Programmed cell death in plant embryogenesis - Journal of Cell Science

INTRODUCTION

During the last two decades, there has been intensive researchon the mechanisms of programmed cell death (PCD). In theanimal life cycle, the earliest manifestations of PCD canalready be seen during embryogenesis, when some cells haveto be sacrificed for the sake of correct embryonic patternformation. Studies with PCD-defective embryos have shownthat the embryonic pattern formation is disturbed and/or theanimal dies prenatally as soon as the PCD pathway isartificially blocked through mutagenesis or targeted genedisruption (White et al., 1994; Kuida et al., 1996). There aretwo main functions that PCD serves in animal embryonicdevelopment: (1) sculpting the embryo, and (2) removal ofunneeded structures (Jacobson et al., 1997; Song and Steller,1999).

Compared to animal embryogenesis, which involves

extensive morphogenesis including cell movement and, inmany cases, germline formation (Müller, 1997), in plants onlythe apical-basal plan, viz. developmental axis with shoot androot meristems, is established during embryogenesis. Mostmorphogenic processes in higher plants occur at the post-embryonic stages, after seed germination (Goldberg et al.,1994); maintenance of a continuous germline was notestablished during plant evolution (Andrews, 1998). Theseobservations point to a relatively simplified embryonic patternformation in plants compared with animals. Consequently, onewould expect a more limited occurrence of PCD in plantembryogenesis.

To date, no systematic studies have been reported that utilizemolecular and biochemical markers of PCD to investigate theimplication of PCD in plant embryogenesis. Curiously,degradation of terminally differentiated suspensor cells, whosetransient active role is promoting the growth of the embryo

4399Journal of Cell Science 113, 4399-4411 (2000)Printed in Great Britain © The Company of Biologists Limited 2000JCS1834

In the animal life cycle, the earliest manifestations ofprogrammed cell death (PCD) can already be seen duringembryogenesis. The aim of this work was to determine ifPCD is also involved in the elimination of certain cellsduring plant embryogenesis. We used a model system ofNorway spruce somatic embryogenesis, which representsa multistep developmental pathway with two broadphases. The first phase is represented by proliferatingproembryogenic masses (PEMs). The second phaseencompasses development of somatic embryos, which arisefrom PEMs and proceed through the same sequence ofstages as described for their zygotic counterparts.

Here we demonstrate two successive waves of PCD,which are implicated in the transition from PEMs tosomatic embryos and in correct embryonic patternformation, respectively. The first wave of PCD isresponsible for the degradation of PEMs when they giverise to somatic embryos. We show that PCD in PEM cellsand embryo formation are closely interlinked processes,both stimulated upon withdrawal or partial depletion of

auxins and cytokinins. The second wave of PCD eliminatesterminally differentiated embryo-suspensor cells duringearly embryogeny.

During the dismantling phase of PCD, PEM andembryo-suspensor cells exhibit progressive autolysis,resulting in the formation of a large central vacuole.Autolytic degradation of the cytoplasm is accompanied bylobing and budding-like segmentation of the nucleus.Nuclear DNA undergoes fragmentation into both largefragments of about 50 kb and multiples of approximately180 bp. The tonoplast rupture is delayed until lysis of thecytoplasm and organelles, including the nucleus, is almostcomplete. The protoplasm then disappears, leaving acellular corpse represented by only the cell wall. Thispathway of cell dismantling suggests overlapping ofapoptotic and autophagic types of PCD during somaticembryogenesis in Norway spruce.

Key words: Plant embryogenesis, Norway spruce, Programmed celldeath, Cell dismantling

SUMMARY

Two waves of programmed cell death occur during formation and

development of somatic embryos in the gymnosperm, Norway spruce

Lada H. Filonova 1, Peter V. Bozhkov 1,*, Vladimir B. Brukhin 1, Geoffrey Daniel 2, Boris Zhivotovsky 3 andSara von Arnold 1

1Department of Forest Genetics, Uppsala Genetic Centre, Swedish University of Agricultural Sciences, Box 7027, S-75007Uppsala, Sweden 2Department of Wood Science, Swedish University of Agricultural Sciences, Box 7008, S-75007 Uppsala, Sweden3Institute of Environmental Medicine, Division of Toxicology, Karolinska Institutet, Box 210, S-17177 Stockholm, Sweden*Author for correspondence (e-mail: [email protected])

Accepted 5 October; published on WWW 16 November 2000

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proper at early stages of embryogenesis (Yeung and Meinke,1993), is cited in modern literature as one of the typicalexamples of PCD in plant development (Jones and Dangl,1996; Beers, 1997), although this issue was examined solely atthe cytological level, in the mid 1970s to early 80s (Nagl, 1976;Nagl, 1977; Gärtner and Nagl, 1980), and has never been re-examined by employing more specific assays for PCD. Onereason may be the difficulty in isolating a sufficient quantityof embryonic tissue at early stages of embryogenesis to be ableto perform such assays. The early zygotic embryo is minuteand is surrounded by endosperm in angiosperms and bymegagametophyte in gymnosperms. Furthermore, even withinthe same plant, different ovules develop asynchronously(Owens, 1995), which hampers collecting embryos at acommon developmental stage. The latter, however, seems to bean important prerequisite for studying PCD, which is likewisea highly asynchronous process by itself; cells within apopulation may begin PCD at very different times, and thelength of the various phases of morphological change can varyfrom cell to cell (Collins et al., 1997; Messam and Pittman,1998).

Somatic embryogenesis represents one form of asexualplant development that can be induced in callus or somatictissues. This provides the means for versatile control overembryo formation and development, and ultimately forisolation of sufficient quantities of developmentallyhomogenous tissues using molecular and morphologicalstage-specific markers. In this connection, we considered thatsomatic embryogenesis would be an attractive tool forstudying PCD in plant embryogenesis. Previous studies in ourlaboratory defined the whole developmental pathway ofsomatic embryogenesis in the gymnosperm, Norway spruce(Pinaceae), through the use of the time-lapse trackingtechnique (Filonova et al., 2000). This pathway involves twobroad phases, which in turn are divided into more specificdevelopmental stages. The first phase is represented byproliferating proembryogenic masses (PEMs), cell aggregatesthat can pass through a series of three characteristic stagesdistinguished by cellular organization and cell number (stagesPEM I, II and III; Fig. 1), but can never develop directly intoa real embryo. The second phase encompasses developmentof somatic embryos. The latter arise de novo from PEM III,

and then proceed through the same, stereotyped sequence ofstages as described for zygotic embryogeny of Pinaceae(Singh, 1978; Fig. 1). Plant growth regulators (PGRs), auxinsand cytokinins, are necessary during the first phase to maintainPEM proliferation, whereas embryo formation from PEM IIIis triggered by the withdrawal of PGRs. Once early somaticembryos have formed, their further development to matureforms requires abscisic acid (ABA; Fig. 1).

