDNA regions that spanned the hypothetical intron junctions. All oligonucleotides were first tested with genomic DNA to insure that the oligonucleotides did not amplify a fragment with genomic DNA. The oligonucleotides used in this analysis are listed in Table 1. Control oligonucleotides were also prepared for the 18S and 26S rRNA genes.
|
In ornamental tobacco, the nectary is embedded in the base of the gynoecium. It encircles the base and has a continuous cuticle over its entire surface (Figs. 1--7). It has two stomatal zones 180° in line with the internal septum (Fig. 8; arrowheads; Thornburg et al., 2003
). All the vascular tissues (veins) emanating from the pedicel below the nectary pass through the gynoecial walls next to the locules and interior to the nectary but do not penetrate the nectary special parenchyma anywhere (Fig. 8; asterisks). Beginning at S2, the special parenchyma accumulates starch. We have begun a series of studies to evaluate development of the floral nectary. We have used macromicroscopy of gynoecia and nectaries throughout nectary development to identify developmental changes in this floral organ.
Between S2 and S9 both the gynoecium and nectary increase in size (G. Ren et al., unpublished data). In addition, the nectary color changes from lime green (S2 and S4; Figs. 1, 2) to yellow (S6; Fig. 3) to a light orange (S9; Fig. 4). Between stages S10 and S12, the nectary matures and becomes a deep, bright orange (Figs. 5--7). Nectar secretion is initiated at late S10 or early S11 and continues beyond S12.
Light microscopy: S9 through S12
Previous studies indicate that approximately 20% of the nectary mass is composed of starch (G. Ren et al., unpublished data) by S9, when it is engorged with starch (Figs. 9, 10, 15, 19). Beginning at S2, the special parenchyma accumulates starch. Starch continues to accumulate through S9 when the nectary changes greatly and the accumulated starch is degraded to produce sugars for nectar production. To better evaluate these changes during this maturation phase of nectary development, we prepared vibratome longitudinal and cross sections through the gynoecium and nectary. These showed that the nectary color changes from the inner part of the nectary toward the single-layered epidermis during S10 and S12 (Figs. 11--14). At S9, the end of the nectary starch filling period, each special parenchyma cell contains plastids that in turn contain multiple starch granules.
|
Electron microscopy: S9 through S12
S9 plastids
The enlarged S9 plastids are uniformly filled with multiple starch grains and have a minimum of lamellae and stroma (Figs. 21, 25). There are very few, small osmiophilic bodies visible in the stroma. There is an average of 8.5 plastids per special parenchyma cell area with each plastid volume averaging about 1000 µm3. Each plastid volume consists of about 78% starch, so that at S9 the ratio of cell volume to starch volume is 5.9 : 1. This figure corresponds well with the previously estimated quantification of starch in nectaries of 20% of total nectary mass, which represents starch (G. Ren et al., unpublished data).
|
The S12 plastids
By S12 (anthesis), many but not all of the plastids in the special parenchyma have lost their starch (Fig. 24), while plastids in other cells have a few starch grains as well as an increased number of crystals (Figs. 28--31). These latter plastids contain many osmiopilic bodies, and there are many crystals that have enlarged and extended the boundary membranes of the plastids (Fig. 31). In addition, stroma lamellae in some plastids have proliferated into what appear to be sheets or reticulate networks in some plastids (not shown). The crystal-containing plastids are consistent with the plastids seen in the Vibratome sections (Figs. 17, 18). If the flowers are not pollinated, these plastids with their crystals continue for at least 48 h before they degenerate. The special parenchyma cells at S12 are highly vacuolated (Fig. 24), but retain their cellular identity with intact nuclei, mitochondria, ER, and vacuoles.
Biochemical and molecular genetic analyses
To accompany these anatomical and developmental results, a series of biochemical and molecular genetic analyses designed to identify and clarify the roles of these compounds in nectary function was conducted. Initially, the yellow pigment that accumulates in the earlier stages and appears at its maximum in the mature S12 nectaries was isolated and characterized.
