Time course of nodule development in the Discaria trinervis (Rhamnaceae) –Frankia symbiosis

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New Phytol. (1999), 141, 345–354

Time course of nodule development in the

Discaria trinervis (Rhamnaceae) – Frankia

symbiosis

CLAUDIO VALVERDE LUIS GABRIEL WALL*

Departamento de Ciencia y TecnologıUa, Universidad Nacional de Quilmes R. SaU enz Penh a180, Bernal (1876), Argentina

Received 15 May 1998; accepted 13 October 1998

The time course of initiation and development of root nodules was investigated in the South American actinorhizal

shrub Discaria trinervis (Rhamnaceae). A local strain of Frankia (BCU110501) which was isolated from D. trinervis

nodules, was used as inoculum. Inoculated seedlings were periodically studied under the light microscope after

clearing with aqueous NaClO. In parallel, semithin and ultrathin sections were analysed by light and electron

microscopy. Infection by Frankia BCU110501 involved intercellular penetration among epidermal and cortical

root cells. Nodule primordia were detected from 6 d after inoculation, while bacteria were progressing through

intercellular spaces of the outer layers of cortical cells. Invasion of host cells by the symbiont occurred 7–9 d after

inoculation, and hypertrophy of the primordium cells was associated with Frankia penetration. Root hairs were

not deformed during the early events of nodule formation. From 13 to 16 d after inoculation, the proximal cellular

zone of the primordia behaved differently from the other tissues after NaClO treatment and remained darkly

pigmented. At the same time, differentiation of Frankia vesicles started to occur inside already infected cells. By

16 d after inoculation, spherical vesicles of BCU110501 were homogeneously distributed in the host cells. These

vesicles were septate and surrounded by void space. Frankia spores or sporangia were not observed in the nodule

tissue. This study has clarified the mode of Frankia penetration in D. trinervis, one of the Rhamnaceae which also

includes Ceanothus. The events involved in infection, nodule induction, host-cell infection and vesicle

differentiation have been characterized and identified as time-segregated developmental processes in the ontogeny

of D. trinervis root nodules.

Key words: actinorhiza, Discaria trinervis, Frankia, intercellular penetration, nodule development.

The sequence of early cytological events in the

formation of N#-fixing root nodule symbioses has

been largely inferred from examination of older

nodule tissue. Few studies have focused on the time

course of nodule development in legumes (Turgeon

& Bauer, 1982; Calvert et al., 1984; Ridge & Rolfe,

1986; Sargent et al., 1987; Eskew et al., 1993).

Similarly, whereas the pathway of infection has been

elucidated in several actinorhizal plants, the time

course of nodule development has received little

attention (Burgess & Peterson, 1987; Berry & Sunell,

1990). All these studies investigated plants that were

infected through root hairs and there is no similar

information for intercellular invasion. An under-

*Author for correspondence (tel 54 11 43657100; fax 54 11

43657101; e-mail lgwall!unq.edu.ar).

standing of the time course of infection and nodule

development could be of great value in the study of

feed-back regulation responses by the plant and

establishing the stage of nodule development at

which signalling between plant and micro-organism

occurs. Furthermore, new information on different

actinorhizal plants is needed to make comparisons

with legumes, in order to analyse how these

symbiotic plants have evolved different or common

strategies for interaction with bacteria.

Actinorhizal plants are able to develop symbiotic

root nodules after infection of their roots by

filamentous gram-positive bacteria of the actino-

mycetic genus Frankia (Baker & Schwintzer, 1990).

Root nodules harbour a specialized living form of the

symbiont, vesicles, where atmospheric N#is reduced,

thus providing the host plant with combined nitro-

gen. In some genera (Casuarina, Allocasuarina)

vesicles are not differentiated and N#fixation occurs


346 C. Valverde and L. G. Wall

directly in the hyphal elements of the bacteria. Actin-

orhizal roots can be invaded by Frankia through one

of two known mechanisms; root hair infection, and

intercellular penetration (Berry & Sunell, 1990).