We have found both direct and indirect indications thatcell death is an important component of the describeddevelopmental pathway (Filonova et al., 2000). First, PEMshave been shown to be composed of two types of cell: smalldensely cytoplasmic cells, which stain red with acetocarmine,and enlarged, highly vacuolated cells, which are more or lesselongated and permeable to Evan’s blue (Fig. 1). The latter,being an isomer of the more commonly used dye Trypan blue,can also be used to distinguish viable from dead or dying cells,based on its inability to pass through an intact plasmamembrane (Huang et al., 1986). Secondly, a proper timing ofsuspensor degradation (within a few days after addition ofABA) has been suggested to be an important factor for normalembryo development. For convenience, we use the term‘suspensor’ (or ‘embryo-suspensor’) in this paper to denote amassive structure that is formed during early zygotic andsomatic embryogeny of gymnosperms and classified, in classicembryology, as secondary suspensor (Singh, 1978; vonAderkas et al., 1991). Thirdly, we have found differentialexpression of monoclonal antibody (mAb) JIM13-reactiveepitope of arabinogalactan proteins (AGPs) in the cell walls ofPEMs and somatic embryos. The vast majority of PEM cellsexpressed this epitope, but none of the cells in the early somaticembryos (Filonova et al., 2000). There is increasing evidencethat AGPs are implicated in PCD in plants (Gao and Showalter,1999; Majewska-Sawka and Nothnagel, 2000), and that it isspecifically the JIM13-reactive epitope of AGPs which marksthe cells destined for PCD (Schindler et al., 1995; Stacey et al.,1995). In this context, the obvious question is whether PEMcells of Norway spruce are committed to die.

The present study tries to answer to this question, and alsoshows how the frequency of PCD in PEMs changes in thecourse of sequential treatments that regulate transition fromPEMs (the first phase) to embryos (the second phase).

L. H. Filonova and others

Fig. 1.A schematic representation of thedevelopmental pathway of somaticembryogenesis in Norway spruce(adapted from Filonova et al., 2000; notdrawn to scale). Proliferation of PEMs isstimulated by auxin and cytokinin. Anindividual PEM should pass through aseries of three characteristic stages (I, IIand III) to transdifferentiate to somaticembryos (SE). At stage PEM I, a cellaggregate is composed of a smallcompact clump of densely cytoplasmiccells adjacent to a single enlarged andvacuolated cell. Similar cell aggregates but possessing more than one vacuolated cell have been classified as PEM II. At stage PEM III, anenlarged clump of densely cytoplasmic cells appears loose rather than compact; polarity is disturbed. Withdrawal of plant growth regulators(PGRs) triggers embryo formation from PEM III, whereas abscisic acid (ABA) is necessary to promote further development of somaticembryos through late embryogeny to mature forms. Shown in red are the cells of PEMs and somatic embryos, which stain in situ red withacetocarmine. Blue colour corresponds to the cells of PEMs and somatic embryos, which stain in situ blue with Evan’s blue. Shown as dashedblue lines in the last but one stage of the pathway are the remnants of the degenerated suspensor in the beginning of late embryogeny.

AUXIN +CYTOKININ no PGR

ABAPEM I

PEM II

PEM III

SE

SE

4401Programmed cell death in plant embryogenesis

Furthermore we have examined a spatial distribution of dyingcells in PEMs and in developing embryos. Finally, we suggestthe cytological pathway of cell dismantling during somaticembryogenesis of Norway spruce.

MATERIALS AND METHODS

Cell culture of Norway spruceEmbryogenic cell line 95.88.17 of Norway spruce (Picea abiesL.Karst.), which has previously been used in time-lapse tracking(Filonova et al., 2000), was also chosen for the present study. Cellaggregates (i.e. a mixture of PEMs and somatic embryos) were storedin liquid nitrogen and thawed 5 months prior to the onset ofexperiments (Norgaard et al., 1993). Culture regrowth was achievedby inoculating cell aggregates on solidified half-strength LPproliferation medium (modified after Bozhkov and von Arnold, 1998)containing 9.0 µM auxin, 2,4-dichlorophenoxyacetic acid (2,4-D),and 4.4 µM cytokinin, N6-benzyladenine (BA). Thereafter, 2 monthsprior to the onset of experiments, proliferating cell aggregates weretransferred to the liquid half-strength LP proliferation medium. Astable proliferating embryogenic suspension culture was establishedby weekly subculturing 3 ml of settled cell aggregates into 47 ml freshproliferation medium in 250-ml Erlenmeyer flasks. This routineminimised the risk of occurrence of intraclonal variation in culturemorphology among different flasks. Suspension cultures were grownon a gyratory shaker (100 rpm) in darkness at 22±1°C.

In order ‘to switch’ the culture from proliferation to development(i.e. to induce somatic embryo formation from PEMs), auxin andcytokinin should be withdrawn (Filonova et al., 2000; Fig. 1). This wasaccomplished through two successive washings of 3-ml settled cellaggregates with 10-ml samples of modified full-strength BMI-S1 PGR-free medium (Bozhkov and von Arnold, 1998) in 15-ml Falcon tubesfollowed by inoculation of 3-ml supernatant-free cell aggregates into47 ml BMI-S1 medium of the same composition in 250-ml Erlenmeyerflasks. The cultures were grown on a gyratory shaker (100 rpm) indarkness at 22±1°C for 1 week until subjected to ABA treatmentenabling further somatic embryo development to mature forms.

ABA treatment involved plating 1 ml of settled cell aggregates ontoWhatman no. 2 filter paper (5.5 cm in diameter) placed on solidifiedBMI-S1 maturation medium with 30 µM ABA in 60-mm Petri plates(10 ml medium per plate; Bozhkov and von Arnold, 1998). The filterpapers with developing somatic embryos were transferred to freshmedium after 2, 4 and 6 weeks. The cultures were incubated in thedark at 22±1°C.

Sampling of cell culturesCell cultures were sampled for subsequent analyses (see followingsections) at four sequential time points, with a progressive increase inthe frequency of somatic embryos compared to PEMs (Table 1):(1) 3 days after subculture in auxin- and cytokinin-containingproliferation medium (designated as 3d 2,4-D+BA), (2) at the end ofsubculture in proliferation medium (designated as 7d 2,4-D+BA), (3)at the end of subculture in PGR-free medium (designated as 7d noPGR) and (4) 7 days after plating on ABA-containing medium(designated as 7d ABA). In addition, individual somatic embryos atadvanced stages of development were collected after 2 and 7 weeksof ABA treatment for terminal deoxynucleotidyl transferase (TdT)-mediated dUTP nick-end labeling (TUNEL) assay.

