CELL BIOLOGY AND MORPHOGENESIS

Ultrastructure and histochemical analysis of extracellular matrixsurface network in kiwifruit endosperm-derived callus culture

M. Popielarska-Konieczna Æ M. Kozieradzka-Kiszkurno Æ J. Swierczynska ÆG. Goralski Æ H. Slesak Æ J. Bohdanowicz

Received: 30 January 2008 / Accepted: 29 February 2008

� Springer-Verlag 2008

Abstract The study used Actinidia deliciosa endosperm-

derived callus to investigate aspects of the morphology,

histology and chemistry of extracellular matrix (ECM)

structures in morphogenically stable tissue from long-term

culture. SEM showed ECM as a membranous layer or

reticulated fibrillar and granular structure linking the

peripheral cells of callus domains. TEM confirmed that

ECM is a distinct heterogeneous layer, up to 4 lm thick

and consisting of amorphous dark-staining material,

osmiophilic granules and reticulated fibres present outside

the outer callus cell wall. ECM covered the surface of cells

forming morphogenic domains and was reduced during

organ growth. This structure may be linked to acquisition

of morphogenic competence and thus may serve as a

structural marker of it in endosperm-derived callus. ECM

was also observed on senescent cells in contact with the

morphogenic area. Treatment of living calluses with

chloroform and washing with ether–methanol led to partial

destruction of the extracellular layer. Digestion with pec-

tinase removed the membranous layer almost completely

and exposed thick fibrillar strands and granular remnants.

Digestion with protease did not visibly affect the surface

layer. Indirect immunofluorescence showed low-methy-

lesterified pectic epitopes labelled by JIM5 monoclonal

antibody. Immunolabelling, histochemistry, and solvent

and enzyme treatments suggested pectins and lipids as

components of the surface layer. These compounds may

indicate protective, water retention and/or cell communi-

cation functions for this external layer.

Keywords Actinidia deliciosa � Morphogenesis �Pectin epitopes � Scanning electron microscopy �Transmission electron microscopy

Abbreviations

2,4-D 2,4-Dichlorophenoxyacetic acid

BSA Bovine serum albumin

CPD Critical point drying

DAPI 40, 6-Diamidino-2-phenylindole dihydrochloride

ECM Extracellular matrix

ECMSN Extracellular matrix surface network

EGTA Ethylene glycol-bis(b-aminoethyl ether)N, N,

N0, N0-tetraacetic acid

PBS Phosphate-buffered saline

PIPES Piperazine-N, N0-bis(2-ethanesulfonic acid)

SB Stabilising buffer

SEM Scanning electron microscopy

TEM Transmission electron microscopy

Introduction

The cell wall of higher plants is formed of dynamic cellular

compartments with structural, protective and growth reg-

ulation functions (Baluska et al. 2003). This active

structure, referred to as the extracellular matrix (ECM)

(Roberts 1994), is an integral part of the ECM–plasma

membrane–cytoskeleton continuum, and plays a funda-

mental role in the reception and transduction of signals

Communicated by D. Somers.

M. Popielarska-Konieczna (&) � G. Goralski � H. Slesak

Department of Plant Cytology and Embryology,

Jagiellonian University, 52 Grodzka St., 31-044 Cracow, Poland

e-mail: m.popielarska-konieczna@uj.edu.pl

M. Kozieradzka-Kiszkurno � J. Swierczynska � J. Bohdanowicz

Department of Plant Cytology and Embryology,

University of Gdansk, 24 Kładki St., 80-822 Gdansk, Poland

123

Plant Cell Rep

DOI 10.1007/s00299-008-0534-9

connected with positional information, recognition, cell

fate determination and consequently plant development.

Recently, the extracellular matrix surface network (EC-

MSN) has drawn considerable attention from researchers.

The chemical composition and structural arrangement of

ECMSN on the cell surface may play a significant role in

morphogenic processes (Verdeil et al. 2001). The external

cell wall is particularly interesting in this regard, as it is

exposed to environmental factors.

