From embryo sac to oil and protein bodies: embryo development in the model legume Medicago truncatula


  • Xin-Ding Wang,

    1. Australian Research Council Centre of Excellence for Integrative Legume Research, School of Environmental and Life Sciences
    Search for more papers by this author
    • These authors contributed equally to this work.

  • Youhong Song,

    1. Australian Research Council Centre of Excellence for Integrative Legume Research, School of Environmental and Life Sciences
    Search for more papers by this author
    • These authors contributed equally to this work.

  • Michael B. Sheahan,

    1. Australian Research Council Centre of Excellence for Integrative Legume Research, School of Environmental and Life Sciences
    Search for more papers by this author
  • Manohar L. Garg,

    1. School of Biomedical Sciences and Pharmacy, The University of Newcastle, Newcastle, 2308, Australia
    Search for more papers by this author
  • Ray J. Rose

    1. Australian Research Council Centre of Excellence for Integrative Legume Research, School of Environmental and Life Sciences
    Search for more papers by this author

Author for correspondence:
Ray J. Rose
Tel: +61 2 49215711


  • The cell and developmental biology of zygotic embryogenesis in the model legume Medicago truncatula has received little attention. We studied M. truncatula embryogenesis from embryo sac until cotyledon maturation, including oil and protein body biogenesis.
  • We characterized embryo development using light and electron microscopy, measurement of protein and lipid fatty acid accumulation and by profiling the expression of key seed storage genes.
  • Embryo sac development in M. truncatula is of the Polygonum type. A distinctive multicellular hypophysis and suspensor develops before the globular stage and by the early cotyledon stage, the procambium connects the developing apical meristems. In the storage parenchyma of cotyledons, ovoid oil bodies surround protein bodies and the plasma membrane. Four major lipid fatty acids accumulate as cotyledons develop, paralleling the expression of OLEOSIN and the storage protein genes, VICILIN and LEGUMIN.
  • Zygotic embryogenesis in M. truncatula features the development of a distinctive multicellular hypophysis and an endopolyploid suspensor with basal transfer cell. A clear procambial connection between the apical meristems is evident and there is a characteristic arrangement of oil bodies in the cotyledons and radicle. Our data help link embryogenesis to the genetic regulation of oil and protein body biogenesis in legume seed.


Extensive examination of embryo morphogenesis since the 19th century has led to a generalized paradigm of the major morphological events involved in plant embryogenesis (Raghavan, 1986; Goldberg et al., 1994). Present-day research is focused on understanding the genetic regulation of embryogenesis (Laux et al., 2004; De Smet et al., 2010), which has primarily been facilitated by the model plant Arabidopsis (Arabidopsis thaliana; Meinke, 1991; Goldberg et al., 1994; De Smet et al., 2010). Detailed studies of embryo morphogenesis in Arabidopsis (Mansfield & Briarty, 1991; Mansfield et al., 1991), including fine structure, show that embryogenesis in this plant is essentially similar to the classic model for embryogenesis, Capsella bursa-pastoris.

Grain legumes are of significance for food, nutrition and fuel, and Medicago truncatula, as a model legume (Rose, 2008), is being used to study the genetic regulation of somatic embryogenesis (Rose et al., 2010) and seed development (Thompson et al., 2009). There have been detailed transcriptomic and proteomic analyses of M. truncatula seed development (Gallardo et al., 2007), providing a legume perspective to complement Arabidopsis research. Although line drawings representing a few stages of embryogenesis were reported for Medicago sativa (Martin, 1914) and the structural development of the M. truncatula pod wall and seed coat has been characterized (Wang & Grusak, 2005), there is no detailed study on the morphological development of the M. truncatula embryo. Such morphological data would provide a suitable base for investigations into somatic embryogenesis and seed development in M. truncatula and provide useful comparative information with soybean (Glycine max), a major crop legume whose embryology (Kennell & Horner, 1985; Chamberlin et al., 1994) and seed molecular biology have been studied (Le et al., 2007).

In this investigation, we provide histological data that illustrate development of the embryo, from embryo sac to mature embryo (including its meristems). We also show details of the morphology and distribution of oil and protein bodies in cotyledons and the radicle. Oil bodies have received little attention in legumes, but through extensive serial sectioning, we reveal new details of their cellular organization. We also investigated the accumulation of storage products in the developing seed. We discuss these data in the context of the developmental biology of legume embryogenesis and highlight the special features of M. truncatula embryogenesis.

Materials and Methods

Plant material

Flowers and developing pods were obtained from glasshouse-grown plants of M. truncatula Gaertn cv Jemalong. The glasshouse had a 14 h photoperiod with a 200 μmol m−2 s−1 lighting intensity and 23 : 19°C day : night temperature regime.