When the root hairs are the target for infection,

they become deformed and branched shortly after in-

oculation with the bacteria. Trapped Frankia hyphae

penetrate and grow basipetally inside the root hair

although they remain encapsulated by a host-derived

membrane and a thin cell wall. In parallel, a centre

of cell division, a prenodule, is generated in the outer

cortex just below the infected root hair. Frankia

continues to grow by cell-to-cell passage throughout

the prenodule and finally reaches the growing nodule

primordium which has already been induced in the

pericycle (Newcomb & Wood, 1987). When Frankia

invades an actinorhizal root intercellularly, root hairs

do not deform or branch, prenodules are not formed

and growth of Frankia is through intercellular spaces

among the cortical cells (Berry & Sunell, 1990).

Whether infection is initiated by root hair pen-

etration or by intercellular invasion, Frankia cells are

never in direct contact with the host-cell cytoplasm.

Discaria trinervis is a South American actinorhizal

shrub, that grows in western Argentina and eastern

Chile, between 31° and 48° S (Tortosa, 1983).

Discaria trinervis belongs to the Rhamnaceae family,

which also includes Ceanothus spp. and other South

American genera. In the field, the root system of D.

trinervis is naturally nodulated by Frankia (Chaia,

1997), but the mode of bacterial penetration into the

root however, is not known.

This paper describes a study of the infection

pathway of D. trinervis, and also a time-course

analysis of nodule formation in this actinorhizal

plant after inoculation with a pure culture of a

Frankia isolated from nodules of a field-grown D.

trinervis.

Plant material

Seeds of Discaria trinervis (Hooker et Arnot) Reiche

were kindly provided by E. Chaia (Universidad

Nacional del Comahue, Argentina). Mature fruits

were collected from the field in 1996 at Pampa de

Huenuleo (41° 10« S, 71° 12« W, Bariloche, Rio

Negro, Argentina). Seeds were separated from dry

fruits, exposed to the sun for 2 h and stored at

®20°C. No loss of germination occurred during

storage. Surface sterilization was performed by

scarification (3 min immersion in 10 M H#SO

%, with

occasional shaking by hand), and then exhaustively

rinsed with sterile distilled water (Chaia, 1997).

Seeds were blotted dry with filter paper, transferred

to vermiculite moistened with modified Evans sol-

ution, diluted to 1}10 strength (1}10 E) supple-

mented with 10 mg l−" of N as NH%NO

$(Huss-

Danell, 1978) and kept at 4°C for 5 d. Germination

and further plant growth were carried out in a

glasshouse, with mean max. and min. temperatures

of 27 and 20°C respectively, 65–95% r.h.

Frankia strain: culture and preparation of inoculum

Frankia strain BCU110501 was kindly provided by

E. Chaia (Universidad Nacional del Comahue,

Argentina). BCU110501 is a local isolate from D.

trinervis nodules, and is infective (Nod+) and effec-

tive (Fix+) in D. trinervis plants (Chaia, 1997). The

inoculum was prepared from bacteria grown for 4 wk

at 28°C in static culture in named minimal medium

BAP (Murry et al., 1984) supplemented with 0±055 M

glucose as the C source (Chaia, 1997). Cells were

sedimented by centrifugation (11000 g for 20 min),

washed with 1}10 E1 mg l−" N as NH%NO

$,

centrifuged again, and resuspended in a small

volume (2–3 ml) of 1}10 E1 mg l−" N. The cell

suspension was then homogenized by repeated

passage through needles of 0±8 and 0±5 mm gauge

(three times through each). Frankia biomass in the

homogenate was estimated as packed cell volume and

total protein was measured with the bicinchoninic

acid method (Nittayajarn & Baker, 1989) using BSA

(Sigma, St Louis, MO, USA) as standard.