To identify the yellow pigment, extracts of the nectary for analysis were prepared. As shown in Fig. 32B, the nectary extract gives a single peak by HPLC analysis. On the same HPLC column, three common carotenoids (lutein, lycopene, and ß-carotene) were run (Fig. 32A). The nectary extracted material co-chromatographed with ß-carotene. Further analysis indicated that this yellow pigment had an identical visible absorption spectrum to ß-carotene (Fig. 32C).
|
|
|
|
). Thus, during floral development the genes encoding these ß-carotene biosynthetic enzymes were regulated (both up and down) in the nectary. A surprising result was that ß-carotene biosynthetic gene expression reached maximal levels by S6, yet the levels of ß-carotene remained low until S9. Because of this observation, the regulation of ß-carotene production must be at a level other than the transcriptional level. While posttranscriptional regulation of those genes (either at the translational or posttranslational level [protein regulatory]) could account for this observation, we believe that a simpler and more plausible explanation is simply the availability of substrates for ß-carotene production. In addition, we evaluated the expression of the carotene dioxygenase, ccd1, which uses carotene as a substrate for the formation of apocarotenoids. This gene was also strongly expressed in S6 and S9 nectaries and its reduced expression continued through S12, but it was turned off after fertilization.
|
|
To evaluate whether sugars can serve as precursors for ß-carotene in the nectary, [14C]-sucrose was fed through the pedicel of immature (S9) flowers and after incubation for 2 d, the nectaries were dissected from the flowers, and the carotenoids were isolated and evaluated by TLC. As can be seen in Fig. 36, Lanes 1 and 2, ß-carotene is the predominant pigment isolated from the nectaries of these flowers. When the thin layer plate was exposed to reveal the location of the isotope, the ß-carotene was highly labeled confirming the biochemistry shown in Fig. 35, indicating that sugars can feed through the MEP pathway into ß-carotene biosynthesis.
|
|
|
To evaluate the patterns of expression for each of these genes, we performed RT-PCR using mRNA isolated from various tissues and at different developmental stages of the nectary. This analysis is shown in Fig. 38B. As was observed for the ß-carotene biosynthetic pathway, the ascorbate biosynthesis genes were also widely expressed throughout the plant. Various levels were found in all of the organs examined. In the nectary, each of these three genes was expressed maximally by S6, and then expression gradually declined. Thus, it seems that as observed for ß-carotene biosynthesis, ascorbate biosynthesis likely takes place in the nectary at early developmental times; however, major levels of ascorbate did not accumulate until after S9, when starch breakdown began. It appears that the biochemical pathways for both ß-carotene and ascorbate are present early in nectary development, yet accumulation of these antioxidants did not occur at high levels until starch breakdown began at about S10. These data indicate that the production of these antioxidants appears to be regulated in late nectary development by substrate availability.
DISCUSSION
The tobacco floral nectary is a highly active gland whose early and intermediate developmental stages (S2 through S9) are characterized by the synthesis of starch in plastids and, to a lesser degree, carotenoids in the plastid thylakoids. These events are marked externally by the increase in sizes of the gynoecium and the basal nectary, and the change in color of the nectary from a lime green (S2) to bright orange (S9). Based on an ongoing study (G. Ren et al., unpublished data), we believe the plastids in the young nectaries are not chloroplasts, based on the lack of distinct grana and their capacity to divide, but are plastids containing photosynthetic pigments with the signals to focus on starch synthesis. At S9, the special parenchyma has become engorged with starch-filled amyloplasts containing a few dense thylakoids. Between S9 and S10, the flower opens, and the secretion of nectar begins and lasts through at least S12.
The events that follow S9 are remarkable. The starch in the amyloplasts is rapidly catabolized to form nectar sugars (G. Ren et al., unpublished data) and a substrate source for ß-carotene and ascorbate biosynthesis. The biosynthesis of the carotenoids is so great that the ß-carotene forms multiple crystals in the majority of plastids. We identify these dual-function plastids that contain both starch and ß-carotene during the later developmental stages as amylochromoplasts. In addition, other metabolic events occur late in nectary development. They are the expression of a novel MYB transcription factor that regulates NEC1 expression prior to anthesis (G-Y. Liu et al., unpublished data), and the resulting NEC1-mediated accumulation of H2O2 in the secreted nectar (Thornburg et al., 2003
).
Durkee (1983)
has characterized several types of floral nectaries developmentally based on whether their internal cells degenerate during secretion, such as holonecrine in soybean (Horner et al., 2003
) or as eccrine when the internal secretory cells retain their integrity for some time following the major secretory phase, as is the case for tobacco. The tobacco nectaries have been shown to continue to secrete beyond S12 if they have not been pollinated.
