The Plant Cell, Vol. 12, 933–947, June 2000, www.plantcell.org © 2000 American Society of Plant Physiologists

Syncytial-Type Cell Plates: A Novel Kind of Cell Plate Involved in Endosperm Cellularization of Arabidopsis

Marisa Otegui

1

and L. Andrew Staehelin

Department of Molecular, Cellular, and Developmental Biology, University of Colorado, Boulder, Colorado 80309-0347

Cell wall formation in the syncytial endosperm of Arabidopsis was studied by using high-pressure-frozen/freeze-sub-stituted developing seeds and immunocytochemical techniques. The endosperm cellularization process begins at thelate globular embryo stage with the synchronous organization of small clusters of oppositely oriented microtubules

(

z

10 microtubules in each set) into phragmoplast-like structures termed mini-phragmoplasts between both sister andnonsister nuclei. These mini-phragmoplasts produce a novel kind of cell plate, the syncytial-type cell plate, from Golgi-derived vesicles

z

63 nm in diameter, which fuse by way of hourglass-shaped intermediates into wide (

z

45 nm in diam-eter) tubules. These wide tubules quickly become coated and surrounded by a ribosome-excluding matrix; as theygrow, they branch and fuse with each other to form wide tubular networks. The mini-phragmoplasts formed between agiven pair of nuclei produce aligned tubular networks that grow centrifugally until they merge into a coherent wide tu-bular network with the mini-phragmoplasts positioned along the network margins. The individual wide tubular net-works expand laterally until they meet and eventually fuse with each other at the sites of the future cell corners.Transformation of the wide tubular networks into noncoated, thin (

z

27 nm in diameter) tubular networks begins at mul-tiple sites and coincides with the appearance of clathrin-coated budding structures. After fusion with the syncytial cellwall, the thin tubular networks are converted into fenestrated sheets and cell walls. Immunolabeling experiments showthat the cell plates and cell walls of the endosperm differ from those of the embryo and maternal tissue in two features:their xyloglucans lack terminal fucose residues on the side chain, and callose persists in the cell walls after the cellplates fuse with the parental plasma membrane. The lack of terminal fucose residues on xyloglucans suggests thatthese cell wall matrix molecules serve both structural and storage functions.

INTRODUCTION

In higher plants, new cell walls are formed during cytokine-sis by fusion of Golgi-derived vesicles into cell plates. Thestructure that gives rise to the cell plate is the phragmoplast,a complex arrangement of microtubules, microfilaments,Golgi-derived vesicles, and endoplasmic reticulum that as-sembles during late anaphase and is dismantled once thenew wall is complete (Staehelin and Hepler, 1996; Smith,1999). The phragmoplast cytoskeleton consists of two op-positely oriented sets of microtubules with their plus ends inthe plane of the cell plate and two corresponding sets of ac-tin microfilaments that do not overlap or directly abut. Thefirst phragmoplast microtubules appear to arise from rem-nants of the mitotic spindle, but all later ones are poly-merized anew, forming a cylinder that consolidates byshortening in length and widening in girth. During cell plateexpansion, the microtubules depolymerize in the center andrepolymerize along the edge, transforming the phragmo-

plast into a barrel-like structure, which marks the growingmargin of the cell plate (Staehelin and Hepler, 1996; Heeseet al., 1998). The high density of microtubules in the phrag-moplast establishes an organelle exclusion zone along thegrowing edges of the cell plate (Samuels et al., 1995).

In somatic plant cells, the development of a preprophaseband signals the future site of fusion of the cell plate with theparental cell wall. A detailed model depicting the structuralevents associated with somatic cell plate formation in high-pressure-frozen/freeze-substituted tobacco root tips andBY-2 cells has been published by Samuels et al. (1995). Twocentral features of this model are the involvement of thin(20-nm-diameter) fusion tubes in the formation of a transientmembrane network from Golgi-derived vesicles and thesubsequent programmed series of maturation steps thatconvert the membrane network into a new cell wall.

In contrast, in syncytial systems such as the nuclear en-dosperm, meiocytes, and gametophytic cells, cytokinesis isuncoupled from mitosis, and no preprophase bands havebeen reported (Brown and Lemon, 1991; Mineyuki, 1999).During endosperm development, the primary nucleus ini-tially undergoes a series of divisions without cytokinesis,

1

To whom correspondence should be addressed. E-mail oteguim@stripe.colorado.edu; fax 303-492-7744.

934 The Plant Cell

forming a syncytium. Shortly thereafter, the cellularizationprocess is initiated by the formation of cell walls betweensister and nonsister nuclei, which are perpendicular (i.e., an-ticlinal) to the syncytial cell wall. Although many studieshave dealt with endosperm cellularization, the formation ofthe very first anticlinal walls between nonsister nuclei has re-mained controversial for more than 90 years (DeMason,1997).

The most widely accepted theory of endosperm cellular-ization, developed from studies on cereals, postulates thatthe first anticlinal cell walls are deposited in the absence ofmitosis and typical phragmoplasts (reviewed in Olsen et al.,1995; Heese et al., 1998; Berger, 1999). In this model, “free-growing” cell walls are postulated to arise from the syncytialcell wall like ingrowths in the absence of typical phragmo-plast arrays and to then extend toward the central vacuole.This mechanism has been compared with the ingrowths thatinitiate cellularization of the blastoderm embryo in Drosoph-ila (Heese et al., 1998). The sites at which these inward-growing cell walls form coincide with the interacting ends ofopposing microtubule arrays that radiate from adjacent nu-clei (Brown et al., 1996a; Olsen, 1998; Olsen et al., 1998).Furthermore, some have speculated that the points of initia-tion of the free-growing anticlinal walls are related to transferingrowths (Dute and Peterson, 1992; Chamberlin et al.,1994). Besides operating in cereals, this mechanism of cellwall formation has also been reported for the endosperm ofArabidopsis

(Mansfield and Briarty, 1990; Brown et al.,1999),

Euphorbia dulcis

(Gori, 1987),

Glycine max

(Dute andPeterson, 1992; Chamberlin et al., 1994),

Haemanthuskatherinae

(Newcomb, 1978),

Helianthus annus

(Newcomb,1973),

Papaver nudicaule

(Olson, 1981),

Phaseolus vulgaris

(Yeung and Cavey, 1988), and

Stellaria media

(Newcomband Fowke, 1973).

