Desiccoplast–etioplast–chloroplast transformation under rehydration of desiccated poikilochlorophyllous Xerophyta humilis leaves in the dark and upon subsequent illumination

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Physiology

Desiccoplast–etioplast–chloroplast transformation under rehydration of desiccated poikilochlorophyllous Xerophyta humilis leaves in the dark and upon subsequent illumination Katalin Solymosi a , Zoltán Tuba b,c,1 , Béla Böddi a,∗ a

Department of Plant Anatomy, Eötvös University, Pázmány P. s. 1/c., Budapest H-1117, Hungary “Plant Ecology” Research Group of the Hungarian Academy of Sciences, Szent István University, Páter K. u. 1., Gödöllo˝ H-2103, Hungary c Institute of Botany and Ecophysiology, Faculty of Agricultural and Environmental Sciences, Szent István University, Páter K. u. 1., Gödöllo˝ H-2103, Hungary b

a r t i c l e

i n f o

Article history: Received 5 September 2012 Received in revised form 29 November 2012 Accepted 29 November 2012 Keywords: Chlorophyll biosynthesis Etioplast Poikilochlorophyllous desiccation-tolerant plant Protochlorophyllide Rehydration

s u m m a r y The transformation of desiccoplasts into etioplasts and the parallel appearance of protochlorophyllide (Pchlide) forms were observed with transmission electron microscopy and 77 K fluorescence spectroscopy, when air-dried detached leaves of the poikilochlorophyllous desiccation tolerant plant Xerophyta humilis were floated in water in the dark. After 1 week of rehydration, pregranal plastids with newly synthesized prothylakoid (PT) lamellae and mainly non-photoactive Pchlide forms developed, while etioplasts with prolamellar bodies (PLBs) and photoactive, 655 nm emitting Pchlide form accumulated primarily in the basal leaf regions after 2 weeks of regeneration. When these latter leaves were illuminated with continuous light for 3 days, the etioplasts transformed into regular chloroplasts and the fluorescence emission bands characteristic of green leaves appeared. These results show that, upon rehydration, the dehydrated chlorenchyma cells are able to regenerate pregranal plastids and etioplasts from desiccoplasts in the dark, which can transform into regular chloroplasts when they are illuminated. This means that the differentiation of pregranal plastids and etioplasts and their greening process is a basic property of fully differentiated cells of X. humilis. Consequently, these processes are not merely characteristic for seedlings with meristematic and differentiating young tissues. © 2013 Published by Elsevier GmbH.

Introduction Desiccation tolerance of seeds, spores and pollen is common and has been studied in detail. Much less is known about this phenomenon in vegetative tissues. Plants tolerating desiccation of their vegetative tissues are called desiccation-tolerant (DT) or resurrection plants (Gaff, 1971; reviewed by Proctor and Tuba, 2002; Rascio and La Rocca, 2005; Moore et al., 2009). Two main strategies have evolved among DT plants (Tuba et al., 1994; Oliver et al., 2000). The so-called homoiochlorophyllous plants (HDTs) normally retain their photosynthetic pigments and membranes upon dehydration. The majority of non-vascular and vascular DT plants belong to this category. The poikilochlorophyllous resurrection plants (PDTs) represent the other category

Abbreviations: Chl, chlorophyll; Chlide, chlorophyllide; DT, desiccation tolerant; Fluor. int. (rel.), relative fluorescence intensity; FM, fresh mass; HDT, homoiochlorophyllous DT; Pchlide, protochlorophyllide; PDT, poikilochlorophyllous DT; PLB, prolamellar body; POR, NADPH:Pchlide oxidoreductase; PT, prothylakoid. ∗ Corresponding author. Tel.: +36 13812165; fax: +36 13812166. E-mail addresses: [email protected], [email protected] (B. Böddi). 1 Deceased author.

(Vassiljev, 1931; Hambler, 1961); they decompose their chlorophylls (Chl-s), their thylakoid membranes disintegrate during dehydration (Hallam and Luff, 1980) and their chloroplasts become desiccoplasts (Tuba et al., 1994, 1996), also termed xeroplasts (Ingle et al., 2008). Different mechanisms have been suggested for the survival of the DT plants. Xerophyta humilis and other Xerophyta species seem to retain their cells and compartmentalization by accumulating proteins, sugars and various osmolytes upon dehydration (Farrant et al., 1999; Farrant, 2000). The chloroplast–desiccoplast transformation has been shown to be reversible, i.e. Chl-s and thylakoids are re-synthesized via reconstitution processes upon rewetting (Tuba et al., 1993a,b, 1994). The ultrastructural changes of the PDT’s chloroplasts upon dehydration and rehydration have been studied in detail for Xerophyta species (Hallam and Luff, 1980; Tuba et al., 1993a,b, 1994; Sherwin and Farrant, 1998; Farrant et al., 1999; Farrant, 2000; Ingle et al., 2008). Changes in the Chl and carotenoid content (Tuba et al., 1994, 1996; Sherwin and Farrant, 1998; Farrant et al., 1999; Farrant, 2000; Ingle et al., 2008), increasing activity of antioxidant enzymes during rehydration (Sherwin and Farrant, 1998; Farrant, 2000), transcriptional and translational activities (Dace et al., 1998; Ingle et al., 2008), as well as CO2 fixation, respiration and photosynthetic parameters (Tuba et al., 1994, 1996, 1997; Ingle et al., 2008) have been analyzed in this genus.