Cell and tissue culture induces different cellular

responses, which mediate adaptations under new environ-

mental conditions. One of them may be the formation of

the ECM surface layer or network on the outer cell wall.

Bobak et al. (2003/4) suggested that ECM formation could

be a kind of stress response of explants elicited by specific

culture conditions. Secretion of ECM could, for example,

protect the surface of callus clusters.

The secretions may also play a role in cell integration

and recognition. Cell morphology and synchrony during

plant regeneration in vitro are controlled by particular

morphogenic programs within which cell–cell communi-

cation is critical to recognition of a cell’s position and

behaviour in early stages of regeneration. Culture condi-

tions may act as a stress signal inducing a morphogenic

program. In such a case, plant cells may follow a devel-

opmental pathway that leads to organogenesis. The

mechanisms that commit a single somatic plant cell to

develop as a morphogenic structure are still poorly

understood and invite further investigations.

Organogenesis (shoot and root formation) and somatic

embryogenesis in vitro have several common features,

including the occurrence of the extracellular matrix.

Organogenesis is considered to be a morphogenic pathway

by which a cell fully expresses its totipotency. Organo-

genesis (as well as somatic embryogenesis) requires a

certain degree of cell dedifferentiation, reinitiation of cell

division, and morphogenic control over cell multiplication

(Pret’ova et al. 2006).

Pectic polysaccharides, which play important structural

roles in the control of cell wall porosity, elasticity and cell–

cell adhesion, are assumed to participate in various dif-

ferentiation mechanisms such as cell elongation, growth

and differentiation (for review see: Willats et al. 2001;

Jarvis et al. 2003). Pectin oligosaccharide fragments also

function as signalling molecules involved in the regulation

of developmental processes (Dumville and Fry 2000).

Several studies have shown that morphogenic processes

such as embryogenesis and organogenesis require pectic

intercellular connections. In suspension-cultured cells of

carrot, Kikuchi et al. (1996) reported significant differences

in the sugar composition of pectic chains between the

stages of the cell developmental pathway. Differences in

methyl esterification of homogalacturonan were found to

accompany somatic embryogenesis competence in Cicho-

rium (Chapman et al. 2000c), Cocos (Verdeil et al. 2001)

and Triticum (Konieczny et al. 2007).

Previous research described the membranous structure

on the surface of endosperm-derived callus in kiwifruit

(Popielarska et al. 2006). Our present work is a detailed

study of ECM formed during culture, using scanning

electron microscopy (SEM) and transmission electron

microscopy (TEM), and a preliminary study by indirect

immunofluorescence microscopy.

Materials and methods

Plant material and culture conditions

Long-term culture (more than 24 months) of endosperm-

derived callus of Actinidia deliciosa cv. Hayward was

initiated and maintained as described previously (Goralski

et al. 2005). Briefly, commercial fruits were kept at room

temperature to allow softening. Pieces of fruit (3 9 3 cm)

were surface-sterilised for 12 min in Ace commercial

bleach diluted 1:1 (v:v) with distilled water and rinsed

three times in sterile distilled water. Endosperm was iso-

lated from seeds and cultured on MS medium (Murashige

and Skoog 1962) with kinetin (5 mg l-1) and 2,4-D

(2 mg l-1) (AEM, Actinidia endosperm medium) to induce

callus. The endosperm-derived callus maintained long-term

on AEM preserved its capacity for regeneration on MS

medium supplied with TDZ (0.5 mg l-1) (RM, regenera-

tion medium). The cultures were incubated at 26 ± 3�C

under a 16 h photoperiod (cool-white fluorescent tubes,

60–90 lmol photons m-2 s-1).

Scanning electron microscopy

Callus samples were prefixed in 5% buffered glutaralde-

hyde (0.1 M phosphate buffer, pH 7.2) for 2 h at room

temperature. After dehydration through a graded ethanol

series, samples were dried with a CPD (CO2 critical-point

drying) system, sputter-coated with gold (Jeol JFC-1100 E

ion-sputtering system) and observed with a scanning

electron microscope (HITACHI S-4700).