Human apoptotic cellsJurkat cells (human leukemia T cell line, obtained from the EuropeanCollection of Cell Cultures, Salisbury, UK) were grown in RPMI 1640medium supplemented with 10% heat-inactivated fetal calf serum, 2mM glutamine, 100 i.u./ml penicillin and 100 mg/ml streptomycin ina humidified atmosphere of 5% CO2 in air at 37°C. Cells were

maintained in logarithmic growth phase by routine passage every 3-4days. Apoptosis was induced by treatment with 250 ng/ml agonisticanti-CD95 mAb (clone CH-11, Medical & Biological Laboratories,Ltd., Nagoya, Japan). 3 hours after incubation, cells were collectedby centrifugation at 1000 g for 5 minutes and the pellet resuspendedin phosphate-buffered saline (PBS) and subjected to a secondcentrifugation. The supernatant was aspirated and the pellet kept at−70°C before DNA was extracted. The fragmented DNA of Jurkatcells served as a positive control for Norway spruce DNA inelectrophoresis experiments.

Protoplast isolation and fractionationProtoplasts were isolated at four sequential time points (Table 1). Thecell wall digesting enzyme solution consisted of 1% (w/v) Cellulase‘Onozuka R-10’ (Duchefa, Haarlem, The Netherlands), 0.25% (w/v)Macerozyme R-10 (Duchefa, Haarlem, The Netherlands), 0.25%(w/v) Driselase (Sigma, St Louis, MO, USA), 5 mM CaCl2 and 0.4M mannitol, pH 5.8. With suspension cultures (i.e. the first three timepoints), cell aggregates were first collected on the top of an 80-µmnylon screen and then transferred to 90-mm Petri plates containingthe enzyme solution (12 ml for 2 g fresh mass of cell aggregates),whereas 7d ABA-cell aggregates were placed directly in the enzymesolution and then resuspended by using a transfer pipette.

Cell aggregates were incubated in the enzyme solution for 1.5 hourson a gyratory shaker (50 rpm) at 25±1°C in the dark. It is significantthat under the specified conditions, only PEM cells releasedprotoplasts, as determined by monitoring cell aggregates incubated inthe enzyme solution at 10-minute intervals using an invertedmicroscope (Axiovert 10, Zeiss, Germany). Cells within the compactembryonal masses of somatic embryos remained intact due to thepresence of the surrounding protoderm layer, whereas long suspensorcells, on the contrary, collapsed in the presence of hydrolytic enzymes.As the proportion of PEMs to somatic embryos dramaticallydescreased from 3d 2,4-D+BA to 7d ABA (Table 1), the fresh massof the culture sample was correspondingly increased at eachsucceeding time point to yield a sufficient number of protoplasts.

Following enzyme treatment, the mixture of protoplasts, embryonalmasses and cell wall debris was filtered through a series of nylonscreens with successive 400, 200, 150, 80, 50 and 30 µm pore sizes.The fraction of large protoplasts separated on the top of the 30 µmscreen was then resuspended in 10 ml of washing solution containing5 mM CaCl2 and 0.4 M mannitol (pH 5.8) in 15-ml Falcon tubes, andwas centrifuged at 80 g for 5 minutes. The supernatant was removedand the pelleted protoplasts were subsequently washed by the sameprocedure twice more in order to remove nucleases (if any werepresent), recently suggested to be contaminating other types ofOnozuka cellulase, produced by Yakult Honsha Co. Ltd., Tokyo,Japan (Bethke et al., 1999).

The mixture of small protoplasts and cell debris passing through the30 µm screen was centrifuged at 80 g for 5 minutes, the supernatantremoved, and the pellet subjected to the above-described three-step

Table 1. Composition of embryogenic cell line of Norwayspruce at four sequential sampling times

Composition, %

Time point PEM I PEM II PEM III SE

3d 2,4-D+BA 26 36 32 67d 2,4-D+BA 8 26 47 197d no PGR 0 6 19 757d ABA 0 0 1 99

The percentage composition of PEMs (at stages I, II and III) and earlysomatic embryos (SE) were determined after observation of at least 500individual cell aggregates with at least four replicates on every occasion.

d, days.

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washing procedure for removing putative nucleases. The pellet wasthen resuspended in washing solution (9 ml washing solution added to1 ml pellet), and gently mixed with an equal volume of 30% (w/v)Ficoll 400 (Amersham Pharmacia Biotech AB, Uppsala, Sweden) inwashing solution. 2 ml of this mixture was layered over 2 ml of 20%Ficoll in 15-ml Falcon tubes and overlaid with 2 ml of 10% Ficoll (bothin washing solution), and then with 0.5 ml of washing solutioncontaining no Ficoll. The resulting discontinuous Ficoll gradient wascentrifuged at 100 g for 20 minutes. Intact small protoplasts werecollected from the two top interfaces with a Pasteur pipette, pooled andwashed three times as previously described, to remove Ficoll.

The time-lapse tracking performed during the protoplast isolationprocedure has shown that small protoplasts were predominantlyreleased from PEM cells, which stain red with acetocarmine, whilelarge protoplasts originated from Evan’s blue-positive cells of PEMs.

The viability of the protoplasts was assessed using fluoresceindiacetate (FDA, Sigma, St Louis, MO, USA) (Widholm, 1972).Samples of small or large protoplasts were incubated for 5 minutes inFDA at 10 µg/ml and mounted on Polysine slides (Menzel-Gläser,Germany). Over 500 protoplasts with at least three replicates werescored on every occasion with a Microphot-FXA (Nikon, Japan)fluorescence microscope, equipped with a super high pressuremercury lamp supply (HB-10101AF) and the standard set of filtersfor fluorescein (B-2A). Both the living protoplasts showing yellow-green fluorescence and the percentage of protoplasts lacking viabilityand excluding FDA (i.e. FDA-negative) were counted.

Samples were taken from the large and small protoplast fractionsfor electrophoretic DNA fragmentation analyses and TUNEL assay.In a separate series of experiments, protoplasts isolated from PEMsat 3d and 7d 2,4-D+BA and 7d no PGR were not fractionated butdirectly subjected to TUNEL assay.

Electrophoretic DNA fragmentation analysesIn order to resolve large DNA fragments, the whole cell culture orfractionated protoplasts were subjected to pulsed-field gelelectrophoresis (PFGE). Samples of the whole cell culture (Table 1)or protoplasts were mixed with an equal volume of molten 2% (w/v)low-melt preparative grade agarose (Bio-Rad, CA, USA) in 0.5× TBEbuffer (Sambrook et al., 1989) and transferred to a mold at 4°C.Solidified agarose blocks were transferred into a lysis buffercontaining 10 mM Tris-HCl (pH 7.6), 100 mM EDTA, 20 mM NaCl,1% (w/v) Sarcosyl and 5 mg/ml proteinase K (BoehringerMannheim), and incubated at 37°C for 40 hours. Electrophoresis wascarried out in 1% (w/v) pulsed field certified agarose (Bio-Rad, CA,USA)-gel using CHEF-DR II apparatus (Bio-Rad). The gels were runat 5.7 V/cm in 0.5× TBE buffer with pulse ramping time from 10 to60 seconds at 14°C for 24 hours. Agarose gels were stained with 1µg/ml ethidium bromide for 30 minutes and destained with distilledwater for 3 hours. For size determination, PFGE Marker I (BoehringerMannheim) was used as standards.