However, the findings of other studies seem to contradictthis prevailing theory of endosperm cellularization. For ex-ample, cytokinesis involving conventional phragmoplastsand cell plate formation between two dividing nuclei havebeen described in the endosperm of wheat (Fineran et al.,1982) and

Ranunculus sceleratus

(XuHan and Van Lammeren,1993). In addition, non-mitosis-related phragmoplasts andcell plates have been observed during endosperm cellular-ization of wheat (Van Lammeren, 1988) and

P

.

vulgaris

(XuHanand Van Lammeren, 1994).

In part, these contradictory reports, some even derivedfrom observations on the same species, may be related tothe difficulty of preserving endosperm cells by using chemi-cal fixatives. To circumvent this problem, we have usedhigh-pressure freezing/freeze-substitution techniques topreserve cellularizing endosperm cells of Arabidopsis. Sam-ples so preserved demonstrate that the initial cellularizationprocess in the syncytial endosperm includes the formationof novel and unique kinds of phragmoplasts and cell plates.We have named these structures mini-phragmoplasts andsyncytial-type cell plates, respectively, to distinguish themfrom the more common somatic-type phragmoplasts and

somatic-type cell plates. Mini-phragmoplasts contain veryfew microtubules, and multiple mini-phragmoplasts work inconcert to produce a syncytial-type cell plate. Although thisprocess is documented only for cell plate formation in themicropylar zone, the same structural intermediates havebeen observed during the initial stages of cell wall formationin the central and chalazal endosperm zones. The subse-quent cell wall growth steps in these latter regions involveother intermediates, which will be reported elsewhere.

RESULTS

The Developing Endosperm

At 70 to 80 hr after anthesis, the syncytial endosperm con-sists of a layer of cytoplasm with free nuclei surrounding thelarge central vacuole (Figure 1A). At this stage, the free nu-clear division cycles have been completed, and no mitoticspindle remains can be detected. The endosperm is sur-rounded by the endothelial cells, which typically exhibit adarkly staining cytoplasm (Figure 1A). Three distinct regionscan be recognized in the highly asymmetric developing en-dosperm of Arabidopsis: the micropylar zone, the centralzone, and the chalazal zone. The cytoplasm-rich micropylarzone completely envelops the developing embryo (Figures1A and 1B). The adjacent central zone consists of a thin pe-ripheral layer of cytoplasm with regularly spaced nuclei anda large central vacuole, whereas the chalazal zone displaysa set of dispersed nuclei within a dense cytoplasm (Figure1A). The latter zone is surrounded in part by the chalazalproliferating cells (Figure 1A).

The micropylar zone can be divided into two different do-mains according to their relation to the central vacuole. Asshown in Figure 1B, one cytoplasmic domain is sandwichedbetween the endothelial cells and the suspensor and part ofthe embryo proper but does not abut the central vacuole,whereas the other domain is bordered by the remaining partof the embryo proper and the central vacuole. Here, we doc-ument the formation of anticlinal cell walls that, when com-pleted, form direct bridging structures between the syncytialcell walls adjacent to the endothelium and the embryo. Wehave also observed the same types of mini-phragmoplastand syncytial-type cell plate during the initial stages of cellu-larization of the central and chalazal zones. However, be-cause the subsequent cell wall growth steps in these latterzones involve numerous other structural intermediates, a fullreport of the cellularization process in the central and cha-lazal zones will be presented elsewhere.

Near the suspensor, the syncytial cell wall facing the en-dothelium exhibits conspicuous transfer ingrowths sur-rounded by numerous mitochondria. The cytoplasm in themicropylar zone typically contains dilated endoplasmicreticulum cisternae, Golgi stacks, and plastids with grana(data not shown).

Syncytial-Type Cell Plate in Arabidopsis Endosperm 935

Structures Associated with Cell Plate Formation in the Micropylar Zone

The cellularization process in the micropylar zone of the en-dosperm begins at the late globular embryo stage with thesynchronous assembly of small groups of oppositely ori-ented microtubules into phragmoplast-like structures andthe formation of cell plates between both sister and nonsis-ter nuclei (Figure 2). Because these “phragmoplasts” andcell plates differ in important respects from those producedby somatic cells, we refer to the structures reported here asmini-phragmoplasts and syncytial-type cell plates. Eachmini-phragmoplast consists of two opposing sets of

z

10microtubules each, and the cell plates are formed by multi-ple mini-phragmoplasts that act in concert between pairs ofadjacent endosperm nuclei. These mini-phragmoplasts ap-pear to deliver Golgi-derived vesicles

z

63 nm in diameter(

SD

6

10,

n

5

20) to the equatorial plane between the nuclei,where their fusion initiates the process of cell plate forma-tion (Figures 3A to 3F). Fusion of these vesicles involveshourglass-shaped intermediates (Figures 3B to 3D). Fusiontubes, the unique 20-nm-wide membrane tubes that medi-ate vesicle fusion in somatic cell plates (Samuels et al.,1995), were never observed.

Fusion of the cell plate–forming vesicles results in the for-mation of wide membrane tubules with a diameter of 45 nm(

SD

6

11,

n

5

15) (Figures 3E and 4), which quickly becomecovered by a fuzzy coat of fine filamentous molecules (Fig-ures 5A and 5B). As this filamentous mesh expands, it ex-cludes ribosomes from the region at which the cell platesare forming. Often, the first cell plate–forming wide tubulesappear aligned with the syncytial phragmoplast microtu-bules (Figure 3E). However, as they fuse with each other toform a branched wide tubule network, they assume a moreequatorial orientation (Figures 3F, 4, 5A, and 5B).

The next stage of syncytial cell plate maturation involvesthe conversion of the wide tubular network into a thin tubu-lar network and the disassembly of the fuzzy coat (Figure

Figure 1.

Light Microscopy of a Longitudinal Section of a Develop-ing Arabidopsis Seed.