Even chromoplast development in a variety of fruits and other plant structures has been followed in other studies (discussed in the introduction), the present study clearly documents that the plastids from the early stages of nectary development through S12 may be considered to have a dual function. We thus give them the name amylochromoplast. Biochemical analysis of carotenoids during the entire period of nectary development (S2 through S12) showed that in the earlier stages carotenoids were formed most likely in the plastid lamellae, and at S9 this process greatly increased with the spectacular formation of ß-carotene crystals from the osmiophilic bodies. We believe these osmiophilic bodies are the later sites of the carotenoids and serve as the centers for ß-carotene crystal development. Their production occurs in all of the nectary cells to a greater or lesser extent, giving the nectaries their distinct orange appearance from S10 through S12. Different methods of preservation and embedments were used to better maintain them. However, they were only undistorted in the fresh Vibratome sections. In that our primary goal was not to better characterize these crystals microscopically, but only to show their presence, we did not pursue cryopreservation and cryosectioning methods.
Thornburg et al. (2003)
concluded that the production of the NEC1 protein was involved in H2O2 production in the nectar. This oxidative compound provides the nectar with an oxidizing medium to serve as an antimicrobial environment to protect the nectar sugars and other diverse components present (Davis et al., 1998
; Pacini et al., 2003
; C. Carter et al., in press
). The expression of this protein and its transcription factor beginning at S9, using in situ hybridization and immunocytochemistry, supports this contention (C. Carter et al., in press
). Because the S9 nectary cells that produce NEC1 remain viable through S12 and beyond, it seems that the cells of the nectary, and possibly the rest of the gynoecium, need to be protected. Our results strongly suggest that the production of ß-carotene and ascorbic acid provide the counterbalancing antioxidants needed to accomplish this protection.
Furthermore, and in concert with a number of other recent studies (Smirnoff and Wheeler, 2000
; Green and Fry, 2005
), some of the ascorbic acid may be converted into calcium oxalate crystals (Horner et al., 2000
), which come to line the gynoecial wall and are interspersed in the septum between the two locules during the later stages of gyneocium development. These crystal-filled idioblasts, containing prismatic and crystal sand crystals are evident in Figs. 10, 12, and 14. These crystals may serve a secondary function as a physical and chemical barrier to protect the gynoecium and the enclosed developing seeds from insects and other organisms that can use them as a source of food (Korth et al., 2006
).
Nectar begins to be secreted by late S10 and early S11. One of the primary characteristics of nectar secretion in ornamental tobacco, in addition to the sugar secretion is the production of H2O2 via the Carter–Thornburg nectar redox cycle. This recently discovered biochemical pathway (Carter et al., 1999
; Carter and Thornburg, 2000
, 2004a
--c
) produces high levels of H2O2 (up to 4 mM) that are antimicrobial and protect the gynoecium and the developing ovules from microbes transferred to the flower by visiting pollinators or by wind/weather. Thus one notable feature of the nectar of ornamental tobacco flowers is the highly oxidative environment found in nectar. As a consequence, the nectary likely produces antioxidants to help alleviate this oxidative stress. We have previously identified ascorbate as the major extracellular antioxidant in soluble nectar (Carter and Thornburg, 2004a
, b
), and we propose that ß-carotene serves as the major intracellular antioxidant in the nectary. Its production peaks at anthesis (S12) exactly when the oxidative stress is greatest.
To evaluate gene expression affecting carotenoid and ascorbate biosynthesis, we isolated 12 cDNAs in these two biosynthetic pathways. Based on the nucleotide sequence of each of these clones, we developed RT-PCR oligonucleotides pairs (Table 1) to permit analysis of this biosynthetic pathway in nectaries. As shown in Fig. 36, the entire pathway of ß-carotene biosynthesis is well expressed in nectaries, and maximal levels of these cDNAs are expressed by developmental S6 to S9. Thus, the levels of mRNAs encoding the ß-carotene biosynthetic enzymes are highly expressed before the maturation phase (S9 to S12), when most of the ß-carotene is produced. Clearly another factor, other than transcriptional regulation, is responsible for regulating the production of carotenoids in the nectary. There are two possible regulatory paradigms that could function in this regard. First, posttranscriptional regulation of those genes, either at the translational level or posttranslational (protein regulatory) level, could account for this observation. However, these phenomena generally do not function to regulate an entire pathway. Further, because low levels of ß-carotene are being produced at these early stages (see Figs. 1--4), we postulate that wholesale down-regulation of the entire pathway is unlikely. A simpler and more elegant explanation is the availability of substrates for ß-carotene production.
In summary, the tobacco nectary represents a very complex factory that simultaneously involves a wide variety of anabolic and catabolic processes that together contribute to nectar production and protection. This study has provided fundamental information for future studies that may help to alter nectar production and composition for better insect visitation, cross pollination, and hybridization of important crop and horticultural plants.