For assessment of internucleosomal fragmentation, DNA from thewhole cell culture, fractionated protoplasts and apoptotic Jurkat cellswas isolated using the proteinase K-based method (Allen and Newland,1998). In the case of whole cell cultures, the cell walls of intact cellaggregates were broken down by a short (30 seconds) destructivepretreatment in 2 ml FastRNA Tubes-Green (BIO 101, Inc., Vista, CA,USA) using Fast Prep FP120 apparatus (Savant Instruments, Inc.,Farmingdale, NY, USA) prior to DNA isolation. For each sample, 4µg DNA was loaded per lane and electrophoresed in 1.8% MetaPhoreagarose (FMC BioProducts, Rockland, ME, USA)-gel at 6 V/cm using0.5× TBE buffer. The gels were run for 3 hours, stained with 1 µg/mlethidium bromide for 30 minutes and destained with distilled water forfurther 3 hours. For size determination, 100 bp DNA Ladder (MBIFermentas, Vilnius, Lithuania) was used as standards.

In situ detection of DNA fragmentation (TUNEL assay)Norway spruce protoplasts, whole mounts of PEMs and early somatic

embryos, as well as sections of somatic embryos at the beginning andend of late embryogeny, were all subjected to TUNEL assay (Gavrieliet al., 1992), using the fluorescein-dUTP-based in situ death detectionkit (Boehringer Mannheim).

Fractions of small and large protoplasts, as well as non-fractionatedprotoplasts, were fixed by mixing with an equal volume of 4% (w/v)paraformaldehyde in PBS (pH 7.2), dropped on Polysine slides anddried for 1 hour at 40°C. The preparations of whole-mount PEMs(sampled at 3d and 7d 2,4-D+BA) and early somatic embryos (sampledat 7d no PGR and 7d ABA) were fixed in 4% (w/v) paraformaldehydein PBS (pH 7.2) for 1 hour under vacuum at 25±1°C. Somatic embryosat the beginning and end of late embryogeny (sampled after 2 and7 weeks of ABA treatment, respectively) were sequentially fixed,dehydrated and embedded as previously described (Filonova et al.,2000), the only difference being that JB-4TM Embedding Kit(Polyscience, Inc., Warrington, CA, USA) instead of Technovit resinwas used in the present study. The embedded embryos were sectionedon a motorised microtome (HM 350, Microm, Germany), and thesections (3.5 µm) were placed on Polysine slides.

The fixed protoplasts were labeled with TUNEL kit diluted 1:2 inreaction buffer (Boehringer Mannheim) (Danon and Gallois, 1998),excluding permeabilization with proteinase K. The whole mountsand sections were labeled with TUNEL kit according to themanufacturer’s protocol, without dilution (Boehringer Mannheim).Negative controls were included by omitting TdT.

For nuclear staining, the samples labeled with TUNEL kit werewashed twice by PBS and stained with 1 µg/ml 4′-6-diamino-2-phenylindole (DAPI, Boehringer Mannheim) in S-buffer (Kuroiwa etal., 1992). Finally all samples were washed with distilled water,covered by Fluorsave (Calbiochem, CA, USA) and examined with aMicrophot-FXA fluorescence microscope using FITC and UV-2A setsof filters for TUNEL detection and DAPI, respectively. Kodak Elite400 film was used for photomicrography.

The percentage of TUNEL-positive protoplasts was determined byscoring over 500 protoplasts with at least three replicates at everyoccasion. Prior to TUNEL assay, individual PEMs at stages I, II andIII, as well as early somatic embryos, were preselected using aninverted microscope. An assessment of the distribution of TUNEL-positive cells within PEMs and somatic embryos was based on theanalysis of least 20 whole-mount or sectioned PEMs or somaticembryos at a certain developmental stage.

Electron microscopyFor ultrastructural analysis, PEMs III and somatic embryos collectedat 7d 2,4-D+BA and 7d no PGR, respectively, were first embedded inthin layers of 0.6% (w/v) Seaplaque agarose (FMC BioProducts,Rockland, ME, USA) dissolved in distilled water (pH 5.8). Preselectedcell aggregates were cut from agarose layers leaving enough agarosearound to protect cells from injury during fixation and embedding.Fixation and dehydration were carried out according to Fowke (Fowke,1995). Samples were embedded in LR-White Hard Grade resin (TedPella Inc., Redding, CA, USA) using an infiltration protocol previouslydescribed for Technovit resin (Filonova et al., 2000).

Cell aggregates were sectioned using a Reichert Ultracut E/FC4D(Leica, Germany) ultramicrotome and sections were collected onformvar coated copper grids. Sections were observed unstained usinga Philips CM/12 transmission electron microscope operated at 60 kV.Observations were recorded using Kodak 4489 film.

RESULTS

Somatic embryo formation is accompanied by amassive DNA fragmentation in PEM cellsIn embryogenic cell cultures of Norway spruce, withdrawal ofPGRs leads to a profound alteration in the balance between



PEMs and early somatic embryos through stimulating somaticembryo formation and correspondingly suppressing PEMmultiplication (7d 2,4-D+BA cf. 7d no PGR; Table 1). Even apartial depletion of auxin and cytokinin towards the end ofsubculture in the proliferation medium results in an appreciableincrease in the frequency of somatic embryos (3d cf. 7d 2,4-D+BA; Table 1). Treatment with ABA promotes continuousembryo development, with a complete disintegration of the fewremaining PEM III (7d ABA; Table 1).

The integrity of DNA in the whole cell culture sampled atfour sequential time points was assessed by PFGE andconventional agarose gel electrophoresis. Two levels of DNAfragmentation were observed in all samples. First, the presenceof large fragments of about 50 kb and under, known to be amarker for apoptosis (Bortner et al., 1995), was detected byPFGE (Fig. 2A). Secondly, when subjected to conventionalagarose gel electrophoresis, isolated DNA from Norway sprucerevealed an oligonucleosomal ladder, similar to that of humanapoptotic cells (Fig. 2B), indicating fragmentation of DNA intomultiples of approximately 180 bp.

The occurrence of DNA fragmentation in the whole cultureconsisting of a number of cell types aggregated in PEMs andsomatic embryos did not, however, suggest how the frequencyof cells with fragmented DNA changes in the course of somaticembryogenesis and whether this frequency correlates withsomatic embryo formation. The differential responses of PEMand embryo cells to hydrolytic enzymes made it possible toisolate intact protoplasts only from PEMs and not fromembryos (see Materials and Methods) for the quantitativeassessment of DNA fragmentation in PEMs with TUNEL.Isolated protoplasts demonstrated a moderate increase inthe frequency of TUNEL-positive nuclei, from 19% to 24%,upon the transition from 3d to 7d 2,4-D+BA, followed by apronounced increase to 50% after withdrawal of PGRs (7d noPGR; Fig. 3). Interestingly, regression analysis has shown verystrong positive correlation between the frequency of somaticembryo formation and the percentage of PEM-protoplastsfragmenting DNA (r= 0.99, P<0.0001; Fig. 3), suggesting thatPCD in PEMs and somatic embryo formation are closelyinterlinked processes, both stimulated upon withdrawal orpartial depletion of PGRs.