Histology of thin sections

Tissue fixation in 4% (w : v) glutaraldehyde and 2% (w : v) paraformaldehyde in 100 mM sodium cacodylate buffer (pH 7.2), dehydration and infiltration in LR White resin was as previously described (Rose et al., 2006). The sectioning of LR White-embedded tissue, the staining of 1 μm sections in toluidine blue and Azur II and light microscopy were also as previously described (Rose et al., 2006). Oil bodies in tissue sections were stained by Nile red (0.02 mg ml−1 in 50% acetone) for 5 min followed by washing in deionized water. Fluorescence imaging was performed using a 50 W Hg lamp, with excitation at 510–560 nm and emission at 590 nm with a Zeiss Axiophot microscope.

Electron microscopy

Tissue (dry seed was first imbibed in water for 5 d at 4°C) was fixed using 4% (w : v) glutaraldehyde and 1.5% (w : v) paraformaldehyde in 25 mM phosphate buffer (pH 7) at 4°C for 4 h, with at least one change of buffer. Tissue was transferred to phosphate buffer, washed once and incubated overnight at 4°C. The tissue was then postfixed in 1% aqueous osmium tetroxide overnight at 4°C and washed twice with distilled water chilled to 4°C. Dehydration was carried out at 4°C with a graded series of 10% (v : v) ethanol with 45 min incubation times. The tissue in 100% ethanol was then left overnight at 4°C. Infiltration into LR White resin was carried out at room temperature on a rotator in a graded series of 20% LR White going from 100% ethanol to 100% LR White with intervals of at least 2 h. There were at least three changes of resin. For the ovules, the resin was infiltrated over 24 h while for the seed it was over at least 7 d. Polymerization was carried out in capsules at 60°C for at least 24 h. Ultrathin sections were cut with a diamond knife and transferred to 200-mesh grids. The sections were stained with uranyl acetate and lead citrate and viewed using a JEOL JEM 1200-EXII electron microscope (JEOL Ltd., Tokyo, Japan). Oil and protein body morphology was quantified using ImageJ (; Abramoff et al., 2004).

Lipid extraction

Lipid extraction was based on the method of Folch et al. (1957). Whole seed was homogenized with chloroform : methanol (2 : 1, v : v) in a final volume 20 times the volume of the sample. After dispersion, the mixture was agitated for 15–20 min in an orbital shaker at room temperature. The homogenate was centrifuged to recover the liquid phase. The solvent was washed with 0.2 volumes of a 0.9% NaCl solution, vortexed and then centrifuged at 3000 g. The upper phase was discarded and the lower phase containing the lipids evaporated under a nitrogen stream.

Fatty acid analysis by gas chromatography

The total fatty acid concentration was analysed by direct transesterification of lipids using a gas chromatograph (GC) as previously described (Koppers et al., 2010). A 4 : 1 (v : v) methanol : toluene mixture (2 ml) containing C21 : 0 or C19 : 0 fatty acids (20 μg ml−1) as internal standards was added to the sample (100 μl). Acetyl chloride (200 μl) was then added to the samples while vortexing, before heating at 100°C for 1 h. The tubes were cooled in water (5 min) and 6% K2CO3 (5 ml) was added before centrifuging (3000 g, 5 min, 4°C). The upper toluene phase was collected and stored in a GC vial at −20°C for analysis.

Methylated total fatty acid samples were analysed by GC using a fixed carbon-silica column (30 m × 0.25 mm, DB-225, J and W Scientific, Folsom, CA, USA). The GC was equipped with a flame ionization detector, autosampler and autodetector. Injector and detector ports were set at 250°C. The oven temperature was 170°C for 2 min, and then increased at 10°C min−1 up to 190°C where it remained stationary for 1 min. The temperature was then increased at 3°C min−1 up to 220°C, which was maintained for a total run time of 30 min per sample. A split ratio of 10 : 1 and an injection volume of 3 μl were used. A known fatty acid mixture was used to identify peaks according to retention time and their concentration was determined using a Hewlett Packard 6890 Series GC with Chemstations Version A. 04.02 (Hewlett Packard, Palo Alto, CA, USA).

Determination of total protein and seed water content

Proteins were quantified essentially as described by Baud et al. (2002). Four seeds were ground in 500 μl of extraction buffer (50 mM HEPES, 5 mM MgCl2, 5 mM dithiothreitol, 1 mM EDTA, 3 mg ml−1 polyvinylpyrrolidone and 10% (v/v) ethylene glycol (pH 7.5)) and then sonicated (Branson Sonifier 450, Branson Ultrasonics, Danbury, CT, USA; 3 × 15 s at 25% duty cycle and output level 2). The extracts were centrifuged (16 000 g, 12 min, 4°C) and the supernatant was used to measure soluble protein by the Bradford Assay (Bradford, 1976). The remaining insoluble protein was extracted by grinding in 500 μl of 1 M NaOH and vortexing for 30 min. The extract was then centrifuged and the protein quantified as described earlier. A BSA standard curve was used to derive actual protein concentrations. Seed water content was calculated from the difference in seed FW and DW. DW was determined after incubation of seeds at 80°C for 2 d.