(A)

Overview. Three distinct regions can be recognized in the devel-oping endosperm: the micropylar zone (MZ) in which the embryo (E)is located, the central zone (CZ), and the chalazal zone (CHZ). Thecentral zone consists of a thin peripheral layer of cytoplasm withregularly spaced nuclei (asterisks) and a large central vacuole (VA).The chalazal zone is partially enveloped by the chalazal proliferatingcells (CHPC). The endosperm is surrounded by the endothelium(EN). Bar

5

50

m

m.

(B)

Detail of the micropylar zone. Two different domains can be dis-tinguished according to their relation with the central vacuole. Onecytoplasmic domain (I) surrounds part of the embryo proper (E) andthe suspensor (S) but does not abut the central vacuole (VA). Theother domain (II) is bordered by the remaining part of the embryoproper and the central vacuole. Endosperm nuclei are indicated byasterisks. EN, endothelium. Bar

5

10

m

m.

936 The Plant Cell

5C). During this process, the diameter of the tubules is re-duced from

z

45 nm to

z

27 nm (

SD

6

6,

n

5

15). Coincidentwith the formation of noncoated thin tubules is the appear-ance of clathrin-coated buds and vesicles, mainly located atthe ends of the thin tubules (Figure 6B).

Although each mini-phragmoplast appears to act inde-pendently, those that form between a given pair of nucleiproduce aligned cell plates that develop synchronously (Fig-ure 4). The wide tubular networks formed by each mini-phragmoplast (Figure 4) grow centrifugally until they mergeinto a coherent cell plate network (Figures 6A and 6B). Be-cause of the variable age of these different network do-mains, cell plate maturation occurs in a patchwork manner,as evidenced by the multiple sites involved in transition fromwide tubules to thin tubules (Figure 6B, open arrows).

One of the most intriguing and defining features of thesyncytial-type cell plate formation is the scarcity of microtu-bules involved in this process. These cell plates arise at sitesdefined by small sets of microtubules organized into mini-phragmoplasts (Figure 3). At later stages, the clusters ofmini-phragmoplast microtubules (4 to 12 microtubules/mini-

phragmoplast) are seen around the margins of the tubularcell plate networks (Figure 6B, dotted circles). In addition, afew single microtubules are seen between the network tu-bules (Figure 6B, arrows). The high frequency with whichvesicles are observed adjacent to both clustered and indi-vidual microtubules (Figure 6A) suggests that their principalfunction is to deliver Golgi-derived vesicles to the formingcell plate.

Lateral growth of the syncytial-type cell plates appears tofollow the same sequence of events described above for theinitiation of the individual tubular networks (Figures 3B to3D). First is the fusion of vesicles (Figure 6A) and the forma-tion of wide tubules by way of hourglass intermediates; thencomes the fusion of these tubules with elements of the widetubular network at the growing edge of the cell plate. The in-corporation of new vesicles to the cell plate by means ofvesicle and tubule fusion processes occurs mostly near thegroups of mini-phragmoplast-associated microtubules.

A consequence of the lack of a broad band of closelypacked phragmoplast microtubules around the edge of syn-cytial-type cell plates is the absence of a structure for push-ing larger organelles out of the way of the growing cell plateedge. Thus, it is not uncommon to see the narrow edge ofan expanding syncytial-type cell plate colliding with a largeorganelle such as a plastid or a mitochondrion (data notshown). The small amount of buckling of the cell plate edgeseen in instances where organelle denting is evident sug-gests that the cell plate margin is quite stiff.

The cytoplasm of the micropylar zone of the endospermcontains a single layer of free nuclei (Figure 7, asterisks). Be-cause this zone forms a cylindrical sheath around the sus-pensor, the nuclei in this sheath are organized in the form ofa two-dimensional array parallel to the syncytial cell wall.Thus, the cellularization of this region of cytoplasm requiresthe formation of multiple cell plates around each nucleus toform a honeycomb-like organization of cell walls perpendic-ular to the syncytial cell walls (anticlinal cell walls). This pro-cess is illustrated in Figure 8, which depicts a tangentialview through the endosperm cytoplasm in the micropylarzone. In particular, Figure 8A shows multiple cell platesaround a single nucleus; moreover, these cell plates are inthe process of linking with each other and with the other ad-jacent cell plates, forming Y- or T-shaped junctions. Athigher magnifications (Figure 8B), this joining is seen to in-volve cell plates that are still in the tubular network stage ofdevelopment. Maturation appears to be slow compared withnetwork growth, as evidenced by the fact that the large cellplates retain mostly a tubular network architecture acrosstheir entire width, even as they fuse with each other (Figure8). Once the adjoining cell plate networks have fused to-gether, they link with the plasma membrane and the syncy-tial cell walls, while simultaneously maturing into fenestratedsheets (Figure 9). Clathrin-coated buds/vesicles are frequentduring this stage (Figures 9A and 9B), and multivesicularbodies are common in the vicinity of the cell plate (Figure9B). The fenestrated sheets finally mature into anticlinal cell

Figure 2. Syncytial-Type Cell Plates Form between Nonsister Nu-clei at the Onset of Cellularization in the Micropylar Zone.

Cell plates are aligned fairly perpendicular to the syncytial cell wallsfacing the embryo (E-SCW) and the endothelium (EN-SCW) (anticli-nal orientation). Arrows indicate syncytial-type cell plates. N, nuclei;S, suspensor; VA, vacuole. Bar 5 2 mm.

Syncytial-Type Cell Plate in Arabidopsis Endosperm 937

walls, which contain numerous primary plasmodesmata (datanot shown).

Composition of Syncytial-Type Cell Plates

To determine whether syncytial-type cell plates differ incomposition from somatic-type cell plates, we have labeled

cryofixed/freeze-substituted and plastic-embedded speci-mens with a series of anti-polysaccharide antibodies and acellulose binding probe (Figures 10 and 11). To assess thexyloglucan content of these cell plates, we used both apolyclonal anti-xyloglucan antibody, which recognizes the

b

-1,4–linked glucose backbone of xyloglucan (Lynch andStaehelin, 1992), and the monoclonal antibody CCRC-M1,which binds to the terminal fucosyl residue of the trimeric

Figure 3. Examples of Assembly of Mini-Phragmoplasts at the Onset of Endosperm Cellularization and Vesicle Fusion Steps.