FOOTNOTES
1 The authors thank the USDA SCA #58-3625-3-104 for partial support of this project to H.T.H., the National Science Foundation Grant #IBN-0235645 to R.W.T., and the Microscopy and NanoImaging Facility and its staff for use of its facilities under the directorship of H.T.H. 
2 Author for correspondence (e-mail: hth{at}iastate.edu
), phone: 515-294-8635, fax: 515-294-1337
LITERATURE CITED
Barua A. B. Olson J. A.. 1998. Reversed-phase gradient high-performance liquid chromatographic procedure for simultaneous analysis of very polar to nonpolar retinoids, carotenoids and tocopherols in animal and plant samples. Journal of Chromatography, B, Biomedical Sciences and Applications 707: 69-79.[CrossRef]
Bechtel D. B. Wilson J. D.. 2003. Amyloplast formation and starch granule development in hard red winter wheat. Cereal Chemistry 80: 175-183.[CrossRef][ISI]
Busch M. Seuter A. Hain R.. 2002. Functional analysis of the early steps of carotenoid biosynthesis in tobacco. Plant Physiology 128: 439-453.
Camara B. Hugueney P. Bouvier F. Kuntz M. Monéger R.. 1995. Biochemistry and molecular biology of chromoplast development. International Review of Cytology 163: 175-247.[ISI][Medline]
Carter C. Graham R. Thornburg R. W.. 1999. Nectarin I is a novel, soluble germin-like protein expressed in the nectar of Nicotiana sp. Plant Molecular Biology 41: 207-216.[CrossRef][ISI][Medline]
Carter C. Healy R. O'Toole N. Naqvi S. S. M. Ren G. Park S. Beattie G. Horner H. T. Thornburg R. W. In press Tobacco nectaries constitutively express a novel NADPH oxidase that is implicated in the defense of floral reproductive tissues. Plant Physiology..
Carter C. Shafir S. Yehonatan L. Palmer R. G. Thornburg R. W.. 2006. A novel role for proline in plant floral nectars. Naturwissenschaften 93: 72-79.[CrossRef][ISI][Medline]
Carter C. Thornburg R. W.. 2000. Tobacco nectarin I: purification and characterization as a germin-like, manganese superoxide dismutase implicated in the defense of floral reproductive tissues. Journal of Biological Chemistry 275: 36726-36733.
Carter C. Thornburg R. W.. 2004a. Is the nectar redox cycle a floral defense against microbial attack?. Trends in Plant Science 9: 320-324.[CrossRef][ISI][Medline]
Carter C. Thornburg R. W.. 2004b. Tobacco nectarin III is a bifunctional enzyme with monodehydroascorbate reductase and carbonic anhydrase activities. Plant Molecular Biology 54: 415-425.[CrossRef][ISI][Medline]
Carter C. Thornburg R. W.. 2004c. Tobacco nectarin V is a flavin-containing berberine bridge enzyme-like protein with glucose oxidase activity. Plant Physiology 134: 460-469.
Castillo R. Fernandez J.-A. Gomez-Gomez L.. 2005. Implications of carotenoid biosynthesis genes on apocarotenoid formation during the stigma development of Crocus sativus and its close relatives. Plant Physiology 139: 674-689.
Cheung A. Y. McNellis T. Piekos B.. 1993. Maintenance of chloroplast components during chromoplast differentiation in the tomato mutant green flesh. Plant Physiology 101: 1223-1229.[Abstract]
Cookson P. J. Kiano J. W. Shipton C. A. Fraser P. D. Romer S. Schuch W. Bramley P. M. Pyke K. A.. 2003. Increases in cell elongation, plastid compartment size and phytoene synthase activity underlie the phenotype of the high pigment-1 mutant of tomato. Planta 217: 896-903.[CrossRef][ISI][Medline]
Davis A. R. Pylatuik J. D. Paradis J. C. Low N. H.. 1998. Nectar-carbohydrate production and composition vary in relation to nectary anatomy and location within individual flowers of several species of Brassicaceae. Planta 205: 305-318.[CrossRef][ISI][Medline]
Durkee L. T.. 1983. The ultrastructure of floral and extrafloral nectaries of Passiflora. In B. Bentley, T. Elias [eds.] The biology of nectaries, 1-29 Columbia University Press, New York, New York, USA..