Growth and inoculation of plants in pouches

Four seedlings at the cotyledonary stage (12–14 d

after the start of germination), were aseptically

transferred to growth pouches (Mega International,

Minneapolis, USA) moistened with 1}10 E1 mg

l−" N. Pouches were kept in the glasshouse through-

out the experiment. Incandescent lamps (400 W,

Osram, Germany) supplemented natural light to

give a photoperiod of 16 h light. After 4 wk of pouch

growth, the position of the root tip (RT) was marked

at inoculation on the pouch with a waterproof marker

pen. Then each seedling was inoculated by dripping

200 µl of the homogenate of Frankia BCU110501

(corresponding to 5 µl of packed cell volume and 20

µg of protein), from the root tip to the uppermost

zone of the root. Pouches were subsequently watered

with 1}10 E1 mg l−" N. When nodules were clearly

visible to the naked eye, the watering solution was

replaced by 1}10 E without N. Each treatment was

run in 10 pouches.

Growth and inoculation of plants in slides

Scarified seeds were tested for sterility by incubation

on solid nutrient broth at 28°C for 48 h, and

germinated under aseptic conditions as described

previously. Two seedlings at the cotyledonary stage,

were aseptically transferred to double length glass


Discaria trinervis nodule development 347

slides covered with a layer of 1}10 E1 mg l−" N in

10 g l−" agar gel (Sigma, St Louis, MO, USA). Slides

with seedlings were wrapped in aluminium foil and

placed vertically in a plastic container with 1}10

E1 mg l−" N, such that the bottom of the agar gel

was immersed to !1 cm in the solution (Wall &

Huss-Danell, 1997). After 7 d of growth, RT was

marked on the reverse of the slides and seedlings

were inoculated by dripping 50 µl of Frankia

BCU110501 homogenate (corresponding to 10 µl of

packed cell volume and 50 µg of protein), basipetally

from the root tip. Slides in the container were kept in

the glasshouse throughout the experiment. Nutrient

solution was renewed every 3–4 d. Inoculated D.

trinervis seedlings were analysed for root hair

deformation at 0, 3, 7 and 14 d after inoculation.

New slides (not previously unwrapped) were directly

inspected each time under the light microscope, and

&four seedlings were observed each time. Con-

taminated seedlings were not used for root hair

analysis.

Time course studies

Time course studies were undertaken on chemically

fixed D. trinervis root pieces removed from the

pouches at intervals after inoculation. Roots were

studied by each fo the following three methods: (1)

clarification with 0±74 M NaClO and staining with

10 g l−" methylene blue; (2) light microscopy of

semithin sections; and (3) electron microscopy of

ultrathin sections. For (1) no less than four seedling

roots were analysed whereas, for (2) and (3), the

biggest nodule primordia from four seedlings were

sampled for fixation.

Light microscopy of clarified roots

Roots were immersed in commercial 0±74 M NaClO

for 15–20 min, rinsed twice with distilled water and

stained with aqueous 10 g l−" methylene blue for 1

min (Truchet et al., 1989). Root pieces were placed

in a few drops of water on a slide, covered with a

cover glass and examined under a Zeiss2 Jenaval

light microscope.

Light microscopy of semithin sections

Selected root segments were fixed in glutaraldehyde

(25 g l−") in 45 mM potassium phosphate, pH 7±2, for

30 min at reduced pressure and then for at least 3–4

h at atmospheric pressure. Fixed root pieces were

post-fixed with osmium tetroxide (20 g l−") and

dehydrated with ethanol (50, 70, 80, 95 and 100%,

by volume). Dehydrated samples were embedded in

Epon-Araldite2 and polymerization was carried out

for 3 d at 70°C. Transverse or longitudinal sections

(1–1±5 µm) were mounted on glass slides, stained

with Methylene blue-Azur II, and examined as

described previously.

Transmission electron microscopy of ultrathin sections

Blocks with samples already used for semithin

sections were selected after light microscopy analy-

sis. Ultrathin sections were cut with glass knives,

stained with 20 g l−" uranyl acetate and 20 g l−" lead

citrate for 2–3 min each, and then analysed in a

JEOL}JEM 1200 EX II transmission electron

microscope.

Root structure and nodulation features

In both the growth pouch system and the slide

system, almost all seedlings of D. trinervis developed

a tap root with few lateral roots. Lateral roots started

to grow rapidly when the tap root reached the end of

the pouch. The tap root stele was diarch and

surrounded by a thick walled endodermis (Fig. 1a).