Identification of oleosin, vicilin and legumin sequences

Protein and transcript sequences were obtained from the Phytozome ( and Genbank ( databases by keyword and BLAST searches, using tentative consensus sequences obtained from the M. truncatula gene index (


Full-length amino acid sequences were aligned using ClustalX 2.0.10 (Larkin et al., 2007). Phylograms were constructed from aligned sequences using the protein maximum likelihood (proml) program in PHYLIP (PHYlogeny Inference Package version 3.69; Felsenstein, 1989) and drawn with Dendroscope (Huson et al., 2007; see Supporting Information, Table S1, for sequences used in construction of the phylogenetic tree).

Gene expression analysis

RNA was isolated from ovules and developing seed using the RNAqueous-4PCR kit (Ambion) and DNase-treated according to the manufacturer’s instructions. Synthesis of cDNA was performed with a SuperScript III first-strand synthesis system (Invitrogen) using 2 μg of total RNA and oligo(dT) primers. The cDNA was diluted 1 : 25 for quantitative real-time polymerase chain reactions (qPCR). All qPCR reactions were prepared using a CAS1200 robot (Qiagen) and run on a Rotor-Gene Q (Qiagen). Primers were designed using Primer3 (Rozen & Skaletsky, 2000; and used to amplify specific oleosins. The legumin and vicilin primers targeted a conserved region common to all members of the respective families and thus have the potential to amplify any of the members within these gene families (Table S2 for primer sequences). Reactions were performed in triplicate (15 μl sample volume) using Platinum Taq PCR polymerase and 2 μM SYTO9 fluorescent dye (Invitrogen). PCR cycling conditions were 94°C for 2 min, followed by 40 cycles of 94°C for 15 s, 60°C for 30 s and 72°C for 30 s.

Disassociation analysis was performed in each run to verify the amplification of a specific product. Gene expression was normalized to the expression of glyceraldehyde 3-phosphate dehydrogenase. PCR efficiency of each run was calculated using the LinRegPCR program (; Ramakers et al., 2003). Relative expression was calculated using the Pfaffl method (Pfaffl, 2001). Results shown are means ± SE of three biological repeats.


The anthers, ovary, ovules and megaspore

Longitudinal sectioning of M. truncatula floral buds permitted visualization of the anthers, stigma, style and ovary, and highlighted a close association between anthers and the stigma (Fig. 1a). The characteristic legume ovary, which forms the pod, had 11 ovules connected to the ovary wall in this section. An image of a single ovule (Fig. 1b) shows the surviving haploid megaspore, with prominent nucleus, that will develop into the embryo sac. The megaspore is surrounded by the nucellus, which is in turn surrounded by the integuments (Fig. 1b).

Figure 1.

The flower – ovary, ovules and megaspore. (a) Stigma, anther, petal and ovary indicated. Note the close proximity of the anther to the stigma. (b) The prominent surviving haploid megaspore (arrow), with its prominent nucleus, will develop into the embryo sac. The megaspore is surrounded by the nucellus (N), which in turn is surrounded by the integuments (I). The inset shows an enlarged view of the megaspore. Bars, 200 μm (a); 20 μm (b); 10 μm (b, inset).

The embryo sac

Using serial sectioning (see Fig. S1), we found the fully developed embryo sac to be of the classical Polygonum-type (Kennell & Horner, 1985; Mansfield et al., 1991), with the surviving haploid megaspore undergoing three mitotic divisions to form the eight nuclei and seven cells of the embryo sac. A single serial section through an embryo sac is shown (Fig. 2a). Sectioning through the entire embryo sac revealed that the eight nuclei could be assigned to the egg cell, two synergid cells, three antipodal cells and the central cell, which contained two polar nuclei (Figs 2b, S1).

Figure 2.

The embryo sac. Serial sectioning (see Supporting Information, Fig. S1) shows the embryo sac contains seven cells, including the egg cell and eight nuclei. (a) Single serial section (section 21 in Fig. S1) showing two polar nuclei in the central cell (CC) and an antipodal nucleus (A). (b) Diagrammatic representation of how the transverse sectioning plane from the micropylar end relates to identification of the eight nuclei of the embryo sac. Bar, 10 μm.

Fertilization and first cell divisions

The zygote has a characteristic large vacuole at the micropylar end (Fig. 3a). The first asymmetric division of the zygote clearly demarcates the larger and more vacuolate basal cell from the densely cytoplasmic apical cell destined to form most of the embryo (Fig. 3b). The apical cell has a ‘dome’ shape representative of the future embryo proper, whereas the basal cell is elongated and contains many small vacuoles. After the second anticlinal division, the size of basal and apical cells becomes similar, with a gradient of vacuolation apparent along the apical–basal axis (Fig. 3c). After further periclinal and anticlinal divisions, the apical cell develops into the early proembryo and the basal cells into a clearly demarcated hypophysis and suspensor (Fig. 4a,b). A clear gradient in vacuolation exists along the apical–basal axis of the developing embryo, with vacuolation minimal in the proembryo and maximal in the most basal cell of the suspensor (Fig. 4a). Plastids containing starch surrounded by a densely stained stroma were also prominent but there were fewer of them in the cells of the proembryo (Fig. 4a). It is clear from Fig. 4(a–b) that the proliferation and enlargement of the suspensor cells and a clearly distinguishable hypophysis (with characteristic cell size, vacuolation and shape) are dominant features of embryogenesis in M. truncatula. In Fig. 4(b), the densely cytoplasmic cells of the embryo and hypophysis are in clear contrast to the vacuolated cells of the adjoining suspensor.