Fraser P. D. Truesdale M. R. Bird C. R. Schuch W. Bramley P. M.. 1994. Carotenoid biosynthesis during tomato fruit development. Plant Physiology 105: 405-413.[Abstract]
Green M. A. Fry S. C.. 2005. Vitamin C degradation in plant cells via enzymatic hydrolysis of 4-O-oxalyl-l-threonate. Nature 433: 83-87.[CrossRef][Medline]
Horner H. T. Healy R. A. Cervantes-Martinez T. Palmer R. G.. 2003. Floral nectary fine structure and development in Glycine max L. (Fabaceae). International Journal of Plant Sciences 164: 675-690.[CrossRef]
Horner H. T. Kausch A. P. Wagner B. L.. 2000. Ascorbic acid: a precursor of oxalate in crystal idioblasts of Yucca torreyi in liquid root culture. International Journal of Plant Sciences 161: 861-868.[CrossRef]
Koltunow A. M. Truettner J. Cox K. H. Walroth M. Goldberg R. B.. 1990. Different temporal and spatial gene expression patterns occur during anther development. Plant Cell 2: 1201-1224.
Kornaga T. Zyzak D. Kintinar A. Baynes J. W. Thornburg R. W.. 1997. Genetic and biochemical characterization of a "lost" unstable flower color phenotype in interspecific crosses of Nicotiana. The WWW Journal of Biology 2: 8..
Korth K. L. Doege S. J. Park S. H. Goggin F. L. Wang Q. Gomez S. K. Liu G. Jia L. Nakata P. A.. 2006. Medicago truncatula mutants demonstrate the role of plant calcium oxalate crystals as an effective defense against chewing insects. Plant Physiology 141: 188-195.
O'Brien S. P. Loveys B. R. Grant W. J. R.. 1996. Ultrastructure and function of floral nectaries of Chamelaucium uncinatum (Myrtaceae). Annals of Botany 78: 189-196.
Osteryoung K. W. McAndrew R. S.. 2001. The plastid division machine. Annual Review of Plant Physiology and Plant Molecular Biology 52: 315-333.[CrossRef][ISI][Medline]
Pacini E. Nepi M. Vesprini J. L.. 2003. Nectar biodiversity: a short review. Plant Systematics and Evolution 238: 7-21.[ISI]
Pyke K. A.. 1999. Plastid division and development. Plant Cell 11: 549-556.
Pyke K. A. Howells C. A.. 2002. Plastid and stromule morphogenesis in tomato. Annals of Botany 90: 559-566.
Pyke K. A. Page A. M.. 1998. Plastid ontogeny during petal development in Arabidopsis. Plant Physiology 116: 797-803.
Reynolds E. S.. 1963. The use of lead citrate at high pH as an electron opaque stain in electron microscopy. Journal of Cell Biology 17: 208-212.
Ronen G. Carmel-Goren L. Zamir D. Hirschberg J.. 2000. An alternative pathway to beta-carotene formation in plant chromoplasts discovered by map-based cloning of beta and old-gold colour mutations in tomato. Proceedings of the National Academy of Sciences, USA 97: 11102-11107.
Ruzin S. E.. 1999. Plant microtechnique and microscopy Oxford University Press, New York, New York, USA..
Smirnoff N. Wheeler G. L.. 2000. Ascorbic acid in plants: biosynthesis and function. Critical Reviews in Biochemical Molecular Biology 35: 291-314.[CrossRef]
Spurr A. R.. 1969. A low-viscosity epoxy resin embedding medium for electron microscopy. Journal of Ultrastructure Research 26: 31-34.[CrossRef][ISI][Medline]
Stpiczynska M. Davies K. L. Gregg A.. 2004. Nectary structure and nectar secretion in Maxillaria coccinea (Jacq.) L.O. Williams ex Hodge (Orchidaceae). Annals of Botany 93: 87-95.
Stpiczynska M. Davies K. L. Gregg A.. 2005. Comparative account of nectary structure in Hexisea imbricata (Lindl.) Rchb.f. (Orchidaceae). Annals of Botany 95: 749-756.
Thornburg R. W. Carter C. Powell A. Mittler R. Rizhsky L. Horner H. T.. 2003. A major function of the tobacco floral nectary is defense against microbial attack. Plant Systematics and Evolution 238: 211-218.[ISI]
Waters M. T. Pyke K. A.. 2004. Plastid development and differentiation. In S. G. Møller [ed.] Plastids, 30-59 Blackwell, Oxford, UK..
Weston E. L. Pyke A. K.. 1999. Developmental ultrastructure of cells and plastids in the petals of wallflower (Erisymum chieri). Annals of Botany 84: 763-769.
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
-carotene desaturase; lyc, lycopene ß-cyclase; ccd1, carotenoid cleavage dioxygenase; nxs, neoxanthin synthase; 26S rRNA; 18S rRNA. GenBank accession numbers for genes are in 