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Please cite this article in press as: Solymosi K, et al. Desiccoplast–etioplast–chloroplast transformation under rehydration of desiccated poikilochlorophyllous Xerophyta humilis leaves in the dark and upon subsequent illumination. J Plant Physiol (2013), http://dx.doi.org/10.1016/j.jplph.2012.11.022

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Chloroplast dedifferentiation and redifferentiation occurs in fully developed vegetative tissues without meristem formation in PDT plants. However, to study chloroplast biogenesis and the formation of the photosynthetic apparatus, dark-germinated plants are often analyzed, which are then illuminated under controlled conditions. Chl biosynthesis and plastid development are lightregulated in angiosperm plants. Therefore, in the absence of light, etioplasts develop instead of chloroplasts. However, this process is relatively slow and requires at least 3–4 days of dark growth (Gunning, 1965; reviewed by Solymosi and Schoefs, 2010). The inner membrane material is usually stored in hexagonal, paracrystalline prolamellar bodies (PLBs) and lamellar prothylakoids (PTs). A step of the Chl biosynthetic pathway, i.e. the transformation of protochlorophyllide (Pchlide) into chlorophyllide (Chlide) is light-activated (reviewed by Schoefs and Franck, 2003; Solymosi and Schoefs, 2010). The reaction is driven by the photoenzyme NADPH:Pchlide oxidoreductase (POR, E.C. 1.3.1.33.), which is accumulated mainly in the prolamellar bodies of dark-grown seedlings (Ryberg and Dehesh, 1986). In the dark, Chl biosynthesis is arrested at Pchlide, which is present in different molecular environments within the etioplasts membranes. Four different molecular populations of Pchlide have been identified with fluorescence spectroscopy at 77 K (Böddi et al., 1992). Due to the interaction of the pigment molecules with other Pchlide molecules (pigment aggregation), with the POR protein and/or other membrane components, e.g. NADPH or NADP+ , the fluorescence emission spectrum of Pchlide is complex. Different chromophore populations – referred to as spectral forms – can be distinguished. Monomer, non-enzyme bound Pchlide molecules with a fluorescence emission maximum at 633 nm have been identified in the PTs (Ryberg and Sundqvist, 1982; Böddi et al., 1989). The pigments of this form are nonphotoactive, i.e. they cannot be transformed immediately into Chlide upon illumination. The presence of this form is characteristic for proplastids of young, developing tissues that contain only PTs (reviewed by Solymosi and Schoefs, 2008). However, the majority of Pchlide molecules is bound to the POR enzyme, which, together with the co-substrate NADPH, forms photoactive ternary complexes (Griffiths, 1978). These ternary complexes can aggregate, and dimers and oligomers are formed with fluorescence emission maxima at 644 and 655 nm, respectively (Böddi et al., 1989, 1992; Wiktorsson et al., 1993; Ouazzani-Chahdi et al., 1998). The enzyme-bound Pchlide molecules are flash-photoactive, i.e. they transform into Chlide even upon 10 ps illumination (Dobek et al., 1981). This process is clearly indicated by the disappearance of the fluorescence bands at 644 and 655 nm and by the appearance of Chlide fluorescence bands at 684 and 690 nm in the fluorescence emission spectra (Böddi et al., 1991). After illumination, the fluorescence emission maximum gradually shifts towards the blue (this process was first described with absorption spectroscopy by Shibata (1957)). The end-product of this process has an emission maximum at 680 nm (Smeller et al., 2003; Solymosi et al., 2007). This spectral shift indicates the disaggregation of the pigments and the POR macrodomains and is characteristic mostly for etiolated leaves. After this disaggregation, the Chlide molecules are released from POR subunits, bind to the chlorophyll synthase enzyme and are esterified, the results of which is the formation of Chl-a (reviewed by Solymosi and Schoefs, 2010). Investigations upon rehydration of desiccated X. humilis leaves in the dark and in the light have shown that the syntheses of D1 protein, digalactosyl-diacylglycerol synthase 1 and the chlorophyllbinding protein Lhcb2 are light-dependent and it is necessary for granum formation. The initial assembly of stromal membranes occurs independently of light and the recovery of photosynthetic activity does not require de novo Chl biosynthesis (Ingle et al., 2008). However, in these studies, only the initial phase (the first 51 h)

of rehydration was analyzed and the transformation of the desiccoplasts into pregranal plastids was described. The formation of etioplasts with PLBs usually requires a longer continuous period of dark-growth (at least 72 h) in etiolated seedlings (reviewed by Solymosi and Schoefs, 2008). In this work, the transformation of the desiccoplasts was studied in detached leaves of air-dried X. humilis, rehydrated in the dark for 2 weeks at the longest and then illuminated for 72 h. Using 77 K fluorescence spectroscopy and transmission electron microscopy, the final steps of Chl biosynthesis and plastid differentiation processes were studied. Materials and methods Plant material and sample treatment The Xerophyta humilis (Bak.) Dub. and Schinz (Velloziaceae) plants were collected from Barakalalo National Park (Limpopo Province, South Africa) and were maintained in a glasshouse as previously described (Dace et al., 1998). Plants were dehydrated by withholding water from the soil, allowing them to dry naturally under ambient conditions in a glasshouse. Air-dried leaves (5% RWC) were collected and posted under anhydrous conditions to ˝ Hungary. However, due to different plant materials used Gödöllo, in the experiments and in the different repetitions, the length of rehydration slightly varied among the samples. For each experiment, 8–10 leaves with 10–15 cm lengths were soaked in tap water and aerated with an aquarium pump in the dark. Samples were collected after 1 h, 1 week and 2 weeks of rehydration as follows: after 1 h, samples were selected from 5 different leaves and segments from the same leaves were used for pigment determination, fluorescence spectroscopic, and electron microscopic investigations. After 1 week dark rehydration, the samples were selected for further experiments with the help of fluorescence spectroscopy, i.e. the 77 K fluorescence emission spectra of leaf segments with 2 mm × 15 mm size were measured. Leaves showing no regeneration were discarded, while those showing emission bands of protochlorophyllide (Pchlide) were analyzed by measuring their pigment contents and studying their ultrastructures. Five regenerated leaves were selected this way, the remaining segments of which were rehydrated for an additional week. Samples were collected from these 2-week-rehydrated leaves the same way and for the same tests as above. All of these experiments were repeated 3 times starting every time with dry leaves. Thus, the results presented in this paper are based on studies of 15 regenerated leaves. Due to the heterogeneity of the leaves, the regeneration of a few leaves was faster; their emission spectra were similar already after 3–4 days rehydration to those of leaves rehydrated for 2 weeks presented in this work. Since this phenomenon was found occasionally, no further analyses were done on these leaves. In some cases, the leaf blades were heterogeneous and they had segments that did not regenerate; these leaves were discarded. After 2 weeks of rehydration in the dark, several leaves were transferred to white (Tungsten lamp) light of 250 ␮mol photons m−2 s−1 for 72 h continuous illumination. Five leaf pieces were measured in each experiment. All samples showed the same tendencies. The figures show representative results. Fluorescence spectroscopy The 77 K fluorescence spectra were measured with a Fluoromax3 (Jobin Yvon-Horiba, France) spectrofluorometer equipped with a low-temperature accessory as in Solymosi and Böddi (2006). The