The cortex consisted of three to six layers of cells

with several intercellular spaces. Sparse cells con-

taining inclusions were seen in the cortex (Fig. 1a).

Flat epidermal cells formed the outermost layer of

root cells. Root hairs had a discontinuous dis-

tribution on the tap root, with hairy zones alternating

with hair-free zones.

Root nodules induced by Frankia BCU110501

usually developed in the tap root as single cylindrical

lobes (Fig. 1b) around RT. Bifurcation of the lobes

was seen in nodules older than 2 months. The

development of D. trinervis nodules was far from

synchronized and a range of developmental stages

was seen at each analysis. Thus, description of

nodule formation was based on the older primordia

of each sampling set.

Root hair deformation

No root hair deformation or branching was observed

72 h after inoculation, either above or below RT

(Fig. 1c) and root hairs remained undisturbed even

after nodule lobes were visible (see, e.g. Fig. 4a).

Nodules developed both in root zones bearing hairs

and in hairless regions. No unspecific root hair

deformation was detected even when seedlings were

contaminated with other bacteria during the ex-

periment (data not shown).

Nodule initiation and development

The normal structures of inoculated D. trinervis

roots remained unaltered between 2 and 5 d after

inoculation. Inoculation caused no root hair de-

formation in the pouch-grown seedlings.



Figure 1. Root structure and nodules of Discaria trinervis.(a) Transverse section an uninoculated root of D. trinervis,grown in the pouch system (for details see Materials and

Methods). Vascular bundles are diarch (protoxylematic

poles are indicated with small arrowheads). (b) Eight wk-

old root nodules of Discaria trinervis grown in the pouch

system and inoculated with a Frankia isolate, BCU110501.

(c) Root hairs of D. trinervis seedlings 72 h after

inoculation. Note that these show no deformation or

branching. Seedlings were grown using the slide technique

(for details see Materials and Methods).

Root nodule primordia were visible as masses of

small meristematic cells arising from the stele 6 d

after inoculation (Fig. 2a). These structures could be

distinguished from lateral root primordia, in which

the root cap primordium was clearly seen in the

outer cortex (Fig. 2b). Neither hypertrophy nor

hyperplasia of adjacent outer cortex cells was

observed. The root surface was not yet seriously

deformed by outgrowing primordia. Semithin trans-

verse sections showed nodule primordia emerging

from the pericycle from one of the two pro-

toxylematic poles of the diarch stele, through the

endodermis. At this stage, primordia were composed

of small cells (Fig. 2c,d). There was no prenodule

formation in the outer cortex. When a nodule

primordium developed in a root-hair bearing zone,

root hairs were not deformed and did not show

abnormal cellular contents. Infection sites were

inferred from a dark staining of the intercellular

spaces between the epidermis and the next cell layer

(Fig. 2d). Frankia hyphae were also detected in these

infection sites (Fig. 2e) but were more clearly

identified in ultrathin TEM sections, where they

were visible embedded in electron-dense material in

the intercellular spaces of the hypodermal root cells

(Fig. 2f). Nodule primordia cells were not invaded

by hyphae of BCU110501 at this time.

Between 9 and 11 d after inoculation, nodule

primordia began to protrude through the epidermis

and were visible on the root surface (Fig. 3a).

Transverse sections showed that nodular proto-

peridermis and vascular bundles had begun to

develop (Fig. 3b). At the ultramicroscopic level,

Frankia hyphae were seen in the intercellular space

between primordia cortical cells, from where they

invaded a host cell in the basal zone of the primordia

(Fig. 3c). During this process, Frankia hyphae did

not disrupt the host plasma membrane, and hyphae

became surrounded by a matrix, probably of host

origin (Fig. 3c). Light microscopy revealed that host

cells in the basal zone of the primordia were

hypertrophic and showed intracellular hyphae (Fig.

3d). Uninfected cells ramined normal-sized. Mul-

tiple infection sites and pathways were usually

identified for each primordium. When two nodule

primordia were detected in the same root transverse

section, they arose opposite the protoxylematic poles

(Fig. 3b). Septate Frankia vesicles were occasionally

seen in infected host cells from the basal zone of a

nodule primordium (Fig. 3d,e).