The immediate question arises as to whether there is acertain cell type within PEMs that undergoes cell death andgenerates fragmented DNA. Since PEMs are composedentirely of two types of cells, which are distinguished by sizeand staining patterns with acetocarmine and Evan’s blue (Fig.1), we investigated the fractionation of PEM-protoplasts bysize and subsequently tested fractionated protoplasts for DNAfragmentation. At each of the four sequential time pointsduring somatic embryogenesis, the culture was sampled torelease protoplasts from PEMs followed by the separation ofthe protoplasts into two fractions: large (>30 µm) and small(<30 µm) protoplasts, respectively (see Materials andMethods). Small protoplasts predominantly contained densecytoplasm (Fig. 4A), and were released from acetocarmine-positive cells (Fig. 1), whereas large protoplasts were for themost part highly vacuolated (Fig. 4C) and originated fromEvan’s blue-positive cells (Fig. 1).

Irrespective of time of sampling, the general viability ofprotoplasts as assessed through FDA staining was alwayssignificantly higher for small than for large protoplasts (Fig.

4E,F). Furthermore, large protoplasts always demonstrated aremarkable increase in the frequency of TUNEL-positivenuclei (30-65%; Fig. 4D,F) compared to small protoplasts (1-18%; Fig. 4B,E). Withdrawal of PGRs (i.e. 7d no PGRs) wasthe treatment resulting in the highest percentage of TUNEL-positive nuclei both in small (18±1.4%; Fig. 4E) and largeprotoplasts (65±2.1%; Fig. 4F).

To reveal the molecular pattern of DNA fragmentationaccounting for a high frequency of TUNEL-positive nuclei in afraction of large protoplasts, the protoplasts and isolated DNA

Fig. 2.DNA fragmentation of the whole embryogenic cell culturesampled at the four sequential time points as shown in Table 1.(A) Pulsed-field gel electrophoresis. Formation of large fragments ofabout 50 kb and under was revealed in all samples. (B) Conventionalagarose gel electrophoresis. Internucleosomal DNA fragmentationgenerating a ladder-like pattern on the agarose gel, similar to that ofhuman apoptotic cells (lane JC; see Materials and Methods), wasfound in all samples. Lanes M, marker DNA (see Materials andMethods). The gels shown are representative; all experiments wererepeated at least three times.

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were subjected to PFGE and conventional electrophoresis,respectively. In PFGE, virtually all the DNA from largeprotoplasts migrated at about 50 kb, with relatively moreprominent bands observed for 3d and 7d 2,4-D+BA (Fig. 4H).Furthermore, regardless of sampling time, a typical ladderingpattern suggestive of internucleosomal DNA fragmentation ina fraction of large protoplasts was shown by conventionalelectrophoresis (Fig. 4J). In contrast, neither 50-kb norinternucleosomal fragmentation was detected byelectrophoresis in fractions of small protoplasts (Fig. 4G,I),which displayed a low frequency of TUNEL-positive nuclei(Fig. 4E). A smear-like pattern of DNA degradation shown byPFGE at 3d and 7d 2,4-D+BA (Fig. 4G) and by conventionalelectrophoresis at 7d ABA (Fig. 4I), is probably associated witha substantial contribution of necrotic cell death by a populationof small protoplasts (as judged from increased proportion ofFDA-negative to TUNEL-positive protoplasts; Fig. 4E).

Spatial distribution of nuclear DNA fragmentation inPEMs and developing embryosBy isolation and fractionation of protoplasts, we haveestablished that there is a certain cell type within PEMs ofNorway spruce that is a primary source of fragmented DNA inthe whole population of cultured PEMs. These observationsdid not, however, allow conclusions to be drawn about thedistribution of such cells in an individual PEM developingthrough a stereotyped series of stages (Fig. 1). Likewise, sincethe protoplasts were isolated exclusively from PEMs, there wasno way to determine whether DNA fragmentation is implicatedin embryonic pattern formation, viz. at early and lateembryogeny stages (Fig. 1).

To address these questions, we collected PEMs and somaticembryos at characteristic developmental stages for in situTUNEL assay (see Materials and Methods). No TUNEL-positive cells were found in PEM I-cell aggregates representingthe earliest stage of PEM (Fig. 5A). The first signs of disturbednuclear DNA integrity were observed only after formation ofadditional enlarged highly vacuolated cells, upon attainingPEM II-developmental level, with very few (usually one) ofsuch cells labeled by TUNEL (Fig. 5B). Thereafter, the numberof TUNEL-positive cells gradually increased with continuedgrowth of PEM. In PEM III-cell aggregates lacking polarisedmorphology, nuclear DNA fragmentation was revealed inseveral enlarged highly vacuolated cells, but this was notobserved in any of the small densely cytoplasmic cells (Fig.5C). Finally, formation of somatic embryos from PEMstriggered by withdrawal of PGRs (Fig. 1) was accompanied bya dramatic increase in the frequency of TUNEL labelingthroughout the entire region of degrading PEM III (Fig. 5D).It is remarkable that embryonal masses of newly formedearly somatic embryos did not show any signs of DNAfragmentation, as indicated from the absence of TUNELlabeling (Fig. 5D).

The next wave of DNA fragmentation during somaticembryogenesis of Norway spruce was related to thedegeneration of embryo-suspensors. The latter process occurredduring early embryogeny, after the suspensors had completedelongation, either in PGR-free medium or after addition ofABA. However, only a limited number of suspensor cellsrevealed TUNEL labeling at any one time, suggesting anasynchronous pattern of PCD (Fig. 6A). Following eliminationof the suspensors, during the beginning of late embryogeny, theembryo appeared completely devoid of fragmented DNA (Fig.6B). Fully developed somatic embryos at the end of lateembryogeny (Fig. 6C) contained only a few TUNEL-positivecells localised within the root cap (Fig. 6D), protoderm (Fig.6E), pith (Fig. 6F) and cotyledons (Fig. 6G).

Ultrastructure of cell dismantlingPreselection of individual PEMs III and somatic embryos forultrastructural analysis permitted observations of a largenumber of cells belonging to two cell types that have beenshown to contain TUNEL-positive nuclei (i.e. enlargedvacuolated cells of PEMs and embryo-suspensor cells) and tocompare their ultrastructure with that of meristematic cellssituated in the interior of clumps of small densely cytoplasmiccells in PEMs or embryonal masses of somatic embryos. Sincetransmission electron microscopy of all these cell types hasrevealed no differences between ultrastructural changesunderlying cell dismantling in PEMs and embryo-suspensor,the following information is consistent for both structures.