Figure 3.

The zygote, first asymmetric division and second anticlinal division. (a) The zygote (arrow) with prominent nucleus and nucleolus and large vacuole (V) in the basal half of the cell. A synergid cell (arrow) is adjacent to the zygote. Remnants of a sperm cell (arrow) are present. (b) Electron micrograph of the first, asymmetric division that gives rise to cells forming the embryo (E) and suspensor (S). (c) Electron micrograph of embryo after second anticlinal division. Bars, 10 μm (a); 5 μm (b, c).

Figure 4.

Embryo development up to the heart stage. (a) Embryo development at 2 d after anthesis (DAA). Developing embryo (E), hypophysis (H) and suspensor (S). (b) Embryo development at 3 DAA. (c) Globular stage embryo at 5 DAA, with prominent hypophysis that will give rise to the root apical meristem. (d) Heart stage embryo at 6 DAA with an extensive suspensor. Bars, 10 μm (a, b); 20 μm (c); 50 μm (d).

Globular stage to cotyledon stage of embryogenesis

By the globular stage of embryo development (Fig. 4c), there is a clear demarcation between the embryo (integrated shoot and root apical regions) and suspensor, with suspensor and embryo cells each possessing a distinct morphology and intracellular organization. The large suspensor cells possess large nuclei and vacuoles and also many dense-staining plastids with small amounts of starch. By the heart stage (Fig. 4d) the cells of the embryo proper are more vacuolate and a long, mostly three-cell-wide, suspensor with large, elongated cells in close contact with the endosperm and seed coat has developed. The elongated suspensor now occupies a morphological niche in the hilum region of the seed coat (Fig. 4d). At the early cotyledon stage, wall ingrowths can be observed in the basal suspensor cell where it abuts the seed coat (Fig. 5a). There are also numerous plastids with small amounts of starch and a densely stained stroma in cells of the suspensor (Fig. 5b). These plastids contrasted the characteristic amyloplasts present within inner cells of the seed coat (Fig. 5c).Intriguingly, the vacuole of suspensor cells appears to have fragmented at this stage of development. Finally, suspensor cell nuclei are much larger (on average, 1.6 ± 0.2-fold larger diameter) than nuclei in cells of the embryo proper (Fig. 5d,e), consistent with endopolyploidy in the suspensor cells.

Figure 5.

The suspensor cells at 8 d after anthesis (DAA). (a) Large basal cell of the suspensor with prominent wall ingrowths (arrows) and plastids (Pl). SC, seed coat. (b) Basal suspensor cells exhibit a large nucleus (N) and nucleolus (Nu), plastids with dense stroma and small amounts of starch. The outlined region is shown in (a). (c) Cells of the seed coat adjacent to the endosperm have wall ingrowths (arrows) and plastids that are typical amyloplasts (A) packed with starch. (d) Large cells of the suspensor with large nuclei (N) and plastids (Pl) with dense stroma and varying amounts of starch. Note the fragmented appearance of the vacuole. (e) Smaller cells of the embryonic radicle have smaller nuclei (N) than the suspensor cells. Bars, 10 μm; (b) is the same scale as (c); (d) is the same scale as (e).

As the cotyledons develop, vacuolation of their cells increases and the suspensor slowly degenerates (Fig. 6a,b). Moreover, by the cotyledonary stage of development, an extensive procambium connects the nascent shoot and root apical meristems (RAMs) (Fig. 6b). With further embryo development, the shoot and RAMs attain a cellular organization characteristic of these primary meristems (Fig. 6c,d). In the case of the shoot apical meristem, the characteristic dome morphology of this meristem has developed, with several densely cytoplasmic cells in the region of the organizing centre apparent at higher resolution (Fig. S2). Similarly, in suitable sections, the quiescent centre and stem cells of the open M. truncatula root meristem can be visualized in a region consistent with our previous in situ hybridization assays for MtWOX5 (Chen et al., 2009; Fig. S3). In addition, the procambium at this stage extends into the cotyledons (Fig. 6d). In a mature embryo, extensive development of the cotyledons and prominent radicle is evident (Fig. 6e).

Figure 6.