Please cite this article in press as: Solymosi K, et al. Desiccoplast–etioplast–chloroplast transformation under rehydration of desiccated poikilochlorophyllous Xerophyta humilis leaves in the dark and upon subsequent illumination. J Plant Physiol (2013), http://dx.doi.org/10.1016/j.jplph.2012.11.022

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excitation wavelengths were 440 nm or 460 nm. The excitation and emission slits were set to 2 nm and 5 nm, respectively. The integration time was 0.1 s. In every case, three spectra were recorded and automatically averaged. The spectra were analyzed with the SPSERV V3.14 program (copyright C. Bagyinka, Institute of Biophysics, Biological Research Centre of the Hungarian Academy of Sciences, Szeged, Hungary). Baseline correction and a combination of three-point and five-point linear smoothing were carried out. The spectra were corrected for the wavelength-dependent variations of the detector’s response. Leaf pieces of 2 mm × 15 mm size were fixed in the sample holder of the spectrofluorometer and immersed in liquid nitrogen for the fluorescence measurements. All manipulations were done in dim green safelight, which was previously tested and did not cause phototransformation of Pchlide. After measuring the 77 K fluorescence spectra, the samples were warmed to −20 ◦ C and then illuminated with white light of 250 ␮mol photons m−2 s−1 for 10 s to test the photoactivity of the different Pchlide forms. The samples were kept in the dark for 10 s after the end of this illumination period, were frozen in liquid nitrogen and the fluorescence emission spectra were registered again. Pigment extraction In some experiments, the pigments were extracted as follows: Dry leaves were soaked in tap water for 1 h, and then the water was removed from their surface with filter paper. The fresh weight was measured and the pigments were extracted with 80% acetone in a mortar. For rehydrated leaves, the pigment extraction was done in dim green light (see above). The extractions were filtered and their fluorescence emission spectra were recorded with 430 or 460 nm excitation. The pigment contents were determined with the help of calibration curves as described in Skribanek et al. (2008). Due to the heterogeneity of the leaves, the pigment content values showed substantial data scattering and could therefore provide only qualitative information.

Fig. 1. 77 K fluorescence emission spectra of desiccated Xerophyta humilis leaves rehydrated in the dark for 1 or 2 weeks and then illuminated for 3 days. A: desiccated leaf, B: leaf after 1 week of rehydration in the dark (basal part of the leaf is shown, however, similar spectrum is found in the apical leaf part), C and D: after 2 weeks of rehydration in the dark, C: apical part of the leaf, D: basal part of the leaf, E: desiccated leaf rehydrated in the dark for 2 weeks and illuminated for 3 days with white light of 250 ␮mol photons m−2 s−1 . The spectra were normalized to their maxima. (The original amplitude of the maximum of spectrum B was one tenth of that of spectra C and D.) Excitation wavelength: 440 nm.

Electron microscopy For transmission electron microscopy, at least 3 different leaf pieces were cut and fixed in 2.5% glutaraldehyde for 3 h in the dark and postfixed in 1% OsO4 for 2 h as in Solymosi et al. (2006a). After dehydration in alcohol series, the samples were embedded in Durcupan ACM resin (Fluka Chemie AG). For all other details of electron microscopic sample preparation see Solymosi et al. (2006a). Three different ultrathin sections were investigated for each sample. 60 pictures were analyzed and for plastid size calculations the standard deviation is presented. The pictures had the same characteristics in each treatment, i.e. the plastid differentiation was synchronized by the rehydration. Therefore, the electron micrographs in Fig. 2 show representative developmental stages of plastids. Results Desiccoplasts in the air-dried leaves No Chl or Pchlide emission bands were found in the fluorescence emission spectra after 1 h of rehydration in the dark or in the light. The recorded signal was a light scattering curve in the 580–780 nm region (Fig. 1, curve A). Despite of the absence of the emission bands in the spectra of the leaf pieces, trace amounts of Chl-a (0.5–2.0 ␮g g−1 fresh mass (FM)) were detected: a band of very low amplitude was observed at 670 nm. The amplitude of this signal was close to the detection limit of the fluorometer.