The cytoplasm of a viable densely cytoplasmic cell containsvery few vacuoles (Fig. 7A), and vacuolation can be defined asan early marker of cell death in our experimental system (Fig.7B). Provacuoles arise from large (diameter 90-130 nm)vesicles budding from the exterior regions of the Golgicisternae (Fig. 7C). The formation of a central autolyticvacuole proceeds not by fusion of provacuoles, but ratherthrough engulfment of large vesicles and portions of cytoplasmby provacuoles (Fig. 7C,D). The formation of plastolysome-like structures from leucoplasts, as earleir described fordegenerating Phaseolussuspensors (Nagl, 1977), was also


60402000

20

40

60

80

Percent TUNEL positive nuclei in PEMs

r = 0.99; P < 0.0001

3d 2

,4D

+B

A 7d 2

,4D

+B

A 7d n

o P

GR

Per

cen

t so

mat

ic

emb

ryo

s

Fig. 3.Correlation of DNA fragmentation in PEMs with somaticembryo formation. Protoplasts were isolated from PEMs at 3d and 7d2,4-D+BA and 7d no PGR. Over 500 non-fractionated protoplastswith three replicates at every occasion were assayed by TUNEL todetermine the percentage of nuclei with fragmented DNA (plottedalong the horizontal axis). Coincident with protoplast isolation,samples of the whole culture were analysed to determine thepercentage of early somatic embryos (plotted along the vertical axis).This included observation of at least 500 individual cell aggregateswith at least four replicates at every occasion.


revealed (Fig. 7E,F). Plastolysome-like structures areprecursors of autolytic vacuoles and consist of a portion ofcytoplasm surrounded by one, or several double membranes;with the sequestered cytoplasm characterised by increasedelectron translucency (Nagl, 1977; Fig. 7E).

In cells of PEMs and embryo-suspensors, provacuolesincrease in number and size towards the cell periphery andprogressively destroy the cytosol and organelles leading to the

formation of large central vacuoles (Fig. 7G,H). At this stage,the cytoplasm often occupies a narrow layer confined betweentonoplast and plasma membrane (Fig. 7H). The tonoplastrupture is delayed until almost complete lysis of the cytoplasmand organelles including the nucleus (Fig. 7I). Once thishappens, the protoplasm disappears leaving a cellular corpserepresented by only the cell wall (Fig. 7J,K).

The earliest morphological sign of nuclear degradation is

Fig. 4.The use of protoplasts forseparation of PEM cells into twofractions, with low and highincidence of DNA fragmentation.Protoplasts were isolated andfractionated by size into twofractions at each of the foursequential time points duringsomatic embryogenesis, as shownin Table 1. The resulting fractionsof small (<30 µm) and large(>30µm) protoplasts were used inDNA fragmentation analyses (seeMaterials and Methods).(A,C) Overview of the fractions ofsmall (A) and large (C)protoplasts, by light microscopy(isolated at 3d 2,4-D+BA).(B,D) Fluorescence microscopy ofthe fractions of small (B) andlarge (D) protoplasts, double-stained by TUNEL (right) andDAPI (left) to detect nuclei withfragmented DNA (greenfluorescence; arrowheads; isolatedat 3d 2,4-D+BA). Bars, 100 µm.(E,F) The average percentage ofFDA-negative and TUNEL-positive protoplasts in thefractions of small (E) and large(F) protoplasts isolated at foursequential time points. Values aremeans ± s.e.m. of at least threereplicates, each containing over500 protoplasts. (G,H) Pulsed-field gel electrophoresis.Formation of large fragments ofabout 50 kb and under wasrevealed in all the samples of largeprotoplasts (H), but in none of thesamples of small protoplasts (G).Lanes M, marker DNA. The gelsshown are representative; allexperiments were repeated at leastthree times. (I,J) Conventionalagarose gel electrophoresis.Internucleosomal DNAfragmentation generated a ladder-like pattern on the agarose gel,similar to that of human apoptoticcells (lane JC in either gel; see Materials and Methods), in all the samples of large protoplasts (J), but in none of the samples of smallprotoplasts (I). Lanes M, marker DNA. The gels shown are representative; all experiments were repeated at least three times.

4406

reflected by the normally round nucleus (Fig. 7A) becomingcrenulated and lobed (Figs 7G, 8A). This is followed bybudding-like segmentation of the lobed nucleus (Fig. 8B). Atthis stage, nuclei display dismantling of the nuclear envelopewithin the areas of segmentation (Fig. 8B) accompanied byclustering of nuclear pore complexes (NPC; Fig. 8C). Theclusters of nuclear pores were found to be closely associatedwith portions of condensed chromatin leaking into cytoplasm(Fig. 8D).

There was no strict interdependency of the timing ofcytoplasm degradation and nuclear segmentation. The lattersometimes occurs at the early stages of cytoplasm degradation,prior to central vacuole formation. In this case, nuclearfragments undergo gradual lysis, as indicated by the presencein provacuoles of remnants of condensed chromatin (Fig. 8D).However, at other times the nuclei remain intact even afterformation of large central vacuole and substantial lysis of thecytoplasm (Fig. 8E). Thus the remnants of chromatin and NPCcan often be seen in the central vacuole at late stages ofcytoplasm degradation (Fig. 8F).

A number of characteristic membrane changes occurduring PCD. Cytoplasmic membranes often form whorlsin dying cells (Fig. 7E). The plasma membrane appearsto remain morphologically intact until rupture of thetonoplast (Fig. 7H). Thereafter it shrinks and lyses,together with remnants of peripheral cytoplasm and someorganelles (Fig. 7I,J).

Most organelles in dying cells are clustered aroundthe nucleus (Figs 7B, 8A). Mitochondria remainmorphologically intact until late stages of cell lysis (Figs7G, 8E). Vesiculation of endoplasmic reticulum (ER)has not been observed. Golgi complexes are activelyparticipating in cell death (see also Henics and Wheatley,1999); they increase in number and produce a collectionof vesicles (Fig. 8D). As mentioned above, large vesicles(diameter 90-130 nm) are involved in provacuoleformation (Fig. 7C). The smaller vesicles (diameter 40-50 nm) are aggregated in clusters that are generallyassociated with ER and randomly distributed in thecytoplasm (not shown).