Transmission electron microscopy

Calluses were fixed in 2.5% formaldehyde (prepared from

paraformaldehyde) and 2.5% glutaraldehyde in 0.1 M

cacodylate buffer (pH 7.0) for 2 h at room temperature.

The samples were rinsed in the same buffer with four

changes (15 min each) and postfixed in buffered 1% OsO4

at 4�C overnight. After rinsing in distilled water, the cal-

luses were treated with 1% uranyl acetate in distilled water

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for 1 h, dehydrated in a graded acetone series and

embedded in Spurr’s resin. Ultrathin sections were cut on a

Sorvall MT-2B ultramicrotome, stained with uranyl acetate

and lead citrate, and examined with a Philips CM 100

transmission electron microscope. Control semithin sec-

tions were poststained with 0.1% Toluidine Blue O and

Sudan Black B (nonspecific staining of lipids).

Solvent and enzyme treatment

To test the chemical composition of the outer surface layer,

calluses of living material were treated with different sol-

vents and enzymes, after which the calluses were fixed in

glutaraldehyde and prepared with a CPD system for SEM

observations.

Lipids were removed by washing the calluses for 15 and

30 min in chloroform and an ether–methanol (1:1, v/v)

mixture. Enzymatic digestion of pectic compounds was

achieved with 1 mg l-1 pectinase (pectinase from Asper-

gillus niger, Fluka) in 0.1 M phosphate buffer (pH 7.5) at

30�C for 30 and 45 min. Protein digestion was achieved

with 1 mg l-1 protease (Protease, type XIX: Fungal,

Sigma) dissolved in 0.1 M phosphate buffer (pH 7.5) at

30�C for 2, 3 or 4 h. Samples in buffer alone were the

control.

Immunolabelling and staining of tissue sections

Small clumps of calli were excised and fixed with 4%

formaldehyde (prepared from paraformaldehyde) and

0.25% glutaraldehyde in stabilising buffer (SB; 50 mM

PIPES buffer, 1 mM MgCl2, 10 mM EGTA, pH 6.8) for

2 h. After washing in PBS, samples were dehydrated in a

graded ethanol series diluted with PBS. Tissue was

embedded in Steedman’s wax as described by Baluska

et al. (1992). In brief, wax was prepared from PEG 400

distearate and 1-hexadecanol (9:1). Tissue was embedded

at 37�C and the wax left to polymerise at room tempera-

ture. Sections of calli 5 lm thick were mounted on slides

coated with Mayer’s egg albumen, dewaxed in absolute

alcohol, passed through a graded ethanol–PBS series and

rinsed in PBS. Specimens were blocked with 5% BSA in

PBS for 1 h at room temperature and incubated in JIM5

monoclonal antibody (diluted 1:20) for 1.5 h. After several

washings, the sections were incubated with anti-rat anti-

body conjugated with Alexa fluor 488 (diluted 1:800) for

1.5 h. Following several washes in PBS, the nuclei were

fluorescence-stained by DAPI for 10 min. Sections were

then stained for 10 min in 0.01% Toluidine Blue to

diminish the autofluorescence of callus tissues. After

washing in PBS, the sections were mounted under cover-

slips using Citifluor antifading solution. Controls were

prepared by omitting the first antibody; no pectin staining

was detected.

Control tissue sections of calli embedded in Steedman’s

wax were treated with Calcofluor White and Aniline Blue

for staining of cellulose and callose, respectively.

Dewaxed and rehydrated sections were stained in 0.1%

Calcofluor White for 15 min at room temperature, washed

with ddH2O and immediately observed using UV light

excitation. Aniline Blue solution (0.005% in 0.15 M

K2HPO4, pH 8.2) was used to examine callose deposition

in callus tissues.