(A) Mini-phragmoplast (arrowhead) consisting of two sets of opposed microtubules (arrows). Golgi-derived vesicles of z63 nm (V) are associ-ated with the mini-phragmoplast microtubules. VA, vacuole.(B) to (D) Vesicle fusion steps showing two Golgi-derived vesicles together (B), an hourglass-shaped intermediate resulting from the fusion oftwo vesicles (C), and a transient structure between an hourglass-shaped intermediate and a wide tubule (D).(E) Wide tubules (WT) are aligned in a parallel orientation to the microtubules (arrows) in a mini-phragmoplast (arrowhead).(F) Fused and branched wide tubules in a mini-phragmoplast (arrowhead).Bars in (A), (E), and (F) 5 200 nm; bars in (B) to (D) 5 50 nm.

938 The Plant Cell

side chain of xyloglucan in the context of an extended xylo-glucan conformation (Puhlman et al., 1991). The anti-xylo-glucan antibody labeled the cell plates strongly, starting withthe earliest stages of their assembly (Figure 10A). Labelingwas also seen over Golgi stacks and vesicles. In contrast,the CCRC-M1 antibody failed to label any structure of thesyncytial endosperm, including cell plates, Golgi stacks, oreven cell walls (Figure 10B). However, CCRC-M1 labelingwas observed over the cell walls of both the embryo and theintegument and endothelial cells surrounding the en-dosperm (Figure 10B). Thus, the lack of CCRC-M1 labelingof the syncytial-type cell plates and the syncytial cell wallsmost likely reflects an absence of the xyloglucan–fucoseepitope in these structures rather than an inability of the an-tibody to bind effectively to the epitopes in our sections.

The presence of methyl-esterified pectins was tested byusing the JIM7 monoclonal antibody (Knox et al., 1990). Thisantibody failed to bind esterified pectins on Epon-embedded(Ted Pella, Inc., Redding, CA) samples, but labeling was ob-tained on tissues embedded in London Resin White (Elec-tron Microscopy Sciences, Fort Washington, PA). Althoughthe preservation of syncytial-type cell plate with this latterresin was poor (data not shown), we were able to identifynewly formed anticlinal cell walls in which JIM7 labeling wasobserved (Figure 10C).

Callose, which was first detected by means of anti-callose(anti–

b

-1,3-glucan) antibody (Meikle et al., 1991) during thewide tubule stage of syncytial-type cell plate formation, in-creases during cell plate maturation (Figure 11A). Strong la-beling was also seen over recently completed cell walls inwhich all fenestrae had closed. On the other hand, no cal-lose labeling was observed over Golgi stacks, vesicles, syn-cytial cell walls, or transfer ingrowths (data not shown).

The presence of low concentrations of cellulose in syncy-tial-type cell plates was suggested by the binding of the cel-lulose probe cellobiohydrolase I–gold (CBHI-gold) (Figure11B). This labeling, however, was much less than that overthe mature cell walls of the surrounding cells (data notshown). Cellulose could first be detected during the wide tu-bular network stage (Figure 11B) and increased through thefenestrated sheet stage. Because no labeling of Golgi

Figure 4. Two Wide Tubular Networks Formed in Two AdjacentMini-Phragmoplasts.

WTN, wide tubular network. Bar 5 200 nm.

stacks or of vesicles was observed, this labeling is highlylikely to have resulted from binding to cellulose and not frombinding of the probe to the backbone of xyloglucan mole-cules, which also consists of

b

-1,4–linked glucan residues.

DISCUSSION

The formation of the very first anticlinal walls between non-sister nuclei in the syncytial endosperm has remained contro-versial for more than 90 years (DeMason, 1997). Thus,despite general agreement about the role of nuclear-basedradial microtubule systems for the positioning of anticlinalcell walls, the mechanism of assembly of these cell wallshas remained an enigma. Briefly, three different theorieshave been put forward to explain this process: (1) extensionof septal-like ingrowths of the syncytial walls in the absenceof mitosis and typical phragmoplasts; (2) conventionalphragmoplast-mediated cell plate formation after mitosis,and (3) non-mitosis-related phragmoplasts and cell plates.For Arabidopsis, two previous studies have reported thatthe first anticlinal cell walls in the endosperm arise from sep-tal-like extensions of the syncytial cell wall (Mansfield andBriarty, 1990; Brown et al., 1999). In the present study,which is based on samples preserved by high-pressurefreezing/freeze substitution rather than chemical fixation, wedemonstrate that the initial anticlinal walls formed during en-dosperm cellularization do not arise from septal-like in-growths but are produced instead by groups of mini-phragmoplasts, which give rise to unique cell plates, syncy-tial-type cell plates, that then mature into cell walls. Themain stages of this process are summarized diagramaticallyin Figures 12 and 13.

Small Clusters of Microtubules Become Organized into Mini-Phragmoplasts during Syncytial-Type CellPlate Development

Cellularization of the endosperm in Arabidopsis starts withthe formation of several mini-phragmoplasts between bothsister and nonsister nuclei (Figures 3A, 3E, and 3F). Each ofthese mini-phragmoplasts comprises two opposing sets of

z

10 microtubules, which during later stages of cell plate de-velopment are seen to persist as distinct morphologicalunits around the cell plate margins (Figure 6B). This particu-lar type of phragmoplast organization has never been ob-served during plant cytokinesis in somatic cells (Baskin andCande, 1990; Heese et al., 1998).

The scarcity of microtu-bules associated with the mini-phragmoplasts during forma-tion of syncytial-type cell plates probably reflects the factthat during the cellularization process, not enough tubulin isavailable to assemble simultaneously five or six conven-tional somatic cell-type phragmoplasts around each en-dosperm nucleus.