A striking differential staining of the proximal

zone of the nodule lobes was observed, after root

clarification, from 13 d after inoculation (Fig. 4a). At

the same time, septate vesicles were seen filling the

hypertrophied host cells in the basal portion of the

lobes (Fig. 4d,e). Spherical vesicles were uniformly

distributed within infected host cells. Vesicles were

surrounded by a void space, which is an artefact

induced by specimen preparation, typically seen in

histological sections of Alnus and several other

actinorhizal nodules. This void space corresponds to

the multi-layered lipid envelop of vesicles (Huss-

Danell, 1997). At this stage of nodule development,

a zonation of the nodules was evident (Fig. 4b). This

consisted of a nodule periderm wrapping and

protecting nodule inner tissues; an apical meristem

leading nodule growth and leaving behind a zone of

cortical cell maturation and differentiation; an

infection zone with hypertrophied cells bearing



Figure 2. Root nodule primordia of Discaria trinervis 6 d after inoculation. (a) Root segment clarified and

stained with Methylene blue (CMB) showing a mass of meristematic cells (*) emerging from below the

endodermis (En). No hypertrophy or hyperplasia of the outer cortical cells was observed close to the nodule

primordia. C, cortex. (b) Non-inoculated CMB-treated root bearing a lateral root primordium. (c) Transverse

section of a nodule primordium. Meristematic cells (*) are shown, arising from the pericycle, opposite to one

of the protoxylematic poles (small arrowhead). The endodermis seems to be disrupted by the growing

primordium (large arrowheads). (d) Enclosed area in (c) where the site of infection was inferred because of

staining of intercellular spaces (arrowhead). *, nodule primordium. (e) Frankia hyphae (small arrowheads) have

colonized the root epidermis of D. trinervis and penetrated the root between epidermal (Ep) and hypodermal

(Hy) cells, through the intercellular spaces, which are filled with dark-stained material (large arrowheads). (f)

TEM of an infection site where transversally sectioned Frankia hyphae (H) are embedded in electron-dense

material (m), located in the space between cortical root cells (C). cw, cell wall.

Frankia hyphae (Fig. 4c) and normal sized un-

infected cells ; finally a proximal zone of hypertro-

phied and infected cells bearing Frankia vesicles in

contact with uninfected cells (Fig. 4d). Vascular

bundles appeared to be fully developed. At the

ultrastructural level, hyphae, and also numerous

vesicles (between 2±5–3±7 µm diameter) containing a

few septa, were present in infected host cells (Fig.

4e). All of the vesicles were surrounded with a void

space.

Several days later, nodules did not differ from

those described for 16-d-old lobes, except for their

bigger size. Sporangia or spores were not detected by

light microscopy of semithin transverse sections.



3a 3b

3c

3d

3e

*

*

100 µm

200 µm

ic

H

H

1 µm

20 µm

ic

uc

uc

ic

H

V

vs

1 µm

Figure 3. Root nodule primordia of Discaria trinervis, 9 d after inoculation. (a) The nodule primordium (*) has

disrupted the epidermis (large arrowheads). Vascular bundles have begun to develop (small arrowhead). (b)

Transverse section of a root bearing two nodule primordia (*), each emerging opposite to one of the

protoxylematic poles. Protoperiderm (large arrowhead) and vascular bundles (small arrowhead) have begun to

develop. Multiple infection sites are seen (arrows). (c) TEM showing the penetration of Frankia hyphae (H)