Cotyledon development and embryo maturation. (a) Embryonic cotyledons (C) have formed at 8 d after anthesis (DAA) in the developing seed. Endosperm (En) is breaking down as it is used as a nutrient source. The suspensor (S) is still present but appears to be degrading. The hilum region of the seed coat is indicated by *. (b) Embryo shown in (a) at a higher resolution. The three primary meristems – shoot apical meristem (SAM), root apical meristem (RAM) and procambium (PC) – are developing. (c) Further seed development at 11 DAA. The endosperm has been further degraded and the cotyledons (C) have become more prominent. The hilum region of the seed coat is indicated by *. (d) Embryo in (c) at a higher resolution. There is further development of the primary meristems and the SAM has developed a characteristic dome shape. (e) An embryo near maturity at 28 DAA. The prominent features are the radicle (R) and storage cotyledons (C). Bars, 200 μm (a, c); 100 μm (b, d); 500 μm (e).

Protein and oil bodies in the cotyledons and radicle

Examining the storage parenchyma of cotyledons stained with toluidine blue and Azur II by high-resolution light microscopy revealed cells packed with darkly stained protein bodies (Fig. 7a). In these cells, a bright border of small globular-like structures surrounded protein bodies and the cell periphery (Fig. 7a). Staining cotyledons for neutral lipid with Nile red (Siloto et al., 2006) also strongly labelled the margin of protein bodies (Fig. 7b) and cell periphery (Fig. 7a). However, we were unable to fully resolve oil body structures from these light micrographs (Fig. 7a). Using electron microscopy revealed that closely packed yet discrete oil bodies always surrounded the densely staining protein bodies (Fig. 8). At lower magnification, the intracellular organization in the storage parenchyma of cotyledons appeared generally uniform, with protein bodies the most prominent feature in these cells (Fig. 8a). At higher resolution, it was clear that relatively few oil bodies were free in the cytoplasm (Fig. 8b). Moreover, and intriguingly, oil bodies not only bounded protein bodies (Fig. 8c), and to a much lesser extent amyloplasts (Fig. 8b), but also lined the plasma membrane (Fig. 8b). Though protein and oil are the dominant storage products, some amyloplasts are present (Fig. 8b). Our observations are consistent with Djemel et al. (2005) in that there is more storage of starch in the seed coat (Fig. 5c) and embryo in early development. Protein bodies in the cotyledons are typically large and have a morphology that varies from circular to kidney-, lens- and dumbbell-shaped (Fig. 8a). Quantitative image analysis indicated protein bodies have an average plan area of 7.2 ± 2.2 μm2 (mean ± SD, = 50 cells), with an average aspect ratio (the quotient of the major and minor axes of an ellipse fitted to the protein body) of 1.5 ± 0.2. By contrast, oil bodies are small and uniformly ovoid (Fig. 8c), with an average plan area of 0.035 ± 0.010 μm2 (mean ± SD, = 185 oil bodies), a diameter of 0.23 ± 0.03 μm and an aspect ratio of 1.4 ± 0.2. In cells of the radicle, oil bodies are also ovoid (with an aspect ratio of 1.3 ± 0.1) and of a similar average size, but exhibited greater size variation (Fig. 9) with an average plan area of 0.032 ± 0.031 μm2 (mean ± SD, = 686 oil bodies). Strikingly, most (76 ± 4%; mean ± SD) oil bodies in these cells preferentially lined the plasma membrane.

Figure 7.

Brightfield and fluorescence images of storage parenchyma cells in mature cotyledons. (a) Light micrograph of a mature cotyledon cell stained with toluidine blue and Azur II. Protein bodies appear dark and are surrounded by a bright bead-like perimeter – the oil bodies. (b) Light micrograph of a mature cotyledon cell stained with Nile red. Red fluorescent lipid is more intense around the dark protein body perimeter and the cell periphery. Protein bodies (P); cell wall (W). Arrowheads point to oil bodies. Bar, 20 μm.

Figure 8.

Electron micrographs of mature cotyledons. (a) Electron micrograph showing the uniformity of mature cotyledon cells. Black protein bodies are prominent with oil bodies appearing as a light staining perimeter. (b) Electron micrograph of single cell showing the black protein bodies (P) and individual oil bodies mainly decorating the periphery of the protein bodies and the plasma membrane (arrows). Starch containing amyloplasts (A) is indicated. (c) High-magnification electron micrograph showing arrangement of individual oil bodies (O) around the protein bodies (P). Bars, 20 μm (a); 5 μm (b); 0.5 μm (c).

Figure 9.

Electron micrograph of seed radicle. Columnar cells in the radicle possess fewer, more heterogeneously sized oil bodies relative to cotyledons, which preferentially line the plasma membrane. Compared with cotyledon cells, root cells contain only few protein bodies (P) which are not surrounded by oil bodies. Bar, 2.5 μm.

These morphological studies of embryogenesis provide a base for understanding how the biogenesis of the major seed storage organelles, oil and protein bodies relates to the embryogenic program in legumes (Table 1). In addition to the morphological studies, we examined lipid accumulation and the expression of oleosin and storage protein genes in relation to embryogenesis.