Syncytial-Type Cell Plate in Arabidopsis Endosperm 939

In somatic cells, the phragmoplast microtubules arise ini-tially from remnants of the mitotic spindle. In developing Ar-abidopsis seeds, however, Brown et al. (1999) haveobserved that after completion of the cycles of free nucleardivisions, no mitotic spindle microtubules remain in the syn-cytial endosperm before assembly of the new cell walls. In-stead, the syncytial endosperm becomes organized intonuclear cytoplasmic domains (Brown and Lemmon, 1992;Pickett-Heaps et al., 1999) that are defined by radial micro-tubule arrays produced by microtubule organizing centersassociated with the external surface of the nuclear envelope(Lambert, 1995). This nuclear-based arrangement of micro-tubules has also been noticed in the syncytial endosperm ofother species, such as rice, barley, wheat, and

P

.

vulgaris

(Van Lammeren, 1988; Brown et al., 1994; XuHan and VanLammeren, 1994; Olsen et al., 1995; Brown et al., 1996a,1996b) as well as in other types of syncytial systems such asdeveloping microspores (Van Lammeren et al., 1985; Brownand Lemmon, 1991). The interaction of opposing microtu-bules emanating from adjacent nuclei has been suggestedto play an important function in determining the sites of non-mitosis-related phragmoplast formation between nonsisternuclei (Van Lammeren, 1988; XuHan and Van Lammeren,1994). Given the structural data presented here, we postu-late that at the onset of cellularization in Arabidopsis en-dosperm, the mini-phragmoplast microtubules arise fromthe opposing overlapping clusters of microtubules radiatingfrom neighbor nuclei (Figure 12). Because the same nuclearcytoplasmic domain organization has been observed in themicropylar, central, and chalazal zones of Arabidopsis en-dosperm (Brown et al., 1999), we think it likely that mini-phragmoplasts originate in the same manner in all of the en-dosperm zones.

Growth of Syncytial-Type Cell Plates Involves Fusion of Vesicles by Way of Hourglass Intermediates, Formation of Wide Tubules, and Fusion of Wide Tubulesinto Networks

One of the most striking differences between syncytial-and somatic-type cell plate formation pertains to themechanism of fusion of the cell plate–forming vesicles. Individing somatic cells, this process involves unique, 20-nm-diameter fusion tubes (Samuels et al., 1995; Verma andGu, 1996), whereas in the syncytial endosperm, fusionseems to occur only between closely opposed vesicles, asevidenced by the hourglass-shaped fusion intermediates(Figures 3B to 3D).

At present, very little is known about the molecules in-volved in the fusion of cell plate–forming vesicles. TheKNOLLE protein, a cytokinesis-specific syntaxin, plays acritical role in membrane fusion events associated with so-matic-type cell plate formation, most likely by participatingin the homotypic fusion of Golgi vesicles in somatic-type cell

Figure 5. Developmental Stages in Syncytial-Type Cell Plate For-mation.

(A) Early stages in vesicle aggregation and wide tubule consolida-tion. Wide tubules are already covered by a fuzzy coat (FC).(B) Wide tubular network covered by a dense fuzzy coat. Some mi-crotubules (MT) and vesicles (V) are associated with the wide tubularnetwork.(C) Different membranous domains in a syncytial-type cell plateshowing the transition from the wide tubular network to the thin tu-bular network (W-TTN) and some noncoated thin tubules corre-sponding to the thin tubular network (TTN). Note that the fuzzy coatis disassembled during the conversion of wide tubules into thin tu-bules.Bars in (A) to (C) 5 200 nm.

940 The Plant Cell

Figure 6. Syncytial-Type Phragmoplast and Cell Plate.

(A) Transverse section of a syncytial-type cell plate. Wide tubules (WT) are seen in the tubular network. Golgi-derived vesicles (V) are associatedwith mini-phragmoplast microtubules (arrows) at the growing cell plate margin. Note that vesicles do not seem to fuse directly with the cell platenetwork but instead fuse first with each other (arrowhead). A mitochondrion (M) and a Golgi stack (G) seem close to the cell plate.(B) Tangential section of a syncytial-type cell plate. Mini-phragmoplasts consisting of 4 to 12 microtubules (dotted circles) are located discontinuouslyalong the margin of the growing cell plate. Some single microtubules (arrows) are interspersed among tubular elements. The nonuniform growth andmaturation of these cell plates are evidenced by the presence of thin tubules (TT) between wide tubular domains (WT). Wide tubule to thin tubule tran-sition sites are indicated by open arrows. Clathrin-coated membrane buds (CB) are associated with thin tubules. FC, fuzzy coat.Bars in (A) and (B) 5 500 nm.

Syncytial-Type Cell Plate in Arabidopsis Endosperm 941

plate formation (Lukowitz et al., 1996; Lauber et al., 1997).Because syncytial-type cell plates are also derived fromhomotypic fusions of Golgi-derived vesicles, one would ex-pect KNOLLE protein or a close homolog to be present informing syncytial-type cell plates. This prediction has beenverified in cellularizing Arabidopsis endosperm cells bymeans of anti-KNOLLE protein antibodies (Lauber et al.,1997). It will be interesting to determine if the

knolle

mutantdisplays the same ultrastructural alterations in syncytial- andsomatic-type cell plates preserved by high-pressure freez-ing/freeze-substitution techniques.

The centrifugal growth of cell plates involves the incorpo-ration of new Golgi-derived vesicles into the cell plate mar-gin.

In syncytial-type cell plates, the vesicles do not seem tofuse directly with the cell plate but instead fuse first witheach other to form wide tubules. Only after vesicles fuse intotubules do they appear to gain the ability to fuse with ele-ments of the wide tubular network in the growing cell platemargin. Thus, incorporation of new material into the growingcell plate seems to encompass two steps: the homotypic fu-sion between Golgi-derived vesicles and the homotypic fu-sion between wide tubules. Two other homotypic fusionsteps occur during syncytial-type cell plate formation: fusionof the early tubular networks of adjacent mini-phragmo-plasts, and fusion of adjoining cell plates at the sites offuture cell corners to form the honeycomb-like wall configu-rations.