DISCUSSION

Hallmarks of PCDDuring PCD, the entire pathway for a cell to pass fromperception of the death signal to clearance of the cell inanimal systems or formation of a functional cell wall-corpse in some plant-specific forms of PCD (e.g.,Groover et al., 1997) can be schematically subdividedinto three phases: induction, execution and dismantling(Depraetere and Golstein, 1998). Induction seems to bethe most conservative phase of eukaryotic PCD, whereasthe execution phase and especially the dismantling phasecan vary greatly from organism to organism and evenfrom tissue to tissue in the same organism, therebysuggesting the existence of multiple morphologicallydescribed types of PCD with a few core mechanisms incommon (Depraetere and Golstein, 1998; Olie et al.,1998; Nicotera et al., 1999). Based on the morphologicalhallmarks of the cell dismantling phase, Clarke (Clarke,

1990) proposed three major types of animal PCD: apoptoticcell death (type 1), autophagic or vacuolar cell death (type 2)and non-lysosomal vesiculate cell death akin to autotrophy(type 3). In Norway spruce somatic embryogenesis, a numberof biochemical and cytological hallmarks of the dismantlingphase of cell death in PEMs and embryo-suspensors have beenrevealed. Altogether, these hallmarks do not fit into any of thethree types of animal PCD proposed (Clarke, 1990), but ratherrepresent overlapped pathways of types 1 (i.e. classicalapoptosis) and 2 (i.e. autophagic or vacuolar cell death; see alsoJones, 2000).

The earliest hallmark of PCD in our experimental systemwas progressive vacuolation of the cytoplasm (Fig. 7). It shouldbe emphasised, however, that most of the differentiated plantcells normally have vacuoles playing active roles in creatingand maintenance of turgor, storage of metabolic products andcell elongation (Marty, 1999); that is, the vacuoles in plants arenot exclusive attributes of a dying cell. In this context, it wasnot a state but rather the kinetics of vacuolation observed in


Fig. 5.Fluorescence microscopy of the spatial distribution of nuclear DNAfragmentation in PEMs. Preparations of whole mounts of PEMs wereassayed by TUNEL (upper part of the panels) and counterstained with DAPI(lower part of the panels). Localization of nuclei with fragmented DNA isrepresented by green fluorescence. The figure is a representative result ofobservation of at least 20 whole-mount PEMs of each stage. (A) PEM I-cellaggregate lacking DNA fragmentation; collected at 3d 2,4-D+BA. (B) PEMII-cell aggregate with one TUNEL-positive nucleus (arrowhead) within anenlarged highly vacuolated cell; collected at 3d 2,4-D+BA. (C) PEM III-cellaggregate with several TUNEL-positive nuclei (arrowheads) within enlargedhighly vacuolated cells; collected at 7d 2,4-D+BA. (D) PEM III-cellaggregate forming two somatic embryos (SE); collected at 7d no PGR. Notethe high frequency of TUNEL labeling throughout the entire region ofdegrading PEM III. Bars, 100 µm.


Norway spruce cells that suggested the occurrence ofautophagic processes specific for type 2 cell death. Indeed,autolytic provacuoles in our experimental system wereproduced either by Golgi complexes, as in animal neuronalcells (Hornung et al., 1989; Clarke, 1990) and insuspensor cells of Tropaeolum majus(Gärtner andNagl, 1980), or through a plant-specific way, as aresult of transformation of plastids intoplastolysome-like structures (Nagl, 1977; Gärtnerand Nagl, 1980). It is remarkable, however, that theplasma membrane and many organelles in dyingNorway spruce cells remained morphologicallyintact until very late stages of cell dismantling, acharacteristic feature of type I or apoptotic celldeath (Clarke, 1990; Depraetere and Golstein,1998).

Two levels of DNA fragmentation revealed inthe present work are both consistent features ofapoptosis, where release of chromatin loop-sized 50-kb fragments is usually followed byinternucleosomal fragmentation (Bortner et al.,1995). Late in apoptosis, nuclear pores undergorearrangement into clusters (Reipert et al., 1996),which was also the case in our experimentalsystem, where NPC clustering occurred in theregions of nuclear segmentation. However, we didnot observe other morphological hallmarks ofapoptotic nuclei, such as strong condensation ofchromatin, pyknosis and nuclear membraneblebbing (Clarke, 1990; Willingham, 1999).

The developmental roles of PCD insomatic embryogenesisIn the present study, two major developmentalroles of cell death have been revealed: degradationof PEMs upon embryo formation and eliminationof suspensors during subsequent embryodevelopment.

Formation of PEMs is a well-knowndevelopmental phenomenon that constitutes atransitional state during somatic embryogenesisof different plant species and represents themechanism of maintaining embryogenic potentialunder sufficient amounts of auxin (in angiosperms)or auxin and cytokinin (in spruce) (Halperin, 1966;de Vries et al., 1988; Pennell et al., 1992; McCabeet al., 1997; Filonova et al., 2000). Time-lapsetracking analysis has previously shown that anindividual PEM of Norway spruce cannot developdirectly to an embryo, but rather has two alternativedevelopmental pathways to execute depending onthe conditions: either to multiply in the presence ofauxin and cytokinin, thereby giving rise to newPEMs, or to transdifferentiate to embryos uponwithdrawal of growth regulators (Fig. 1; Filonovaet al., 2000). The latter pathway, as indicated fromthe present study, is accompanied by massive PCDin PEMs (up to 50% TUNEL-positive cells foundat 7d no PGR; Fig. 3). Furthermore, a strongpositive correlation has been shown betweenpercent of TUNEL-positive cells in PEMs and the

frequency of somatic embryos in the whole culture (Fig. 3);i.e. the PEM dies when its transient role to maintain theembryogenic potential is accomplished.

In the developmental context, the transition from the PEM

Fig. 6.Fluorescence microscopy of spatial distribution of nuclear DNAfragmentation in developing somatic embryos. Preparations of whole mounts (A) orsections (B-G) of somatic embryos were assayed by TUNEL (upper part of thepanels) and counterstained with DAPI (lower part of the panels). Localization ofnuclei with fragmented DNA is represented by green fluorescence. The figure is arepresentative result of observation of at least 20 whole-mount or sectioned somaticembryos at each stage. (A) Whole mount of somatic embryo during earlyembryogeny displaying TUNEL labeling (arrowheads) in several cells withinsuspensor (s); em, embryonal mass; collected at 7d no PGR. (B) Longitudinalsection of somatic embryo in the beginning of late embryogeny showing a lack ofDNA fragmentation; collected after 2 weeks of ABA-treatment. (C) Longitudinalsection of somatic embryo at the end of late embryogeny (DAPI staining only)showing the regions where DNA fragmentation could sometimes occur (arrowspointing to D-G); collected after 7 weeks of ABA-treatment. (D) root cap (RC);(E) protoderm (PD); (F) pith (PH); (G) cotyledons (C). Bars, 100 µm.