Table 1.   Schedule for embryo development
Stage of embryo developmentTime after anthesis (d)Figure numbersCharacteristics
  1. RAM, root apical meristem; SAM, shoot apical meristem.

First cell division 03(b)Asymmetric; basal cell is larger with many small vacuoles
Second cell division 13(c)Gradient in vacuolation evident along apical–basal axis
Early embryo 24(a)Four-cell proembryo, hypophysis and prominent suspensor evident. Vacuoles and many plastids present in prominent suspensor
Early embryo 34(b)Multicellular proembryo and hypophysis are densely cytoplasmic
Globular stage 54(c)Hypophysis is prominent. Suspensor cells are large with many plastids
Heart stage 64(d)Hypophysis now integrated into the embryo proper. Suspensor is composed of elongated cells
Early cotyledon stage – Suspensor 85(a,b,d)Suspensor cells are large with large nuclei, many plastids and wall ingrowths in basal cell
Early cotyledon stage – Embryo proper and seed 86(a,b)
Cotyledons have developed. Early SAM and RAM connected by the procambium. Endosperm is being absorbed. Seed coat cells adjacent to endosperm have amyloplasts and wall ingrowths
Cotyledon development116(c,d)Cotyledons develop, associated with vascularization. Characteristic organization of SAM and RAM is evident
Accumulation of protein and oil in cotyledons12-2810
Protein bodies with vicilin and legumin and oil bodies with characteristic lipid content develop
Mature embryo286(e),
Cotyledons fully developed. Oil and protein bodies prominent. Some amyloplasts present. Oil bodies localized around protein bodies and adjacent to plasma membrane. Radicle is prominent with oil bodies adjacent to plasma membrane

Fatty acid biosynthesis and expression of oleosin genes

The majority (94%) of fatty acid present in the pool of M. truncatula seed lipid, based on transesterification of lipid, consisted of palmitic (C16 : 0), oleic (C18 : 1n-9), linoleic (C18 : 2n-6) and linolenic (C18 : 3n-3) acids, with linolenic acid alone comprising 36% of total fatty acid (Fig. 10a). Accumulation of lipid begins 14 d after anthesis (DAA) and continues throughout cotyledon development (Fig. 10a,b). Using BLAST searches, we identified four oleosin genes in M. truncatula. Oleosins are proteins embedded within the phospholipid monolayer of oil bodies and act to prevent oil body coalescence (Huang, 1992). Phylogenetic analysis indicated that the four M. truncatula oleosins we identified were present in two clades (Fig. 11a). Expression profiling by qPCR revealed that, similar to lipid accumulation, oleosin genes begin expression at 14 DAA with continued high expression until 32 DAA (Fig. 11b). The timing of oleosin gene expression thus parallels that of lipid accumulation.

Figure 10.

Accumulation of the major fatty acids and total protein during seed development (a) Four fatty acids – palmitic (C16 : 0), oleic (C18 : 1n-9), linoleic (C18 : 2n-6) and linolenic acid (C18 : 3n-3) – accounted for 94% of the final lipid fatty acid content of seeds. (b) Accumulation of total lipid fatty acid. (c) Accumulation of protein. (d) Seed water content. Values are means ± SE, = 3.

Figure 11.

Phylogram of oleosins and oleosin, vicilin and legumin expression in seed development. (a) Phylogram of selected oleosins from oilseed species. Four Medicago truncatula oleosin proteins encoded by Mt1g11190, Mt3g133000, Mt5g036320 and Mt6g005060 were identified and reside in clades I and II of the tree (see Supporting Information, Table S1, for sequences used in construction of the phylogram). (b) Expression profile of representative oleosins from clades I and II, and vicilin and legumin gene families during seed development. Values are means ± SE (= 3). See Table S2 for primer sequences used for quantitative real-time polymerase chain reaction.

Expression of storage protein genes

Total seed protein increases throughout seed development (Fig. 10c) concomitant with decreasing seed water content (Fig. 10d). We identified vicilin and legumin genes using sequence information available in public databases (Phytozome, GenBank and the M. truncatula gene index) and from published data (Thompson et al., 2009). Expression of the legumin gene family occurs subsequent to 14 DAA and reaches maximum expression at c. 22 DAA. Expression of the vicilin gene family commences earlier than legumin at 12 DAA and peaks at 18 DAA. Following peak expression, the expression of both legumin and vicilin gene families rapidly declines (Fig. 11b). Note that the expression pattern of individual legumin and vicilin genes generally mirrored that obtained by profiling the entire family (data not shown). The expression of the storage protein genes therefore declines before oleosin gene expression and lipid accumulation are complete.