Conserved Structures Allow for the Identification of Corresponding Stages in Somatic- and Syncytial-Type Cell Plates

The assembly of both somatic- and syncytial-type cellplates involves transient membrane networks that undergoprogrammed changes during cell plate maturation, but asdocumented in this report, the geometries of these networksdiffer between the two systems. In view of the appearanceof a characteristic, dense fuzzy coat on both the wide tubu-lar network from syncytial-type cell plates (Figure 5B) andthe tubulo-vesicular network from somatic-type cell plates(Samuels et al., 1995), we postulate that these two mem-brane networks represent corresponding stages in cell plateformation. The assembly of a very dense fibrous coat ontothe surface of the initial membrane networks may serve tomechanically stabilize these delicate membrane systemsuntil the cell wall–forming polysaccharides can take over thisfunction (Staehelin and Hepler, 1996). The persistence of adense fibrous coat on the wide tubules throughout thelengthy wide tubular network stage of syncytial-type cellplate formation (Figures 5B, 6A, and 6B) supports the argu-ment of the membrane-strengthening function of the fibrouscoat material. With the importance of this fuzzy coat materialnow firmly established for both types of cell plates, futurestudies need to identify the molecules involved in the forma-tion of this coat.

The next stage in cell plate maturation, loss of the fuzzycoat and conversion of the associated membrane systemsinto a thin tubular network, is another feature in common inthe two types of cell plates (Figures 5C and 6B; Samuels etal., 1995, Figures 4 and 5B). These changes are accompa-nied by two other developments in common: the appear-ance of clathrin-coated buds and vesicles, and the rapidbuildup of callose deposits within the thin tubules. Whereasthe former structures appear to be responsible for removingselected membrane proteins and excess membrane lipidsfrom the cell plate networks, the initial accumulation of cal-lose within the membrane tubes may provide the mechani-cal support that allows the safe dismantling of the fuzzycoat.

Syncytial-Type Cell Plates Mature Heterogeneously and More Slowly Than Do Somatic-Type Cell Plates

Syncytial-type cell plates mature in a patchwork manner,showing different domains at various developmental stagesat any given time. This particular pattern of maturation is re-lated to the fact that these cell plates are produced by multi-

Figure 7. Syncytial-Type Cell Plates Ready to Fuse with a SyncytialCell Wall in the Cytoplasmic Domain That Does Not Abut the CentralVacuole.

See Figure 1 for an overview. Endosperm nuclei are indicated by as-terisks. Syncytial-type cell plates are marked by arrows. E, embryoproper; EN, endothelium; S, suspensor. Bar 5 10 mm.

942 The Plant Cell

ple mini-phragmoplasts that act in concert but, because ofthe small number of microtubules, have limited capabilitiesfor vesicle transport. Thus, initially, at the onset of cellular-ization, each mini-phragmoplast gives rise to its own tubularnetwork, which then expands and fuses with other tubularnetworks to establish a large consolidated tubular cell platenetwork (Figures 6B and 8) that can be converted into a newcell wall. This multiple origin is one of the principal causes ofthe patchwork maturation of syncytial-type cell plates, but itis not the only one. A second cause is the mechanism of lat-eral cell plate enlargement, which continues to involve multi-ple independent mini-phragmoplasts located around the cellplate margins. Because transport of vesicles to the cell plateappears to occur along the microtubules of these mini-phragmoplasts, and because vesicle fusion and fusion of

wide tubules with the growing edge also occur close to themini-phragmoplasts, the enlargement of cell plates appearsto be confined to localized domains around their margins.

This localized growth phenomenon leads to the questionof how cell plate expansion is controlled. Given the discretenature of the mini-phragmoplasts associated with syncytial-type cell plates, and considering the short life (

t

1/2

) of the mi-crotubules (Hepler and Hush, 1996), the even growth of thecell plate margins could involve either gradual, microtubuleturnover–mediated shifts in mini-phragmoplast distributionor the complete breakdown of entire mini-phragmoplastsand the creation of new ones in different locations. Only ob-servations on appropriately labeled living cells will enable re-searchers to distinguish between these alternatives.

Our micrographs also demonstrate that syncytial-type cellplates mature more slowly than somatic-type cell plates.This is shown by the extended duration of the wide tubularnetwork stage of syncytial-type cell plate formation, even af-ter fusion of adjacent syncytial-type cell plates (Figure 8).After all adjoining cell plates have linked to form a honey-comb-like configuration, they fuse with the parental plasmamembrane and the syncytial cell wall. This fusion step coin-cides with conversion of the tubular network cell plate into afenestrated sheet and, finally, into an anticlinal cell wall. A

Figure 9. Fusion of Syncytial-Type Cell Plates with the SyncytialCell Wall.

(A) Transverse section of a flattened fenestrated sheet (FS) that hasalready fused (arrowhead) with the syncytial cell wall (SCW) facingthe embryo (E).(B) Tangential section of a syncytial-type cell plate at the fenestratedsheet stage. Some tubules (arrows) are still observed. Clathrin-coated structures associated with the cell plate and multivesicularbodies (MVB) are commonly observed at this stage.CB, clathrin-coated bud; FS, fenestrated sheet. Bars in (A) and(B) 5 100 nm.

Figure 8. Fusion of Syncytial-Type Cell Plates.

(A) Syncytial-type cell plates (arrows) forming simultaneously aroundan endosperm nucleus (N). All of the cell plates exhibit a tubular net-work architecture, and where they have joined, they give rise to Y- orT-shaped junctional domains. EN, endothelium; VA, vacuole.(B) Detail of the network (arrows) resulting from the fusion of severalsyncytial-type cell plates.Bars in (A) and (B) 5 2 mm.

Syncytial-Type Cell Plate in Arabidopsis Endosperm 943

similar pattern of cell plate maturation after fusion with theparental cell wall has been demonstrated during the finalphases of somatic-type cell plate assembly (Samuels et al.,1995) and has been related to the presence of maturationalfactors in the cortical division site (Mineyuki, 1999).

Xyloglucans in Syncytial-Type Cell Plates and Endosperm Cell Walls Are Devoid of Fuscose Residues

The strongest and most consistent labeling of all stages ofsyncytial-type cell plate formation as well as of Golgi stacksand Golgi-derived vesicles was obtained with the anti-xylo-glucan backbone antibodies. This result is consistent withprevious observations on somatic-type cell plates (Moore etal., 1986) and with the finding that xyloglucan is synthesizedby Golgi-localized enzymes and delivered to the cell plate invesicles (Moore et al., 1986; Zhang and Staehelin, 1992). Incontrast, the same structures are not labeled with the anti-xyloglucan side chain antibody CCRC-M1, which recog-nizes the terminal fucose residues on the trisaccharide side

Figure 10. Labeling Cell Plates and Cell Walls with Anti-Xyloglucanand Anti-Pectic Polysaccharide Antibodies.