Results

Scanning electron microscopy

The surface of morphogenic callus domains was coated

with secretions. The ECM over callus cells varied in

structure (Fig. 1a–h). The external matrix appeared

mostly as a compact membranous layer (Fig. 1a, c, d). In

some regions not covered by the membranous layer,

fibrillar structures of the extracellular network (especially

between cells; Fig. 1b) and granular mucilage-like secre-

tions on the cell surface were exposed. Transitions from

compact to fibrillar structure were also noted (Fig. 1e, f).

Fragments of membranous layer were observed in shoot

regions, but did not cover the developing organs. This

type of ECM coated callus domains and round paren-

chymatic cells (single or in clusters) localised in

morphogenic domains (Fig. 1g, h).

Transmission electron microscopy

TEM of peripheral callus cells showed vesicles and chan-

nels of endoplasmic reticulum parallel to and in the close

vicinity of the cell wall. Mitochondria were round or oval

in section. Dictyosomes and numerous ribosomes were

observed (Fig. 2a–c). Microbodies and plastids with a

simple system of lamellae and a simple large starch grain

were present occasionally (Fig. 2d). At the plasmalemma,

exo- or endocytosis was observed, giving the plasmalemma

a crenelated appearance (Fig. 2b–d, f). Adjacent to the

inner surface of the cell wall were vesicles which may be

from Golgi bodies, indicating some exocytotic activity

(Fig. 2d). Endocytosis was seen as the formation of a

coated vesicle from a coated pit (Fig. 2f). Large vacuoles

and sequestration of cytoplasm were noted in some

peripheral cells (Fig. 2e).

More detailed TEM showed a conspicuous layer

composed of amorphous material with fibrillar and

spherical components, covering morphogenic cell clusters

from long-term culture (Fig. 2a–e). In some cases it

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contained amorphous dark and grey patches or globular

deposits similar in electron density and appearance to

lipid bodies. Under high magnification there were many

clearly visible small osmiophilic granules attached to

fibers extending from the outer cell wall of the callus cell

(Fig. 2a–c). The fine ECM layer featured grey granulation

Fig. 1 Electron micrographs of

callus surface and

organogenesis (a–h). Callus

surface was covered with

membranous (indicated by

ecm), fibrillar (indicated by

arrows) and granular (indicated

by arrowheads) structures (a–f).Fibrillar structures and granules

of mucilage-like secretion

forming a network at site of

cell–cell adhesion (b).

Enlargement of detail c (in

window) shows fibrillar nature

(f) of cell surface enveloped

with membranous component of

ECM (d). Transformation of

membranous-fibrillar structures

of ECM (e–f). Organogenic

structures (o), trichomes (t),remnants of membranous

structures (short arrows) and

parenchymatic cells (asterisks)

adjacent to developing organ

and callus domains (g–h). Barsrepresent 2 lm for b, d and f;10 lm for c and e; 50 lm for h;

100 lm for a; and 500 lm for g

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Fig. 2 Ultrastructure of callus

surface (a–e) and intercellular

space (f). Osmiophilic dark

granules and grey patches or

deposits (indicated by arrows)

as components of ECM (ecm)

attached to fibers (indicated by

f) extending from outer cell wall

(indicated by cw) (a).

Reticulated appearance of

extracellular matrix (b–c); note

crenelated plasmalemma

(indicated by arrowheads) and

fibers extending from outer cell

wall; visible vesicles and

channels of endoplasmic

reticulum (indicated by er), part

of dictyosome (d) and

mitochondrion (indicated by m)

(c). Vesicles (asterisks) adjacent

to inner cell wall (d); visible

plastid (p) with lamellae and

starch grains, microbody (b),

mitochondria, lobes (n) of

nucleus, numerous ribosomes

and fine extracellular matrix.

Parenchymatic, senescent cell

was covered with granular,

amorphous and ‘‘peeling’’

extracellular matrix (e); sindicate sequestration of

cytoplasm and v vacuole.

Ultrastructure of intercellular

space (is) (f); note crenelated

plasmalemma with coated pits,

endoplasmic reticulum and

electron-dense layer (shortarrows) as part of cell wall.

