Degeneration of oil bodies by rough endoplasmic reticulum -associated protein during seed germination in Cannabis sativa

Abstract

Introduction

Seed plants accumulate nutritional sources such as protein, lipids and carbohydrates in the endosperm or cotyledon for germination and post-germinative growth of the seedlings. Photosynthetic sugars are often polymerized into starch in amyloplasts or converted into lipids and stored as lipid droplets (LDs) in seeds. LDs are present in all plant cell types, ranging from a few LDs per cell in leaves to thousands of LDs per cell in seeds (Kang et al. 2022). LDs are often referred to in the literature by various terms, for example, lipid bodies, oil bodies, oleosomes or spherosomes depending on the characteristics of the species (Hsieh and Huang 2004; Purkrtova et al. 2008; Sánchez-Albarrán et al. 2019; Zienkiewicz and Zienkiewicz 2020; Nazari et al. 2022; Guzha et al. 2023). These organelles are surrounded by a half membrane and characteristically integrated with specific structural proteins (Tzen et al. 1993; Thiam et al. 2013; Huang 2018; Ischebeck et al. 2020).

LDs are not merely energy storage organelles, but also dynamic structures involved in diverse cellular metabolisms like membrane remodelling, regulation of energy homeostasis, stress responses and coordination between different organelles (Choi et al. 2022; Bouchnak et al. 2023). In addition, they play crucial roles at key sites in the freezing tolerance of seeds, engaging in direct interaction with glyoxysomes for seedling lipid degradation and producing antifungal compounds in leaves (Shimada et al. 2018; Olzmann and Carvalho 2019).

Lipids found in oilseeds are composed of a hydrophobic core filled with triacylglycerols (TAG), which are the most common storage lipids (Quettier and Eastmond 2009; Horn et al. 2013; Goold et al. 2015). The structural proteins of LDs commonly contain three membrane proteins known as oleosin, caleosin and steroleosin (Hsiao and Tzen 2011; Chapman et al. 2012; Huang 2018; Shimada et al. 2018). They are anchored in the phospholipid monolayer by a hydrophobic α-helical hairpin domain with a proline knot, and the C- and N-termini face of the cytosol (Alexander et al. 2002; Capuano et al. 2007; Purkrtova et al. 2008).

Oleosins are the most abundant integral membrane proteins of LDs in oilseeds. Particularly, the lipid droplet-associated proteins stabilize the LDs and prevent the coalescence or aggregation of this organelle in mature seeds (Huang 1992, 1994; Miquel et al. 2014). Hence, they are important regulators of LD dynamics; their ubiquitination, extraction and proteasomal degradation precede LD breakdown (Deruyffelaere et al. 2015; D’Andrea 2016).

Storage oil mobilization usually begins with seed germination. As a carbon or energy source in the germinating seeds, storage oil contributes to providing free-fatty-acids released from TAG by lipase or sugars through free-fatty-acid degradation by β-oxidation with subsequent gluconeogenesis (Graham 2008; Hielscher et al. 2017). The pathway of storage lipid conversion to sugars was examined in germinating lupin seeds (Borek and Ratajczak 2010; Borek et al. 2017). Subsequently, all the storage compounds are remobilized during post-germinative growth (Babazadeh et al. 2012; Miray et al. 2021).

Recently, some researchers reported on the LD degradation system in plants (Farquharson 2018; Kretzschmar et al. 2018; Traver and Bartel 2023). Since LDs are strongly associated with the endoplasmic reticulum (ER) (Suzuki 2017), this association has been observed at the electron microscopy level in many organisms (Fujimoto et al. 2013; Ohsaki et al. 2017; Sui et al. 2018; Renne et al. 2020; Kang et al. 2022; Zhang et al. 2022). During germination and seedling establishment, glyoxysomal enzymes degrade oil bodies to release storage lipids in seeds (Graham 2008; Shimada et al. 2018). Peroxisome contains the triacylglycerol lipase SUGAR-DEPENDENT1. This lipase is associated with the surface of the peroxisomes, and it is translocated to the oil body surface during seedling establishment (Thazar-Poulot et al. 2015).

Cannabis seeds contain approximately 18–30 % protein, 30–40 % oil and 25–34 % carbohydrate (Leonard et al. 2020; Vasantha Rupasinghe et al. 2020). Much of the knowledge of LD function in plants comes from studies of oilseeds (Laibach et al. 2015; Pyc et al. 2017b; Chen et al. 2023). Despite the importance of storing fats, oils and wax in seeds, our knowledge of the specificities of lipid metabolism remains uncertain.

This study represents a fundamental step towards the morphological elucidation of the mobilization mechanism of storage compounds in seeds. This research aimed to determine (i) the structural characteristics of storage compounds in the cotyledons of Cannabis, (ii) the degradation pathway leading to the β-oxidation of storage oil during seed germination, and (iii) the relationship between storage organelles such as oil bodies and protein bodies in oilseeds.