(A) Transverse section of a wide tubular network and Golgi stacks(G) labeled with anti-xyloglucan antibody (ANTI-XG). Arrows indicatelabeled Golgi stack.(B) Section labeled with CCRC-M1 (anti-xyloglucan fucose residues)antibodies. Note the lack of label on the syncytial-type cell plate(SCP) and the syncytial cell wall (SCW) and the strong labeling of theadjacent endothelium cell walls (CW). The arrowhead indicates thesite of fusion between the syncytial-type cell plate and the syncytialcell wall. EN, endothelium; END, endosperm.(C) Transverse section of a recently formed anticlinal cell wall la-beled with JIM7. VA, vacuole.Bars in (A) to (C) 5 200 nm.

Figure 11. Labeling Cell Plates and Cell Walls with Anti-Callose An-tibodies and CBHI-Gold Probe.

(A) Tangential section of a thin tubular network and some wide tu-bules labeled with anti-callose antibodies. Two clathrin-coatedmembrane structures (CB) are seen budding off some tubules in thecell plate.(B) Transverse section of a wide tubular network labeled with CBHI-gold (arrows).Bars in (A) and (B) 5 200 nm.

944 The Plant Cell

chain of xyloglucan (Puhlman et al., 1991). Given that all ofthe walls of the cells surrounding the endosperm were posi-tively labeled with this antibody, we can rule out the possi-bility that CCRC-M1 antibodies were unable to bind thexyloglucan–fucose epitope in our samples. Thus, we con-clude that the xyloglucan molecules produced by Arabidop-sis endosperm lack terminal fucose residues.

This finding is not completely unexpected, because theseed storage form of xyloglucans in nasturtium (Tropaeolummajus), legumes, and other plants is essentially devoid ofterminal fucose residues (Reid, 1985). However, the nastur-tium embryo produces two kinds of xyloglucans during de-velopment (Desveaux et al., 1998): a structural fucosylatedxyloglucan that is delivered to the primary cell wall, and astorage nonfucosylated xyloglucan that accumulates in em-bryo cells in the periplasmic space between the plasmamembrane and the primary cell wall. Our labeling data showthat in contrast to nasturtium, in Arabidopsis endospermneither the xyloglucans associated with the primary cellwalls nor the xyloglucans in the cell plates become fucosy-lated. The absence of labeling with the CCRC-M1 antibod-ies on Golgi stacks, Golgi-derived vesicles, and cell plates in

Arabidopsis endosperm supports the idea that neither trans-port nor deposition of xyloglucan in the extracellular matrixinvolves transient fucosylation of xyloglucan followed by adefucosylation step.

The lack of terminal fucose residues on the trisaccharideside chain of xyloglucan has several interesting functionalimplications. As shown by Levy et al. (1991, 1997), the pres-ence of these terminal fucose residues leads to the stabiliza-tion of the straight, flat conformation of xyloglucan, whichcan bind to cellulose fibrils. In the absence of these fucoseresidues, the xyloglucan molecules tend to assume atwisted conformation that cannot bind to cellulose. The in-ability to bind to cellulose makes such fucose-less xyloglu-can more accessible to hydrolytic enzymes and thereforemore suitable as a storage form of polysaccharide. At thesame time, the lack of stable xyloglucan binding to cellulosemicrofibrils will lead to cell walls that are less cross-linkedand hence mechanically weaker, as evidenced by the de-creased tensile strength of the cell walls of the mur1 cell wallmutant (Zablackis et al., 1996). Based on these consider-ations, the cell walls of Arabidopsis endosperm probablyalso lack tensile strength, but because the endosperm issurrounded and protected by mechanically strong tissues,this weakness seems to have no deleterious consequences.Indeed, the lack of strength and rigidity may be critical forendosperm function, given that the endosperm cell wall hasto expand locally and give way as the embryo enlarges andimpinges on the endosperm space.

Callose Serves Different Functions in Syncytial-Type Cell Plates and Endosperm Cell Walls

Callose is first detected during the wide tubular networkstage of syncytial-type cell plate formation, increases duringthe formation of the thin tubular networks and fenestratedsheets, and persists in the mature endosperm cell wall.Whereas these findings parallel those reported for somatic-type cell plates, the presence of callose in the mature cellwalls constitutes a unique property of endosperm cell walls(Stone and Clarke, 1992; Brown et al., 1997). The callose as-sociated with somatic-type cell plates has been postulatedto perform two functions (Samuels et al., 1995): (1) to me-chanically stabilize the delicate membrane tubules as thefuzzy coat is dismantled and the cell plate–forming mem-branes are matured by the selective removal of membranecomponents by way of clathrin-coated vesicles, and (2) toprovide the spreading force that widens the tubules andconverts the network into a fenestrated sheet and a continu-ous cell wall. Support of these interpretations has comefrom studies with the drug caffeine, which prevents callosesynthesis when somatic-type cell plates are forming andleads to their fragmentation, and from the demonstrationthat callose forms a membrane surface–adhering layer dur-ing the formation of fenestrated sheets (Samuels et al.,1995; Samuels and Staehelin, 1996). Given the similarity of

Figure 12. Diagram Showing Microtubule Organization in the Syn-cytial Endosperm at the Onset of Cellularization and the Putative Or-igin of the Mini-Phragmoplasts from Opposing Overlapping Clustersof Microtubules.

Microtubules radiating from microtubule-organizing centers (MTOCs)in the nuclear envelope define the nuclear cytoplasmic domains(NCDs) and the sites of syncytial-type cell plate formation. MP, mini-phragmoplasts; N, nucleus.

Syncytial-Type Cell Plate in Arabidopsis Endosperm 945

the membrane structures with which callose is associatedduring syncytial-type cell plate formation and given the ki-netics of callose deposition, we postulate that callose per-forms the same membrane stabilization and thin tubuleexpansion functions in both types of cell plates.