The electron micrographs indicated the presence of small (average diameter: 2.5 ± 0.5 ␮m) desiccoplasts with large groups of electrondense plastoglobuli and a poorly developed inner membrane system. Concentrically arranged vesicles were often found among the few inner membranes (Fig. 2A), but sometimes almost no membranes and only plastoglobuli were observed (Fig. 2B). Vacuoles with intact tonoplast membranes and mitochondria were also easily distinguishable in the cells as in Sherwin and Farrant (1998), Farrant et al. (1999) and in Farrant (2000). Desiccoplast–proplastid/etioplast transformation in leaves after 1 week rehydration in the dark After 1 week of rehydration in the dark, a fluorescence emission band at 634 nm, characteristic of Pchlide, appeared (Figs. 1, curve B and 3A, solid line) in the spectra of each segment of the leaves. The amplitude of this band was very low; it was about 10 times lower than those in spectra of the 2-week rehydrated leaves (see below). For better demonstration, this spectrum was normalized to its maximum. No fluorescence emission bands corresponding to Chl pigments were detected even in this magnified spectrum. When the excitation was shifted to 460 nm, an emission band was found at 642 nm and a shoulder at 655 nm (Fig. 3, inset, solid line). The Pchlide forms with emission maxima at 634 and 642 nm were non-photoactive, while the amplitude of the 655 nm band slightly decreased upon illumination (however, no significant increase was found above 670 nm upon illumination) (Fig. 3A, dashed lines). The plastids of the dark samples contained

Please cite this article in press as: Solymosi K, et al. Desiccoplast–etioplast–chloroplast transformation under rehydration of desiccated poikilochlorophyllous Xerophyta humilis leaves in the dark and upon subsequent illumination. J Plant Physiol (2013), http://dx.doi.org/10.1016/j.jplph.2012.11.022

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Fig. 2. Electron micrographs of desiccated leaves of Xerophyta humilis after 1 h (A and B), 1 week (C and D), 2 weeks (E–G) of rehydration in the dark in tap water and after subsequent illumination for 3 days with white light of 250 ␮mol photons m−2 s−1 (H). A: typical desiccoplasts with concentrically arranged vesicles and clusters of electrondense plastoglobuli in well-preserved cytoplasm. B: section of a desiccoplast with plastoglobuli, no inner membranes can be observed in the plastid. C: plastid with single prothylakoid membranes and plastoglobuli. D: plastid with developing prothylakoids and with a few bithylakoids indicated by arrows and shown in the inset. E and F: Etio-desiccoplasts from the apical part of the leaf. G: etio-desiccoplast from the basal part of the leaf. Prolamellar bodies (E and G), perforated prothylakoids (E–G) and starch grains (G) are characteristic. H: typical chloroplast with stroma thylakoids and low grana. The bars indicate 1 ␮m. Cw: cell wall, M: mitochondrium, N: nucleus, Pg: plastoglobuli, Plb: prolamellar body, Pt: prothylakoids, RER: rough endoplasmic reticulum, S: starch, V: vacuole, Ve: vesicle.

Please cite this article in press as: Solymosi K, et al. Desiccoplast–etioplast–chloroplast transformation under rehydration of desiccated poikilochlorophyllous Xerophyta humilis leaves in the dark and upon subsequent illumination. J Plant Physiol (2013), http://dx.doi.org/10.1016/j.jplph.2012.11.022

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Etioplast formation in leaves after 2 weeks of rehydration in the dark After 2 weeks of rehydration, intense Pchlide bands at 634 and 655 nm were found in the fluorescence emission spectra (Figs. 1, curves C and D, 3B and C, solid lines). The amplitude ratio of the fluorescence bands at 634/655 nm was different in the basal and apical parts of the leaves (compare curves C and D in Fig. 1). The basal parts (Fig. 1, curve D) accumulated more photoactive Pchlide forms with an emission maximum at 655 nm than the apical sections (Fig. 1, curve C). After illumination, the band at 655 nm disappeared and a new emission band appeared at 679–680 nm observed with 440 nm excitation (Fig. 3B and C, dashed lines), showing the production of Chlide. The excitation with 460 nm indicated that the non-photoactive Pchlide form with emission maximum at 642 nm found already in the spectra of 1week-rehydrated leaves (Fig. 3A, inset) was still present; and the production of the 690 nm (Fig. 3B, inset) and shorter wavelength (Fig. 3C, inset) emitting Chlide forms was found. The broadness of these latter emission bands and the differences in their maxima indicated the spectral heterogeneity of the newly formed Chlide. The plastids of leaves rehydrated for 2 weeks in the dark often contained PLB-like structures (Fig. 2E and G), which were, however, not so regular as those of dark-germinated Poaceae seedlings (Gunning, 1965; reviewed by Solymosi and Schoefs, 2008). PT lamellae, which were often perforated, stretched out from the PLBs (Fig. 2E and G). The plastids of the apical part of the leaves (Fig. 2E and F) had light stroma but did not contain starch grains. In this region, many etioplasts contained clusters of large plastoglobuli (Fig. 2E) characteristic of the desiccoplasts. In the basal part of the leaves, the stroma of the plastids was relatively electron-dense. Thus, the membranes were not always very easily distinguishable. However, the central PLB and the radial PT membranes were characteristic in these plastids and they also often contained starch (Fig. 2G). The average plastid size did not change during rehydration in the dark, but remained 2.5 ± 0.5 ␮m in both leaf segments. Compared with the very small (trace) Pchlide content of leaves rehydrated for 1 week, the Pchlide content increased to 5–7 ␮g g−1 FM in samples collected after 2 weeks of rehydration, while the Chl content remained unchanged (0.5–2 ␮g g−1 FM). Etioplast–chloroplast transformation in dark-rehydrated and subsequently illuminated leaves

Fig. 3. 77 K fluorescence emission spectra of desiccated Xerophyta humilis leaves rehydrated for 1 week (A) or 2 weeks (B and C) in the dark (solid lines) and those of the same leaves illuminated for 10 s with white light of 250 ␮mol photons m−2 s−1 (broken lines). B: spectrum of the apical part of a leaf, C: spectrum of the basal part of a leaf. The spectra were recorded with 440 nm (main figures), and 460 nm (insets) excitation wavelengths and were normalized at their maxima. A: The maximal amplitudes of the signals were very low (about 10 times lower than those shown in other samples) in the experimental spectra.

several large, electron-dense plastoglobuli and single, often perforated PT membranes (Fig. 2C and D). Interestingly, in some cases bithylakoids were also observed (Fig. 2D, inset). No membrane vesicles were present in these plastids, but no changes were observed in the average plastid diameter (2.5 ± 0.5 ␮m) when compared to those of air-dried leaves. These leaves contained 0.5–2.0 ␮g g−1 FM Chl (similarly to the dry leaves) and traces of Pchlide were detected.