Our study (Table 1) provides a developmental and cellular base with which to integrate transcriptomic and proteomic investigations of M. truncatula seed development (Gallardo et al., 2003, 2007; Firnhaber et al., 2005; Thompson et al., 2009). In addition, knowledge of zygotic embryogenesis is important for proteomic and transcriptomic studies of somatic embryogenesis in M. truncatula (Imin et al., 2005; Mantiri et al., 2008; Rose et al., 2010). While much of the developmental biology we present here is consistent with the current understanding of angiosperm embryogenesis, features of embryogenesis in M. truncatula provide additional perspectives to the process.

The embryo sac and early embryogenesis

The M. truncatula embryo sac derives from the functional megaspore (Fig. 1b), similar to M. sativa (Reeves, 1930). By sectioning from the micropylar to chalazal end of the embryo sac, we identified that the M. truncatula embryo sac has a Polygonum-type organization (Reiser & Fischer, 1993) similar to Arabidopsis (Mansfield et al., 1991) and is the usual organization in soybean (Kennell & Horner, 1985) and other legumes (Rodriguez-Riano et al., 2006). The Polygonum-type arrangement involves the antipodals, polar nuclei of the central cell, synergids and the egg cell (Fig. 2b).

The M. truncatula zygote has a characteristic intracellular asymmetry with a large vacuole at the micropylar end. The positioning of the vacuole is significant in the subsequent development of the embryo (Schulz & Jensen, 1968). As evident from Fig. 3b, the first division is asymmetric and produces a densely cytoplasmic apical cell and a basal cell with numerous vacuoles. This division is important in establishing an axis of polarity and hence determines subsequent embryo and suspensor development.

Embryo development

Following the first asymmetric division, the apical cell develops into the majority of the embryo, while the basal cell forms the suspensor. The embryo body plan is rapidly established (Fig. 4a) with the development of the embryo proper and the suspensor. The embryo proper appears similar to Arabidopsis (Mansfield & Briarty, 1991). However, at this stage of embryo development, differences between Arabidopsis and M. truncatula become apparent. Whereas the Arabidopsis suspensor consists of a single file of cells, the M. truncatula suspensor comprises a large number of highly vacuolated cells. Moreover, in Arabidopsis the hypophysis develops from division of the upper cell of the suspensor (Mansfield & Briarty, 1991). Our observations of four-cell-stage M. truncatula embryos and subsequent development (Figs 3c, 4a,b) are consistent with hypophysis development from the basal cell of the first apical division and the development of the two cells below this into the suspensor. Cells of the embryo proper are densely cytoplasmic and, similarly, cells of the hypophysis have fewer vacuoles whereas cells of the suspensor are highly vacuolated. Our interpretation of early embryogenesis in Fig. 4a,b is that it is the dome-shaped apical region, together with the adjoining multicellular hypophysis, that develops into the fully formed embryo – consistent with the subsequent development of the embryo into globular (Fig. 4c) and heart stages (Fig. 4d). It is unclear from the study of Chamberlin et al. (1994) how the hypophysis develops in soybean. Previous studies on a range of legumes do not identify the hypophysis – referring only to the proembryo or embryo proper and the suspensor (Martin, 1914; Weinstein, 1926; Yeung & Clutter, 1978). Indeed, in Phaseolus vulgaris, Weinstein (1926) reported no distinct separation between the embryo proper and the suspensor, a finding consistent with investigations of Phaseolus coccineus (Yeung & Clutter, 1978). Current understanding in Arabidopsis indicates that the hypophysis gives rise to the quiescent centre and root cap (Laux & Jurgens, 1997). Our observations of M. truncatula (Fig. 4a–d) are consistent with the hypophysis being a precursor to the major part of the RAM and root cap.

The body of evidence suggesting that the suspensor plays a role in early embryo development in legumes (Yeung & Clutter, 1978, 1979) concurs with our observations in M. truncatula. In Phaseolus, the large suspensor cells are endopolyploid, with nuclei containing polytene chromosomes (Nagl, 1969). In M. truncatula, the large nuclear volume (relative to cells of the embryo proper) is consistent with endopolyploidy, although we saw no evidence for polytene chromosomes as occurs in P. vulgaris. It seems reasonable that the suspensor plays a role in the uptake and production of substances required for embryo development until cotyledon formation, consistent with the development of substantive wall ingrowths in the basal suspensor cell (Yeung & Clutter, 1979). There is considerable structural diversity in legume suspensors (Le et al., 2007), but the relationship between the suspensor and embryo or seed size is not clear. Interestingly, the M. sativa suspensor with a single file of cells and an elongated basal cell (Martin, 1914) contrasts with that in M. truncatula. Recent transcriptome investigations in soybean have shown that the suspensor has a transcriptional profile different from the rest of the seed (Le et al., 2007).