However, whereas callose disappears from somatic-typecell plates during their conversion into cell walls, the callosein syncytial-type cell plates not only persists throughout allstages of cell plate formation but also is maintained after cellwalls have matured. The presence of callose in mature en-dosperm cell walls has also been reported for rice and othercereals (Stone and Clarke, 1992; Brown et al., 1997). Onepossible function of callose in endosperm cell walls is toserve, like the fucose-less xyloglucan, as a carbohydrate re-serve for the growing embryo. In addition, because of itsgel-like physical properties, callose could further increasethe plasticity of the endosperm cell walls to allow for the re-modeling of the endosperm tissue around the rapidly ex-panding embryo.

METHODS

Seeds of Arabidopsis thaliana (Landsberg erecta wild type) wereplanted in Metro-mix 200 growing medium (America Clay Works,Denver, CO) with Arabidopsis controlled-release fertilizer 17-6-12 1minor (Lehle Seeds, Round Rock, TX). Plants were grown in continu-ous fluorescent lighting, at a temperature of 27 6 18C and a relativehumidity of 45%. Developing seeds were excised at various hoursafter anthesis.

High-Pressure Freezing/Freeze Substitution

Whole developing seeds were removed from the siliques. They wereimmediately loaded into sample hat holders that had been coatedwith a solution of 100 mg of lecithin per mL of chloroform. The hold-ers were filled with a solution of 15% dextran or 0.1 M sucrose. Thesamples were frozen in a Baltec HPM 010 high-pressure freezer(Technotrade, Manchester, NH) and then transferred to liquid nitrogenfor storage. Substitution was performed in 2% OsO4 in anhydrous

Figure 13. Stages of Syncytial-Type Cell Plate and Cell Wall Formation.

The model depicts the main stages of syncytial-type cell plate and cell wall formation in Arabidopsis endosperm, with a newly formed anticlinalcell wall connecting the syncytial cell walls facing the endothelium and the embryo (E). CB, clathrin bud; CW, cell wall; FC, fuzzy coat; HGI, hour-glass intermediate; M, ribosome-excluding matrix; MP, mini-phragmoplast; MT, microtubules; PM, plasma membrane; SCW, syncytial cell wall;V, vesicles; WT, wide tubules.

946 The Plant Cell

acetone at 2808C for 72 hr, followed by slow warming to room tem-perature over 2 days. After several acetone rinses, some sampleswere teased from the holders and infiltrated in Epon or Spurr’s resin(Ted Pella) according to the following schedule: 5% resin in acetone(4 hr), 10% resin (12 hr), 25% resin (12 hr), 50% (24 hr), 75% (24 hr),and 100% (24 hr). Polymerization was performed at 708C for Eponand 608C for Spurr’s resin. Other samples were washed in ethanol af-ter the acetone rinse and then embedded in London Resin White(Electron Microscopy Sciences) (25% in absolute ethanol, 50%,75%, and 100%; 12 hr at each concentration); polymerization wasperformed at 608C for 24 hr.

Immunolabeling and Cellobiohydrolase I–Gold Probe

Callose was localized by using a monoclonal antibody to b-1,3-glu-can from Biosupplies Australia (Parkville, Victoria, Australia). Xyloglu-cans were localized by using the polyclonal antibody anti-xyloglucan(Moore et al., 1986) and the CCRC-M1 monoclonal antibody (Puhlmanet al., 1991). Methyl-esterified pectins were localized by using theJIM7 monoclonal antibody (Knox et al., 1990).

Cellobiohydrolase I–gold was prepared as described by Krishnamurthy(1999). Cellobiohydrolase I was a gift of Dr. M. Shulein (Novo Nordisk,Danbury, CT) and was conjugated to 15-nm-diameter colloidal gold(BBInternational; Ted Pella).

Sections from Epon resin blocks (anti-callose, anti-xyloglucan, andCCRC-M1) or from LR White–embedded material (JIM7) were col-lected on formvar-coated nickel grids. Samples were blocked for 30min with a 5% (w/v) solution of nonfat dried milk in PBST (Tween-20,0.1%) (for anti-callose antibody and anti-xyloglucan antibody) or a3% (w/v) solution of the same components (for CCRC-M1 and JIM7).Primary antibodies anti-callose (1:50 in PBST), anti-xyloglucan (1:10in PBST), CCRC-M1 (1:10 in PBST), and JIM7 (1:1) were applied for45 to 60 min at room temperature (anti-callose and anti-xyloglucan)or overnight at 48C (CCRC-M1, JIM7). The sections were washed in astream of PBST containing 0.5% Tween-20 and then transferred tothe secondary antibodies (goat anti–mouse IgG for anti-callose andCCRC-M1, goat anti–rabbit IgG for anti-xyloglucan, and goat anti–ratIgG for JIM7) conjugated with 15-nm-diameter colloidal gold for 1 hr.

Controls in each case received the same treatments except thatthe primary antibodies were omitted.

In all cases, sections 70- to 90-nm thick were cut on a Leica Ul-tracut R and then stained with 2% uranyl acetate in 70% methanolfor 10 min followed by Reynold’s lead citrate (2.6% lead nitrate and3.5% sodium citrate, pH 0.12) for 4 min. Samples were observedwith a Philips (Hillsboro, OR) CM10 microscope. In some cases, se-rial sections were studied to try to elucidate the three-dimensionalconfiguration of some cellular structures.

ACKNOWLEDGMENTS

We thank Dr. Andreas Nebenfüehr for valuable comments on themanuscript. The donation of cellobiohydrolase I by Dr. M. Shulein(Novo Nordisk) is gratefully acknowledged. This work was supportedby National Institutes of Health Grant No. 18639 and U.S. Depart-ment of Agriculture Grant No. 96-35304-3710 to L.A.S. andFullbright/Antorchas Foundation and Comision Nacional Investiga-ciones Cientificas y Tecnicas, Argentina, Postdoctoral Fellowshipsto M.O.

Received February 2, 2000; accepted April 20, 2000.

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DOI 10.1105/tpc.12.6.933 2000;12;933-947Plant Cell

Marisa Otegui and L. Andrew StaehelinArabidopsis

Syncytial-Type Cell Plates: A Novel Kind of Cell Plate Involved in Endosperm Cellularization of

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