The Frankia isolate BCU110501 indues develop-

ment of N#-fixing root nodules in the native South

American actinorhizal plant, D. trinervis (Chaia,

1997). Our work shows that roots of D. trinervis are

infected by intercellular penetration (Miller & Baker,

1985b), as occurs in the genus Ceanothus (Liu &

Berry, 1991), which also belongs to the family

Rhamnaceae. Previous work on other actinorhizal

plants indicates that the pathway of infection is

determined by the host plant (Miller & Baker,

1986; reviewed by Huss-Danell, 1997). The cytology

and ultrastructure of nodules of D. trinervis are

similar to those reported for nodules of Discaria

toumatou (Newcomb & Pankhurst, 1982). However,

our observations over an extended time period, have

enabled us to identify the sequence of various

individual developmental stages within the whole

process of nodule formation in the D. trinervis–

BCU110501 symbiosis. The first stage of root

infection and nodule induction takes c. 6 d from the

time at which the two partners make contact at

inoculation (Fig. 2). After Frankia hyphae reach the

root surface, they adhere to the epidermis and enter

the root by penetration, first between epidermal cells

and then between outer cortical cells. The root hairs

do not express any response during these early

interactions with the bacteria, and they remain

undeformed throughout the process of nodulation

(Fig. 1c). This observation is consistent with the

observed mode of root infection. Root hairs are not

involved in the initiation of nodulation in other

actinorhizal plants where intercellular invasion of

the symbiont occurs (Berry & Sunell, 1990). Never-

theless, root cells of D. trinervis sense the presence of

the invading bacteria and secrete locally an electron-

dense material into the intercellular spaces. As has

also been reported for Elaeagnus angustifolia (Miller

& Baker, 1985a), this extracelular material was

apparently not digested by the bacteria (Fig. 2f ).

Frankia BCU110501 can progress through this

material without leaving a clear zone around it. At

the time when hyphae are moving between outer

cortical root cells, development of the nodule

primordium is induced within the pericycle, imply-

ing that some kind of signal transduction occurs

between symbionts. Masses of small cells with dense

cytoplasm emerge through the endodermis from

protoxylematic poles of the diarch stele.

The hyphae of Frankia reaches the basal zone of

into a nodule cortical cell (ic), from the intercellular space. Intracellular Frankia hyphae are surrounded by a

capsule (arrowhead). (d) Longitudinal section of a young nodule where basal cortical cells (ic) of the

primordium are infected with Frankia BCU110501 hyphae (arrows). Note the hypertrophy of the infected

cells. Uninfected cels (uc) remained normal-sized. Frankia hyphae can be seen between cortical cells

(arrowheads). (e) Frankia vesicles (V) were occasionally detected in a TEM of an infected host cell. The vesicle

was septate (small arrowhead) and surrounded by a void space (vs). H, hyphae.

the primordium and penetrates the differentiated and

mature host cells of the nodule parenchyma (Fig. 3).

This process appears to induce an increase in the size

of the host cell, whereas uninfected cells retain their

normal dimensions. Within the nodule cells, Frankia

is embedded in a matrix and always surrounded by a

membrane of plant origin. The central vascular

bundle of the young nodule develops, as does the

protoperiderm, which will constitute the outermost

cell layer of the nodule lobe. Vesicle differentiation

in Frankia BCU110501 seems to occur soon after

host-cell invasion, since a septate vesicle was seen

immediately after hyphae had penetrated into a

nodule basal cell.

Vesicle differentiation results in numerous spheri-

cal and septate vesicles, uniformly distributed

throughout the cytoplasma of the host cells (Fig. 4).

Concomitantly, the basal zone of the lobes becomes

opaque after oxidation by hypochlorite. Nitrogen

fixation is probably occurring at this stage since

acetylene reduction activity was detected 1 wk later-

(data not shown). At 2 wk after inoculation, the first

nodules are easily identified as single cylindrical lobes

in the root system. Nodule emergence over the same

time scale was reported in Sepherdia argentea seed-

lings inoculated with Frankia HFPGpI1 (Racette

& Torrey, 1989). At the microscopic level, all tissues

described in other actinorhizal root nodules, such as

vascular tissue, nodule apical meristem, periderm

and infected cortical tissue (Baker & Schwintzer,

1990), have been developed at this stage. Further

nodule growth results in increased size and no

change in histology. This D. trinervis–BCU110501

symbiosis is phenotypically Spore− under laboratory

conditions, as no sporangia or spores were detected

in the nodule. This observation agrees with field

studies of rhamnaceaen nodules (Chaia, 1993, 1997).