Bars represent 1 lm for a–dand f; and 5 lm for e

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and more or less osmiophilic patches against a relatively

electron-transparent background (Fig. 2d). The cell wall

of senescent cells was also covered with ECM, but with

an amorphous, granular and ‘‘peeling’’ appearance

(Fig. 2e). In larger intercellular areas near the callus

surface the fibril-like material filling the space had a

different appearance. The fibrillar structures composing

the network were adjacent to an electron-dense layer of

the cell wall (Fig. 2f).

Solvent and enzyme treatment

Treating living callus with protease did not remove the

surface layer (Fig. 3a, b), but washing in chloroform for

15 min altered the external matrix pattern: the coherent

layer of secretion clearly visible before treatment showed

many small holes (Fig. 3c), which increased in number and

size after extraction for 30 min (Fig. 3d–f). Digestion of

living kiwifruit callus with pectinase for 30 and 45 min

caused considerable but not complete degradation of the

membranous layer (Fig. 3g), exposing thick fibrils formed

of thin filaments and granular remnants (Fig. 3h, i).

Immunolabelling and staining of tissue sections

The membranous layer covering the callus surface pos-

sessed pectins recognized by the monoclonal antibody

JIM5 (Fig. 3j, k), but larger intercellular spaces also

showed JIM5-labelled low-esterified pectins at the cell wall

junctions. Cellulose microfibrils and hemicellulose in the

cell wall were marked by Calcofluor White (b-1,4-glucan

binding agent). After its application, no fluorescence of the

surface layer appeared (Fig. 3l). Staining with Aniline Blue

revealed that the cell wall of some cells contained a callose

component. Single or unorganised groups of cells (data not

shown) were observed near the callus surface.

Discussion

Our previous histological analysis and SEM study of

morphogenic endosperm-derived callus of A. deliciosa

described the membranous layer covering the callus sur-

face and termed it extracellular matrix (ECM) (Popielarska

et al. 2006). Our present work used different microscopic

techniques to study the structure and chemical composition

of the material coating the callus surface.

SEM showed heterogeneous material covering the callus

surface. Some parts of the callus were coated by a smooth

membranous layer, other regions by fibrillar and granular

structures forming a network similar to that observed

during in vitro induction of somatic embryogenesis,

androgenesis and organogenesis in different species such as

Coffea arabica (Sondahl 1979), Cichorium (Dubois et al.

1991, Chapman et al. 2000a, b), Papaver somniferum

(Ovecka and Bobak 1999), Cocos nucifera (Verdeil et al.

2001) Fagopyrum tataricum (Rumyansteva et al. 2003),

Drosera spathulata (Bobak et al. 2003/2004), Triticum

aestivum (Konieczny et al. 2005), Brassica napus (Na-

masivayam et al. 2006) and A. deliciosa (Popielarska et al.

2006). Most data concern the formation, structure and

chemical composition and function of EMC in somatic

embryogenesis. In peripheral cells of Cichorium somatic

embryos, Chapman et al. (2000a, b) described a surface

network consisting of mucilage-like secretions and fibrillar

structures, giving the outer wall a rough texture. Bobak

et al. (2003/4, 2004) documented fibrillar ECM linking the

surface cells of somatic embryos of Drosera sphatulata.

Similarly, net-like fibrillar material was found in the gaps

between epidermal cells during acquisition of embryogenic

competence in Brassica napus embryoid culture (Namasi-

vayam et al. 2006). The fibrillar structure covered

androgenic embryos of Triticum (Konieczny et al. 2005,

2007). The induction of embryogenic competence in Cocos

calli was also associated with the presence of a fibrillar

layer fully coating embryogenic cells just before their first

division (Verdeil et al. 2001).

The data confirming the occurrence of ECM during in

vitro organogenesis are scarce (Ovecka and Bobak 1999;

Konieczny et al. 2007). Ovecka and Bobak (1999) reported

that the ECM coating adventitious buds and somatic

embryos of Papaver differed in structure. The ECM

observed during androgenic shoot and embryo develop-

ment in Triticum was uniform (Konieczny et al. 2007).