West & Harada (1993), in a general consideration of angiosperm embryogenesis, point to the partitioning of the embryo into protoderm, ground tissue and procambium. It is noteworthy that the procambium is the first pluripotent tissue to develop in M. truncatula, preceding both the shoot apical meristem (SAM) and the RAM. This sets the scene for future plant development, with a continuum between the early SAM, procambium and RAM (Fig. 6b) emerging before a fully developed SAM and RAM are apparent (Fig. 6d). It is noteworthy that during in vitro culture, in suitable experimental systems, the procambium can initiate embryo (Guzzo et al., 1994) or RAM (Rose et al., 2006) development. When embryogenesis completes, fully developed cotyledons, with their storage products and the radicle are evident (Fig. 6e). We find that the radicle is large and well developed in M. truncatula and, presumably, this enables rapid establishment of the seedling in soil after germination.

Oil and protein bodies are numerous in the mature cotyledons, but there are few starch-containing amyloplasts (Fig. 8b), which contain much less starch than the amyloplasts present in the hilum region of the seed coat (Fig. 5c). In early embryogenesis the suspensor has many plastids containing small amounts of starch, contrasting with the embryo axis. In the heart stage embryo, plastids are more abundant and contain small amounts of starch. As the cotyledons develop there appears to be a slow accumulation of starch in the plastids but little or no plastid proliferation.

The oil and protein bodies of the mature cotyledons of the seed

Protein bodies occupy most of the cell in M. truncatula cotyledons. The current understanding is that protein bodies develop by trafficking of protein in endoplasmic reticulum-derived vesicles into storage vacuoles (Herman, 2008). Oil bodies surround protein bodies and line the plasma membrane, but are only infrequently free in the cytoplasm. While oil and protein bodies were visualized many years ago (e.g. Lott & Buttrose, 1978), the cellular arrangement of oil bodies in plant cotyledons has received essentially no attention (e.g. Lott & Buttrose, 1978; Hsieh & Huang, 2004; Herman, 2008). Our observations clearly show that the arrangement of oil bodies is not random. Potentially, the low concentration of fatty acid per unit DW in M. truncatula cv Jemalong (8.8–11.6%) may make it easier to visualize oil body arrangement (Tonnet & Snudden, 1974; Djemel et al., 2005). The Lott and Buttrose electron microscopy study (1978) on protein bodies in several legumes shows oil bodies arranged around the perimeter of protein bodies, with the uniformity of oil body size and shape dependent on the species. The Lott & Buttrose (1978) study, however, focused on protein bodies with no discussion of oil bodies. The arrangement does not reflect some common physicochemical relationship to all cellular membranes as the regular arrangement does not hold for amyloplasts or nuclei. At higher resolution there certainly appears to be a very close association, although it is clear that there is no fusion in the same way that there is no coalescence between oil bodies.

The arrangement of oil bodies we observed most likely results from the presence of oleosin in the oil body membranes. Oleosin prevents coalescence between oil bodies, and indeed down-regulation of oleosin in Arabidopsis and soybean leads to increased coalescence and decreased oil production (Siloto et al., 2006; Schmidt & Herman, 2008). Interestingly, the arrangement of oil bodies on the inside of the plasma membrane may have a cellular protective effect during seed desiccation. Accordingly, knockdown of oleosin in soybean, which leads to the formation of giant oil bodies, results in few, if any, viable cotyledon cells after hydration and germination (Schmidt & Herman, 2008). In cells of the radicle, where there are fewer oil bodies per cell than in the cotyledons, the preferential alignment of oil bodies along the plasma membrane might play an additional role. Here, oil bodies at the plasma membrane potentially provide a localized resource of easily manipulated carbon skeletons, associated with lipid and phospholipid, to facilitate rapid expansion of the plasma membrane in enlarging radicle cells during germination.

Higher oil content in Brassica napus is associated with higher oleosin (Hu et al., 2009), and oil body size and shape relate to the oil : oleosin ratio (Hsieh & Huang, 2004). A feature of the oil bodies in M. truncatula is their small size and regular ovoid shape compared with the much larger protein bodies of irregular shape. In M. truncatula, we identified four oleosin genes that fall into two distinct clades (Fig. 11a). It would be of interest to determine how isoforms of oleosin from these distinct clades contribute to the cellular arrangement of oil bodies.

The change in legumin and vicilin expression during seed development is similar to that reported for M. truncatula (cv Jemalong) by Gallardo et al. (2007), with vicilin expression peaking before legumin expression. The timing of oleosin, vicilin and legumin biosynthesis (Fig. 11b) is consistent with a highly coordinated production of both oil and protein bodies. Indeed, there is a remarkable coordination between lipid accumulation and oleosin expression and in the expression of the major storage proteins. In Vicia faba, antibody studies show that legumin and vicilin occur in the same protein bodies (Graham & Gunning, 1970). The coordination also involves acquiring a characteristic protein : oil ratio. M. truncatula can usefully serve as a model to explore more fully the coordination of oil and protein body biogenesis in legumes.


This research was supported by an Australian Research Council (ARC) Centre of Excellence Grant (CEO348212) to R.J.R. and by an ARC Australian Postdoctoral Fellowship (DP0770679) to M.B.S.