Recent molecular phylogenetic studies, based on

the gene sequence encoding the large subunit of

chloroplastic rubisco (rbcL) of actinorhizal and non-

actinorhizal relatives, together with new symbiotic

properties (Swensen, 1996; Swensen & Mullin,

1997) have hypothesized four branches (clades) of

independent evolution of actinorhizal plants. Our

findings on the mode of infection in D. trinervis

confirm the assumption that phylogenetically

clustered actinorhizal genera placed within clade II,

which grouped Discaria close to Trevoa, Colletia,

Ceanothus, HippophaeX , Elaeagnus and Shepherdia,

share the mode of bacterial penetration into the host

roots, but also other anatomical features of the



Figure 4. Root nodule lobes of Discaria trinervis, 16 d after inoculation. (a) A lobe showed differential staining

in its basal zone after root segment was clarified and stained with methylene blue (CMB) (arrowhead). (b)

Longitudinal section of a D. trinervis root bearing three nodule lobes. Zonation of the lobe can be seen at this

stage: periderm (large arrowhead), vascular bundles (small arrowhead), apical meristem (*), meristem derived

cortical cells (mc), cortical cells infected with Frankia hyphae (ic), cortical cells infected with Frankia vesicles

and hyphae (vc). (c) Magnified enclosed area A from (b), showing details of the transition limit between zones

of newly mature cortical cells (mc) and infected cortical cells (ic). (d) Magnified enclosed area B from (b),showing details of the lobe zone where Frankia vesicles are differentiated (small arrowheads). Vesicles were

uniformly spread into the hypertrophied host cell. (e) TEM showing Frankia vesicles (V) and hyphae (H) in

nodule cortical cells. Vesicle diameter ranged from c. 2±5 and 3±7 µm. Usually, one to three septa were observed

(small arrowheads) in each vesicle.



N2 fixation (nif)

Pigmentation of nodule basal cells

Frankia vesicle differentiation (ved)

Host cell invasion by Frankia (hin)

Nodule initiation in pericycle (noi)

Root intercellular infectionby Frankia (inf)

Preinfectionevents

Root adhesion (roa)

Root colonization (roc)

No root hair deformation (rhd–)

Inoculation

0 3 6 9 13 16

Time (days)

?

?

Figure 5. Time course of developmental events in the ontogeny of Discaria trinervis nodules. Abbreviations

of the stages (between parentheses) are according to recently proposed terminology (Akkermans & Hirsch,

1997). Time scale is linear and ticks correspond to experimental observations (Figs 1–4). Question marks (?)

denote uncertainty in the time scale of certain processes.

infected cells and vesicle structure (Swensen &

Mullin, 1997).

Finally, we propose a model summarizing our

description of the nodulation process in D. trinervis

(Fig. 5). This model also includes some preliminary

results on N#

fixation and the regulation of nodule

development. These are indicated as question marks

at the end of the ‘ inf ’ and ‘noi ’ processes. This

model might be useful in interpreting further studies

on the mechanisms of induction of nodule genes

(Pawlowski & Bisseling, 1996) and on the regulation

of nodulation (Wall & Huss-Danell, 1997).

We are greatly indebted to E. Chaia (Universidad Nacional

del Comahue, Argentina) for providing D. trinervis seeds

and the Frankia BCU110501 isolate. We thank the

Instituto de Bioquı!mica y Biologı!a Molecular (IBBM,

U.N.L.P., Argentina) which kindly gave us the oppor-

tunity to perform a greater part of this work. Financial

support was obtained through a grant from Universidad

Nacional del Comahue and Universidad Nacional de

Quilmes (Argentina). C. Gonzalez is thanked for her

technical assistance with sample processing for micro-

scopy. Part of this work was performed at the Swedish

University of Agricultural Sciences, Umea/ , Sweden, with

financial support from The Swedish Foundation for

International Cooperation in Research and Higher Edu-

cation (through a grant to Dr K. Huss-Danell). We also

thank Dr K. Huss-Danell for valuable comments on an

early draft of the typescript, and R. Ariet for her review of

the spelling. C.V. holds a fellowship from Consejo

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Documents

Time course of nodule development in the Discaria trinervis (Rhamnaceae) –Frankia symbiosis