When the 2-week-rehydrated leaves were exposed to continuous light of 250 ␮mol photons m−2 s−1 for 72 h, fluorescence emission bands were observed at 685, 695 and 730 nm, characteristic for green leaves (Fig. 1, curve E). Regular chloroplasts developed in these leaves with plenty of stacked thylakoid membranes (grana). Due to the high electron density of their stroma, however, the contrast of these membranes was poor (Fig. 2H). Plastid size increased during illumination (average length: 3.3 ± 0.8 ␮m, average width: 0.9 ± 0.3 ␮m), and plastids became more ovale (average length/width ratio: 0.3). Starch grains were often observed in these plastids. During the 3 days of illumination, the Chl content increased to about 400–450 ␮g g−1 FM, which was obviously enough for the formation of regular chloroplasts. Further accumulation was observed during 7 days illumination; about 1200 ␮g g−1 FM Chl was found, but no remarkable changes took place in the chloroplast ultrastructure or in the fluorescence spectra. Discussion Considering the global importance of PDT plants (Alpert, 2000; Porembski, 2007), the physiology of the DT-tolerant vegetative

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organs is of great importance. The fast recovery of photosynthesis during rehydration allows important production rates, even in very dry seasonally long (5–10/11 months) areas. The special feature of the leaves of these plants is the formation of desiccoplasts (Tuba et al., 1993b). The question can be raised about their physiological role in the regreening leaves during rehydration. In this work, detached leaves which had fully differentiated and no meristematic cells were used. Thus, the carrying on of proplastids and their pigments via cell division from meristematic cells, which occurs, for example, during embryo development (Böddi et al., 1999) or bud break (Solymosi et al., 2006a, 2012), can be excluded. The results described here show the regeneration of cells and plastids and de novo synthesis of different compounds (pigments and plastid inner membranes) during the rehydration process in fully differentiated tissues. The fluorescence measurements here (Fig. 1, curve A) show that the photosynthetic pigments decompose during the degreening process, as reported previously (Tuba et al., 1996; Farrant, 2000), which is very important for the photoprotection of the chlorenchymatic tissues. The presence of chlorophyllous pigments in photochemically inactive structures would provoke photodamage via production of reactive oxidative substances (ROS) (Farrant et al., 2003). The high density of plastoglobuli in the desiccoplasts of dry (inactive) and rehydrated (transforming) desiccoplasts brings up the possibility that they contain storage materials and precursors, which can be used directly in the formation of PLB and PT membranes in the dark and of thylakoid membranes in the light. On the other hand, plastoglobuli may play a role in photoprotection during light stress (reviewed by Bréhélin et al., 2007). The early phases of plastid differentiation (observed after 1 week of dark rehydration, Fig. 2C and D) are in good agreement with data obtained by Ingle et al. (2008). However, in this work, the accumulation of non-photoactive Pchlide pigments was observed after 1 week of rehydration (Figs. 1 and 3), indicating de novo pigment (i.e. Chl) biosynthesis after this period. Furthermore, occasionally the stacking of two thylakoids (that might be considered as low grana or bithylakoids) was also observed (Fig. 2D, inset). The transformation of desiccoplasts into pregranal plastids (or poorly developed etioplasts) first with few PT-s only (Fig. 2C and D), and then to etioplasts with PLBs (Fig. 2E and G) resembles the proplastid–etioplast transformation process (reviewed by Solymosi and Schoefs, 2008). Similar, poorly developed etioplasts are present also in epicotyls of dark-geminated pea (Böddi et al., 1994), in innermost leaf primordia of buds (Solymosi et al., 2006a, 2012) and cabbage head (Solymosi et al., 2004). The formation of Pchlide forms during the desiccoplast–etioplast pathway follows the general rules observed in leaves of very young seedlings, i.e. first the short-wavelength (here 634 nm emitting), non-photoactive Pchlide form is produced (Schoefs et al., 1994, 2000) (Figs. 1, curve B and 3A), then the gradual accumulation of the flash-photoactive 655 nm emitting form proceeds (Figs. 1, curves B–D, 3B and C). A unique feature of the etioplasts relative to desiccoplasts is the appearance of the 642 nm emitting, non-photoactive form (Fig. 3A). It cannot be a variant of the Pchlide dimer with emission maximum at 644 nm described earlier in leaves of etiolated seedlings because the latter was proven to be fully photoactive (Böddi et al., 1991). The 642 nm emitting and non-photoactive Pchlide form resembles the POR-Pchlide-NADP+ dimer (Franck et al., 1999). The presence of this form indicates a shortage of NADPH in desiccated and dark-rehydrated leaves. This shortage can also be the reason for the loose structure of PLBs observed in this work because NADPH has an important structural role in maintaining the highly regular structure of the PLBs (Ryberg and Sundqvist, 1988; Solymosi et al., 2006b). Another possibility is that the pigment in this form is a