The TEM of endosperm-derived callus revealed cells

with high secretory activity, accompanied by the formation

of conspicuous ECM on their surface, characterised by the

crenelated appearance of the plasmalemma, the presence of

vesicles at the plasmalemma–cell wall interface, and

numerous ribosomes, mitochondria, and plastids with

storage starch. Peripheral parenchymatic cells had features

typical of senescent cells, such as large vacuoles and

sequestration of cytoplasm.

The presence of a distinct layer on the kiwifruit callus

surface was confirmed by TEM. This external layer,

composed of fibrils, deposits and granules, corresponds to

the membranous layer visible in SEM. Namasivayam et al.

(2006) reported similar TEM observations of ECM in

Brassica. They observed granules associated with fibres

extending from the outer cell wall, and grey patches as a

component of this outer layer. With TEM we found dif-

ferences in ECM structure depending on the callus cell

type. The cells with morphogenic ability were covered with

fine or reticulated ECM, while non-embryogenic cells and

senescent cells had granular ECM with amorphous deposits

and a ‘‘peeling’’ appearance. According to Rumyantseva

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et al. (2003), loss of embryogenic competence is preceded

by modification of ECM structure from fibrillar to gluelike,

observable by SEM. The fibrillar structures that we

observed forming a network in intercellular spaces were

similar to the reticular filling previously noted in histo-

logical sections (Popielarska et al. 2006). The electron-

dense layer between the cell wall and the network in the

intercellular spaces probably was due to condensation of

cell wall components.

The chemical composition of ECM is still not well

known. Some pectin polymers (Verdeil et al. 2001; Kon-

ieczny et al. 2007) and arabinogalactan proteins (Samaj

et al. 1999a, b; Chapman et al. 2000a; Konieczny et al.

2007) have been reported as components of secretions

covering cells with morphogenic potential.

Our data from the protease treatment showed no visible

changes in ECM appearance, in agreement with results on

Triticum (Konieczny et al. 2005). In Cichorium (Dubois

Fig. 3 SEM images obtained after treatment with different solvents

and enzymes solutions of living calluses (a–i) and fluorescence

microscopy images of the callus surface (j–l). Surface view of callus

after 4 h of protease treatment (a–b), after washing with chloroform

for 15 min (c) and 30 min (d–f); visible fibrillar structures under

partially degraded external layer (f). Disappearance of surface layer

after 45 min of digestion with pectinase (g–i); note granular remnants

(arrowheads) and thick fibrils (arrows) composed of thin filaments

(i). Immunofluorescence due to monoclonal antibody JIM5 (recog-

nizing low-esterified pectins) binding surface layer (short arrows) and

intercellular spaces (asterisks) (j–k); k combined with DAPI staining;

arrowheads indicate nuclei. Staining with the b-1,4-glucans binding

fluorescence dye, Calcofluor White (l); no fluorescence of the cell

walls. Bars represent 1 lm for b, c and f; 2 lm for e and i; 10 lm for

h; 20 lm for d and g; 50 lm for a and k; 50 lm for h; and 100 lm

for g–j and l

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123

et al. 1992), however, the proteinaceous nature of ECM

was confirmed by protease digestion. Later studies revealed

the presence of specific arabinogalactan proteins in the

ECM on the cell surface in regenerative callus of Zea

(Samaj et al. 1999a, b), Cichorium (Chapman et al. 2000a),

and Triticum (Konieczny et al. 2007).

According to Samaj et al. (1999a, b) arabinogalactan

proteins and weakly methylated pectins probably play an

important role in cell–cell adhesion and plant morpho-

genesis, so this extracellular matrix may be involved in

recognition of morphogenic cells and regulation of early

embryogenic and/or morphogenic stages.

In Triticum, solvent treatment degraded the coherent

layer of ECM to smooth-textured fibrils (Konieczny et al.

2005), but holes and other damage were not observed.

Our experiment with solvent and histochemical staining

with Sudan Black B (data not shown) suggested that

some lipophilic substances are part of the ECM in ki-

wifruit. In TEM, some globular deposits of the external

layer were similar in electron density to cytoplasmic lipid

bodies.