4408 L. H. Filonova and others

Fig. 7.The ultrastructure of cytoplasmic changes revealed during cell dismantling. For transmission electron microscopy analysis, PEMs IIIand somatic embryos were collected at 7d 2,4-D+BA and 7d no PGR, respectively (see Materials and Methods). The micrographs illustrate thecommon changes characteristic for cells of both PEMs and embryo-suspensors. (A) A normal densely cytoplasmic cell with a round nucleus(n). cw, cell wall. (B) Formation of numerous vacuoles around the nucleus (n) as an early marker of PCD. (C) Large vesicles (diameter 90-130nm; arrowheads) budding from the Golgi (G) cisternae and participating in provacuole (pv) formation. Note the inclusions of such vesicles(arrows) in provacuoles. (D) Engulfment of large vesicles (arrows) and portions of the cytoplasm (asterisk) by provacuoles (pv).(E) Plastolysome-like structures (pl) containing more electron-translucent cytoplasm. Inset shows a higher magnification of plastolysome-likestructure surrounded by a double membrane (arrowhead). Asterisk denotes a whorl formed by the cytoplasmic membrane. (F) Thetransformation of leucoplasts (lc) into plastolysome-like structures. (G) Cell at moderately advanced stage of PCD. The fragment of thecytoplasm with lobed nucleus (n) is held against the cell wall (cw) by a central vacuole (cv). Note that the mitochondria (arrows) remainmorphologically intact. (H) Almost complete lysis of the intracellular components including nucleus by expanding central vacuole (cv). Thecytoplasm is reduced to a narrow layer confined between tonoplast (tp) and plasma membrane (pm). (I) Cell at a late stage of PCD, after therupture of tonoplast. Plasma membrane (pm) is shrunken and detached from the cell wall (cw). Note the presence of a few organelles remainingattached to the plasma membrane. (J) Complete disappearance of organelles and lysis of fragmented plasma membrane (pm). (K) Cellularcorpse represented by cell wall (cw) only. Bars, 2.5 µm (A,B,E,G-K); 0.5 µm (C,D,F, inset in E).


phase to the somatic embryo phase, its rigid control by the levelof PGRs, and the concomitant destruction of structures setapart in the previous phase, as found in our experimentalsystem, are all essential criteria of the well-knownphenomenon of metamorphosis; a stereotyped sequence ofevents during the transition from larvae to adult forms inamphibian and insect development (Müller,1997). Similar to the PEM-to-embryotransition in Norway spruce, classicmetamorphosis involves extensive hormone-dependent remodelling of tissues and organs(Jiang et al., 1997; Weeks, 1999), wherePCD plays an important constructive role(Schubiger et al., 1998; Su et al., 1999). Wecould find only one process in plantdevelopment showing similarities bothwith PEM-to-embryo transition in Norwayspruce and with animal metamorphosis.That is a unique type of embryogenydescribed in Paeonia spp. where viablemultiple embryos form from a massivestructure (called the proembryo), whichsubsequently becomes highly vacuolatedand ultimately collapses (Yakovlev andYoffe, 1957; Cave et al., 1961). It wassuggested that embryogeny in gymnospermsand Paeonia represents a case of parallelevolution (Cave et al., 1961).

Another site of PCD found in the presentwork was the embryo-suspensor. We haverevealed an asynchronous pattern of PCD inNorway spruce suspensor, indicated by thepresence of just a few TUNEL-positive cellsin the samples of early somatic embryos(Fig. 6A). The embryo-suspensor isconsidered to be the first terminallydifferentiated structure produced during plantdevelopment. In normal embryogenesisprograms, the suspensor accomplishes itsfunction and ultimately dies at a certainstage of embryo-proper development, i.e. atthe end of the heart stage in angiosperms(Yeung and Meinke, 1993) and at the end ofearly embryogeny in gymnosperms (Singh,1978). The proper timing of suspensordegradation appears to be an importantprerequisite for correct embryonic patternformation. In Arabidopsis, two types ofembryo-specific mutants developed as aresult of misregulated PCD in the suspensorand thus disturbed interactions betweenembryo proper and the suspensor have beencharacterised. One mutant (twin) displays anabnormal proliferation of suspensors givingrise to multiple embryos (Vernon andMeinke, 1994; Zhang and Somerville,1997), while another mutant (raspberry)fails to undergo the transition from globularto heart stage and continues to proliferateboth in suspensor and embryo-properregions (Yadegari et al., 1994). We have

previously shown that intensive radial growth, delay in theonset of histogenesis and finally disintegration of embryonictissues in B-type somatic embryos of Norway spruce wascorrelated with prolonged longevity of suspensors (Filonova etal., 2000).

There were a few dying (TUNEL-positive) cells found to be

Fig. 8.The ultrastructure of nuclear changes revealed during cell dismantling. Fortransmission electron microscopy, PEMs III and somatic embryos were collected at 7d2,4-D+BA and 7d no PGR, respectively (see Materials and Methods). The micrographsillustrate the common changes characteristic for cells of both PEMs and embryo-suspensors. (A) Lobing of nucleus (n) at moderately advanced stage of PCD. cw, cell wall.(B) Budding-like nuclear segmentation resulted in a separation of nuclear fragment(asterisk; inset shows a higher magnification) into adjacent cytoplasm. Notable is thedismantling of nuclear envelope (arrows in inset). (C) Clustering of nuclear pore complex(NPC) during nuclear segmentation. (D) Nuclear degradation detected at early stages ofcytoplasm lysis, before formation of central vacuole. Note the inclusions of condensedchromatin (arrowheads) within provacuoles (pv), and the clustering of nuclear pores (box)associated with condensed chromatin leaking into cytoplasm. Inset shows a highermagnification of the Golgi complex (G; asterisk in the main panel) being converted into acollection of vesicles. (E) Formation of central vacuole (cv) that includes an intact nucleus(n) surrounded by a sheath of cytoplasm. Arrows denote morphologically intactmitochondria. (F) The remnants of condensed chromatin and NPC (arrows in inset) incentral vacuole (cv) at late stage of PCD. Bars, 2.5 µm (insets, 0.5 µm).

4410

set apart as a result of differentiation of embryonal mass tissuesat final stages of somatic embryo development (Fig. 6C-G).These incidents of PCD are likely to be associated with theformation of provascular tissue in the pith region (Carland etal., 1999), transformation of protoderm to epidermis (Lackieand Yeung, 1995), the onset of cotyledon senescence (Smart,1994) and sloughing of the root cap cells (Wang et al., 1996).

In conclusion, somatic embryogenesis in Norway spruceinvolves two successive waves of PCD, which are responsiblefor degradation of PEMs and elimination of embryo-suspensors. These cell suicide events ensure normalprogression of somatic embryogenesis, i.e. transition fromPEMs to somatic embryos and correct embryonic patternformation, respectively. Further studies will be aimed atelucidation of the molecular mechanisms of developmentalcell death in our experimental system through targetedstimulation and inhibition of key biochemical pathways thatare known to be recruited during the classical caspase-dependent apoptosis (Wertz and Hanley, 1996) and therecently characterised autolytic type of PCD leading to planttracheary element formation (Woffender et al., 1998; Grooverand Jones, 1999).

We thank Dr Alan Jones (North Carolina University, Chapel Hill)for stimulating discussions and criticism and Dr Lisa Butterworth(Uppsala Genetic Centre) for advice with pulsed-field gelelectrophoresis. This work was supported by Thematic researchproject (joint The Swedish University of Agricultural Sciences/TheForestry Research Institute of Sweden), Jakob WallenbergsFoundation and the Swedish Institute.

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Programmed cell death in plant embryogenesis - Journal of Cell Science