Pchlide ester that is not accepted as substrate by the POR enzyme (Griffiths, 1980). However, Pchlide esters are present in very low amounts and in monomeric forms having emission maxima at 628 or 633 nm in etiolated leaves (reviewed by Solymosi and Schoefs, 2008). There was an increasing gradient in the relative amounts of the flash-photoactive 655 nm emitting Pchlide form from the tip towards the base of leaves (Figs. 1 and 3), but this gradient appeared only in 2-week-old dark-rehydrated leaves. A similar gradient is common in etiolated leaves of seedlings from the Poaceae family. It is in connection with the different ages of cells along the leaf blades; in the basal region, the cells are young and the eldest cells are located on the tip (Robertson and Laetsch, 1974; reviewed by Solymosi and Schoefs, 2008). The amount of the 655 nm emitting form has an optimum during the dark development. In very young cells, the amount of POR and Pchlide is low; no POR oligomers or PLBs are formed (Klein and Schiff, 1972; reviewed by Solymosi and Schoefs, 2008). In senescent cells, however, the PLB membranes may be loose or decomposed and the POR oligomers disintegrate, too. The test plant in this work, X. humilis, has base-growing leaves i.e. with younger cells present at the leaf base. The high metabolic activity of these cells is also illustrated by the appearance of starch grains in the etioplasts of the basal cells (Fig. 2G), also observed by other authors after short periods of dark-rehydration (Ingle et al., 2008). The appearance of the starch grains indicates that the desiccated leaves contain storage material, which is used in the rehydration process, when the biochemical mechanisms become active and can regenerate the cell components. Since no starch grains were found in the desiccoplasts of fully dehydrated leaves, and no photosynthetic starch can be produced in the dark, their appearance needs a complex metabolic network, i.e. starch formation from other stored materials. These materials could be stored in the plastoglobuli, the number of which decreased during the rehydration process (Fig. 2F and G), or more likely, they could derive from osmolytes characteristic for this species (Farrant et al., 1999; Farrant, 2000).

Fig. 4. Schematic representation of plastid development occurring during rehydration of dry Xerophyta humilis leaves. 1: desiccoplast stage in dry leaves with vesicular inner membranes and clusters of plastoglobuli. 2: beginning of (pro)thylakoid organization: perforated lamellar sheets are formed, occasionally the stacking of two membrane sheets can be observed, this step is similar for plastid development occurring in the dark or in the light. 3: chloroplasts are formed with grana and stroma thylakoids in the light (in case of Xerophyta rather low grana are characteristic for the chloroplasts). 4: etioplasts develop in the dark from desiccoplasts and have prolamellar bodies and prothylakoids. Upon illumination, the transformation of etioplasts into chloroplasts occurs.

Please cite this article in press as: Solymosi K, et al. Desiccoplast–etioplast–chloroplast transformation under rehydration of desiccated poikilochlorophyllous Xerophyta humilis leaves in the dark and upon subsequent illumination. J Plant Physiol (2013), http://dx.doi.org/10.1016/j.jplph.2012.11.022

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Upon illumination of the dark-rehydrated leaves, we observed the formation of the photosynthetic apparatus, the appearance of PSI and PSII fluorescence and the disappearance of Pchlide (Fig. 1, curve E). In parallel, the etioplast–chloroplast transformation was also observed (Fig. 2H). Similarly, chloroplast formation was observed when the leaves were rehydrated in the light (Tuba et al., 1993a,b; Dace et al., 1998; Ingle et al., 2008). The observed plastid differentiation patterns studied in the differentiated cells of the vegetative photosynthetic tissues of dehydrated Xerophyta leaves are summarized in Fig. 4. The indicated stages of chloroplast redifferentiation (i.e. the desiccoplast–chloroplast transformation involving pregranal plastids, Fig. 4, stages 1→2→3) are well-known from literature data (e.g. Ingle et al., 2008) and are in good agreement with data presented for the gradual dedifferentiation of chloroplasts upon dehydration of Xerophyta villosa (Hallam and Luff, 1980; Sherwin and Farrant, 1998). During dehydration, chloroplast dedifferentiation occurs through similar stages, but in the opposite direction (Fig. 4, stages 3→2→1). Thus, the transformations indicated with arrows are in this case reversible. Stages 1 and 2 are similar to the initial steps of proplastid–chloroplast transition (described also by Ingle et al., 2008); the vesicles and single thylakoids can be also considered as primary grana and the plastids called pregranal plastids (Whatley, 1977, 1979). However, in PDT plants, chloroplast dedifferentiation and redifferentiation, chlorophyll breakdown and resynthesis occur in a fully developed vegetative tissue without simultaneous meristematic dedifferentiation and redifferentiation of the cells. In these plants, the above-mentioned processes are induced merely by desiccation and rehydration, therefore these plastids are termed desiccoplasts (Tuba et al., 1994, 1996) if differentiated in the light. In this work, we have proved that after sufficiently long dark rehydration periods (e.g. 1 or 2 weeks or earliest after 3 days) also the desiccoplast–etioplast differentiation pathway can be observed (Fig. 4, stages 1→2→4). The etioplasts (or etio-desiccoplasts) transform into chloroplasts upon illumination (Fig. 4, stages 4→3) similarly to processes observed during the greening of etiolated plants. The results of this work show a new phenomenon: the fully developed vegetative cells/tissue of desiccated PDT X. humilis leaves have the ability to carry out plastid differentiation, and their plastids have high metabolic flexibility by which they can adapt to different environmental conditions. Acknowledgments The authors are grateful to Prof. Jill Farrant (University of Cape Town) for providing us with the plant materials, to Csilla Jónás and Katalin M. Gergely (Eötvös University) for their helpful assistance in the electron microscopic sample preparation, and to Gyula Rabnecz (Szent István University) for transportation of the plant material. The second author of this work died on 4 July 2009, and we dedicate this paper to his memory. References Alpert P. The discovery, scope, and puzzle of desiccation tolerance in plants. Plant Ecol 2000;151:15–7. Böddi B, Lindsten A, Ryberg M, Sundqvist C. On the aggregational states of protochlorophyllide and its protein complexes in wheat etioplasts. Physiol Plant 1989;76:135–43. Böddi B, Ryberg M, Sundqvist C. The formation of a short-wavelength chlorophyllide form at partial phototransformation of protochlorophyllide in etioplast inner membranes. Photochem Photobiol 1991;53:667–73. Böddi B, Ryberg M, Sundqvist C. Identification of four universal protochlorophyllide forms in dark-grown leaves by analyses of the 77 K fluorescence emission spectra. J Photochem Photobiol B: Biol 1992;12:389–401. Böddi B, McEwen B, Ryberg M, Sundqvist C. Protochlorophyllide forms in non-greening epicotyls of dark-grown pea (Pisum sativum). Physiol Plant 1994;92:160–70.