Pectin polymers have been found in ECM of embryo-

genic cells of Daucus (Iwai et al. 1999), Cichorium

(Chapman et al. 2000c), Cocos (Verdeil et al. 2001) and

Triticum (Konieczny et al. 2007). Fernando et al. (2007)

reported that cells involved in regeneration and formation

of organogenic protuberances in Passiflora are covered

with pectin lying just outside the cell wall. In wheat, pec-

tinase digestion resulted in complete disappearance of

ECM and the collapse of embryogenic cells (Konieczny

et al. 2005). In our experiment only partial disappearance

of ECM was observed. Pectinase treatment exposed gran-

ular remnants on the cell surface and filaments forming

thick fibrils at sites of cell adhesion, visible by SEM. We

suggest that pectin polymers form the fibrils extending

from the outer cell wall and observed in TEM as a com-

ponent of ECM. Our observations with cellulose-binding

fluorescent dye ruled out cellulose polymers as an element

of ECM structure. We did identify a callose component in

morphogenic callus domains of kiwifruit. Deposition of

such polysaccharides around embryogenic cells has been

suggested as an early structural marker for these cells

(Samaj et al. 2006).

The characteristics of pectin depend, for the most part,

on the extent of methylesterification of the carboxyl groups

of polygalacturonic acid. Numerous data indicate that the

pectic polysaccharides in the wall of young or actively

growing cells are highly methylesterified, whereas the

walls of mature or senescent cells contain highly acidic

pectins (Iwai et al. 1999). Several authors found that

labelling with JIM5, a monoclonal antibody specific for

low-esterified epitopes of pectin, is restricted mostly to

some cell junctions and the lining of intercellular spaces

between mature cells of parenchyma (Knox et al. 1990;

Guillemin et al. 2005). Pectin polysaccharides with high

methylesterification were found only in compact calli,

limited mostly to the fibrillar material filling expanded

areas between large cells at the edges of the calli (Liners

et al. 1994). In our experiment we recognised low-methy-

lesterified pectins detected with JIM5 in intercellular

spaces and also on the surface layer. Similarly, Verdeil

et al. (2001) used JIM5 antibody to detect pectin epitope in

the external coating of fibrillar matrix in coconut callus. In

Zea, however, highly esterified pectins recognised by JIM7

antibody were localised to external material and to cell–

cell adhesion sites in clumps of embryogenic cells, while

JIM5 recognising low-esterified pectins did not label EC-

MSN (Samaj et al. 2006). According to Konieczny et al.

(2007), in meristematic tissue of Triticum the outer wall of

surface cells and the continuous layer over the cells was

highly reactive to anti-pectin JIM7. The reverse was

described in Cichorium (Chapman et al. 2000c), where

immunolocalisation of an epitope recognised by JIM5

revealed that the supraembryonic network of somatic

embryos was not esterified. These results indicate that the

regulation of pectin localisation within the ECM differs

between monocotyledonous and dicotyledonous species

(Samaj et al. 2006), and our data support this.

We suggest that low-esterified pectin and probably some

lipophilic substances are components of ECM in kiwifruit-

derived callus. The ECM structure depends on cell activity

and can vary on the surface of the same piece of callus. The

possible role of the above-described external material

during kiwifruit regeneration is uncertain. Since pectins

function in cell-cell adhesion and are a reservoir of sig-

nalling molecules, ECM may be involved in integration

and recognition of morphogenic cells within multicellular

callus domains. The stress response induced by specific

culture conditions and the protective function of this

external material must also be considered.

Acknowledgments The authors are grateful to Prof. Dr. El _zbieta

Kuta (Jagiellonian University) for critically reading the manuscript

and making valuable suggestions. The SEM images were made in the

Laboratory of Field Emission Scanning Electron Microscopy and

Microanalysis at the Institute of Geological Sciences of the Jagiel-

lonian University. We thank Jadwiga Faber for her expert technical

assistance.

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