7

Böddi B, Lindsten A, Sundqvist C. Chlorophylls in dark-grown epicotyl and stipula of pea. J Photochem Photobiol B: Biol 1999;48:11–6. Bréhélin C, Kessler F, van Wijk KJ. Plastoglobules: versatile lipoprotein particles in plastids. Trends Plant Sci 2007;12:260–6. Dace H, Sherwin HW, Illing N, Farrant JM. Use of metabolic inhibitors to elucidate mechanisms of recovery from desiccation stress in the resurrection plant Xerophyta humilis. Plant Growth Regul 1998;24:171–7. Dobek A, Dujardin E, Franck F, Sironval C, Breton J, Roux E. The first events of protochlorophyll(ide) photoreduction investigated in etiolated leaves by means of the fluorescence excited by short, 610 nm laser flashes at room temperature. Photobiochem Photobiophys 1981;2:35–44. Farrant JM. A comparison of mechanisms of desiccation tolerance among three angiosperm resurrection plant species. Plant Ecol 2000;151:29–39. Farrant JM, Cooper K, Kruger LA, Sherwin HW. The effect of drying rate on the survival of three desiccation-tolerant angiosperm species. Ann Bot 1999;84: 371–9. Farrant JM, Vander Willigen C, Loffell DA, Bartsch S, Whittaker A. An investigation into the role of light during desiccation of three angiosperm resurrection plants. Plant Cell Environ 2003;26:1275–86. Franck F, Bereza B, Böddi B. Protochlorophyllide-NADP+ and protochlorophyllideNADPH complexes and their regeneration after flash illumination in leaves and etioplast membranes of dark-grown wheat. Photosynth Res 1999;59: 53–61. Gaff DF. Desiccation tolerant flowering plants in southern Africa. Science 1971;174:1033–4. Griffiths WT. Reconstitution of chlorophyll formation by isolated etioplast membranes. Biochem J 1978;174:681–92. Griffiths WT. Substrate-specificity studies on protochlorophyllide reductase in barley (Hordeum vulgare) etioplast membranes. Biochem J 1980;186:267–78. Gunning BES. The greening process in plastids, 1. The structure of the prolamellar body. Protoplasma 1965;60:111–30. Hallam ND, Luff SE. Fine structural changes in the mesophyll tissue of the leaves of Xerophyta villosa during desiccation. Bot Gaz 1980;141:173–9. Hambler DJ. A poikilohydrous, poikilochlorophyllous angiosperm from Africa. Nature 1961;191:1415–6. Ingle RA, Collett H, Cooper K, Takashi Y, Farrant JM, Illing N. Chloroplast biogenesis during rehydration of the resurrection plant Xerophyta humilis: parallels to the etioplast–chloroplast transition. Plant Cell Environ 2008;31: 1813–24. Klein S, Schiff JK. The correlated appearance of prolamellar bodies, protochlorophyllide species, and the Shibata shift during development of bean etioplasts in the dark. Plant Physiol 1972;49:619–26. Moore JP, Le NG, Brandt WF, Driouich A, Farrant JM. Towards a systems-based understanding of plant desiccation tolerance. Trends Plant Sci 2009;14:110–7. Oliver MJ, Tuba Z, Mishler BD. The evolution of vegetative desiccation tolerance in land plants. Plant Ecol 2000;151:85–100. Ouazzani-Chahdi MA, Schoefs B, Franck F. Isolation and characterisation of photoactive complexes of NADPH:protochlorophyllide oxidoreductase from wheat. Planta 1998;206:673–80. Porembski S. Tropical inselbergs: habitat types, adaptive strategies and diversity patterns. Rev Bras Bot 2007;30:579–86. Proctor MCF, Tuba Z. Poikilohydry and homoihydry: antithesis or spectrum of possibilities? Tansley review no. 141. New Phytol 2002;156:327–49. Rascio N, La Rocca N. Resurrection plants: the puzzle of surviving extreme vegetative desiccation. Crit Rev Plant Sci 2005;24:209–25. Robertson D, Laetsch WM. Structure and function of developing barley plastids. Plant Physiol 1974;54:148–59. Ryberg M, Dehesh K. Localization of NADPH:protochlorophyllide oxidoreductase in dark-grown wheat (Triticum aestivum) by immuno-electron microscopy before and after transformation of the prolamellar bodies. Physiol Plant 1986;66:616–24. Ryberg M, Sundqvist C. Spectral forms of protochlorophyllide in prolamellar bodies and prothylakoids fractionated from wheat etioplasts. Physiol Plant 1982;56:133–8. Ryberg M, Sundqvist C. The regular ultrastructure of isolated prolamellar bodies depends on the presence of membrane-bound NADPH-protochlorophyllide oxidoreductase. Physiol Plant 1988;73:218–26. Schoefs B, Franck F. Protochlorophyllide reduction: mechanisms and evolution. Photochem Photobiol 2003;78:543–57. Schoefs B, Garnir HP, Bertrand M. Comparison of the photoreduction of protochlorophyllide to chlorophyllide in leaves and cotyledons from dark-grown bean as a function of age. Photosynth Res 1994;41:405–17. Schoefs B, Bertrand M, Franck F. Spectroscopic properties of protochlorophyllide analyzed in situ in the course of etiolation and in illuminated leaves. Photochem Photobiol 2000;72:85–93. Sherwin HW, Farrant JM. Protection mechanisms against excess light in the resurrection plants Craterostigma wilmsii and Xerophyta viscosa. Plant Growth Regul 1998;24:203–10. Shibata K. Spectroscopic studies on chlorophyll formation in intact leaves. J Biochem 1957;44:147–73. Skribanek A, Solymosi K, Hideg É, Böddi B. Light and temperature regulation of greening in dark-grown ginkgo (Ginkgo biloba L.). Physiol Plant 2008;134: 649–59. Smeller L, Solymosi K, Fidy J, Böddi B. Activation parameters of the blue shift (Shibata shift) subsequent to protochlorophyllide phototransformation. Biochim Biophys Acta 2003;1651:130–8.

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G Model JPLPH-51670; No. of Pages 8 8

ARTICLE IN PRESS K. Solymosi et al. / Journal of Plant Physiology xxx (2013) xxx–xxx

Solymosi K, Böddi B. Optical properties of bud scales and protochlorophyll(ide) forms in leaf primordia of closed and opened buds. Tree Physiol 2006;26: 1075–85. Solymosi K, Schoefs B. Prolamellar body: a unique plastid compartment, which does not only occur in dark-grown leaves. In: Schoefs B, editor. Plant cell organelles – selected topics. Kerala: Research Sign Post; 2008. p. 151–202. Solymosi K, Schoefs B. Etioplast and etio-chloroplast formation under natural conditions – the dark side of chlorophyll biosynthesis. Photosynth Res 2010;105:143–66. Solymosi K, Martinez K, Kristóf Z, Sundqvist C, Böddi B. Plastid differentiation and chlorophyll biosynthesis in different leaf layers of white cabbage (Brassica oleracea cv. capitata). Physiol Plant 2004;121:520–9. Solymosi K, Bóka K, Böddi B. Transient etiolation: protochlorophyll(ide) and chlorophyll forms in differentiating plastids of closed and breaking leaf buds of horse chestnut (Aesculus hippocastanum). Tree Physiol 2006a;26: 1087–96. Solymosi K, My´sliwa-Kurdziel B, Bóka K, Strzałka K, Böddi B. Disintegration of the prolamellar body structure at high concentrations of Hg2+ . Plant Biol 2006b;8:627–35. Solymosi K, Smeller L, Ryberg M, Sundqvist C, Fidy J, Böddi B. Molecular rearrangement in POR macrodomains as a reason for the blue shift of chlorophyllide fluorescence observed after phototransformation. Biochim Biophys Acta 2007;1768:1650–8. Solymosi K, Morandi D, Bóka K, Böddi B, Schoefs B. Plastids, photosynthetic pigments and pigment forms in leaf primordia of buds with various structures and different developmental stages. Planta 2012;235:1035–49.

Tuba Z, Lichtenthaler HK, Csintalan Z, Pócs T. Regreening of desiccated leaves of the poikilochlorophyllous Xerophyta scabrida upon re-hydration. J Plant Physiol 1993a;142:103–8. Tuba Z, Lichtenthaler HK, Maróti I, Csintalan Z. Resynthesis of thylakoids and chloroplast ultrastructure in the desiccated leaves of the poikilochlorophyllous plant Xerophyta scabrida upon re-hydration. J Plant Physiol 1993b;142:742–8. Tuba Z, Lichtenthaler HK, Csintalan Z, Nagy Z, Szente K. Reconstitution of chlorophylls and photosynthetic CO2 assimilation upon rehydration of the desiccated poikilochlorophyllous plant Xerophyta scabrida (Pax) Th. Dur. et Schinz. Planta 1994;192:414–20. Tuba Z, Lichtenthaler HK, Csintalan Z, Nagy Z, Szente K. Loss of chlorophylls, cessation of photosynthetic CO2 assimilation and respiration in the poikilochlorophyllous plant Xerophyta scabrida during desiccation. Physiol Plant 1996;96:383–8. Tuba Z, Smirnoff N, Csintalan Z, Szente K, Nagy Z. Respiration during slow desiccation of the poikilochlorophyllous desiccation tolerant plant Xerophyta scabrida at present-day CO2 concentration. J Plant Physiol Biochem 1997;35:381–6. Vassiljev JM. Über den Wasserhaushalt von Pflanzen der Sandwüste im sudöstliche Kara-Kum. Planta 1931;14:225–309. Whatley JM. Variations in the basic pathway of chloroplast development. New Phytol 1977;78:407–20. Whatley JM. Plastid development in the primary leaf of Phaseolus vulgaris: variations between different types of cell. New Phytol 1979;82:1–10. Wiktorsson B, Engdahl S, Zhong LB, Böddi B, Ryberg M, Sundqvist C. The effect of cross-linking of the subunits of NADPH:protochlorophyllide oxidoreductase on the aggregational state of protochlorophyllide. Photosynthetica 1993;29:205–18.

Please cite this article in press as: Solymosi K, et al. Desiccoplast–etioplast–chloroplast transformation under rehydration of desiccated poikilochlorophyllous Xerophyta humilis leaves in the dark and upon subsequent illumination. J Plant Physiol (2013), http://dx.doi.org/10.1016/j.jplph.2012.11.022