Materials and Methods

The achenes of fibre hemp cultivar (Cannabis sativa cv. ‘Chungsam’) obtained from Dangjin Agricultural Technology Centre (DATC), South Korea were used in this study. Dangjin area located in Chungcheong Province in South Korea (37°03ʹN, 126°51ʹE) provides favourable environmental conditions for high-quality hemp seeds. These achenes were collected from an approved farm by DATC 8 months prior. They were stored in a seed storage chamber of DATC at 4 °C. Twenty achenes were germinated for two days on sheets of filter paper moistened with sterile water in glass Petri dishes (150 mm × 20 mm) in an incubator with 65 % relative humidity and 20 °C under darkness (Blandinières et al. 2021). The embryo samples were obtained from the germinating seeds at various times in the growth phase; early (12h), middle (18 h) and late stage (24 h) after germination.

For light microscopy, the seeds were dissected with a razor under the stereoscopic microscope and fixed for 2 h in 2 % glutaraldehyde in 25 mM phosphate buffer, pH 7.2. After being rinsed with deionized water, they were post-fixed for 1 h in 2 % osmic acid and dehydrated with a graded ethanol series (50, 70, 80, 90, 95, 100 % ethanol). Then the samples were embedded in Spurr’s resin for 14 h and polymerized for 48 h at 60 °C. Semithin sections of 0.4 μm in thickness were cut on an ultramicrotome (Reichert Ultracut S, Leica, Germany) with glass knives and stained with toluidine blue-basic fuchsin. For the histochemical study, the fresh sections of the embryo tissue were touched on the slide glasses and stained with Sudan III, Alcian blue and Astra Blue. All the samples were observed and photographed using a light microscope (Axiophot II, Zeiss, Germany).

For scanning electron microscopy (SEM), the achenes were fixed in the same protocol described in the light microscopy sample preparation. Then the samples were transferred into isoamyl acetate. The samples were subjected to critical point drying with pressurized liquid carbon dioxide (Bioradical E3000, Bio-Rad, USA). The dried specimens were mounted on aluminium stubs, coated with gold-palladium in a sputter coater (JFC-1110E, JEOL, Japan), and photographed in a FE-SEM (JSM-6700F, JEOL, Japan) at 15 kV.

For TEM, the seeds were treated with the SEM fixation method described above. The materials were dehydrated with a graded ethanol series and replaced with propylene oxide. Subsequently, ultra-thin sections of 70 nm thickness were cut with a diamond knife (Micro Star SU-30, Ted Pella, USA) using an ultramicrotome (Reichert Ultracut S, Leica, Germany) and sections were collected on 300 mesh copper grids. The sections were stained for 20 min with 1 % uranyl acetate and for 10 min with 1 % lead citrate. Image acquisition was performed with a transmission electron microscope (JEM-2000 EX II, JOEL, Japan) at 80 kV.

Results

Cannabis achenes have a hard pericarp encasing a single seed. In this study, the achenes varied in length from 4 to 5 mm, and in diameter from 3 to 4 mm (Fig. 1A and ​andC).C). The seed consisted of an endosperm and an embryo with two cotyledons and a radicle (Fig. 1B and ​andD).D). The axis of the Cannabis embryo was curved and contained a U-shaped feature (Fig. 1B). The tip of the radicle and cotyledons were oriented toward the stylar end of the achene (Fig. 1B and ​andD).D). When the germination began, the radicle emerged from the pericarp at the stylar end and split the seed coats into halves that were attached at the base (Fig. 1C). The scanning electron micrographs of the Cannabis seed showed that it consisted of endosperm, two distinctive cotyledons (outer, and inner cotyledon) and a radicle in a piece of the embryo (Fig. 2A). Specifically, the endosperm was confined to a peripheral region between the inner cotyledon and radicle in the mature seed (Fig. 2B). The deshelled seed was smooth and oval or orbicular in form as well as the enclosed seed (Fig. 3A). The epidermal cells of the embryo were rectangular in shape and arranged end to end in rows. They were equal approximately 15 µm in width but differed in length, the longer one being 60 µm and the shorter 14 µm. (Fig. 3B). The cotyledonary cells, functioning as storage, contained numerous oil bodies and protein bodies (Fig. 3C).

The cotyledons comprised several layers of parenchymatous cells (isodiametric cells) and two or more layers of palisade cells (Fig. 4A). TEM images revealed that the protein bodies of cotyledon cells measured between 2.5 and 3.5 µm in diameter (Figs. 4C, 5A–C). Large protein bodies are surrounded by many oil bodies ranging from 0.7 to 1.8 µm in diameter. However, the size of the extracted oil bodies varied from 0.1 to 2 µm (Fig. 5F). Oil bodies in Cannabis seed appeared spherical and were sporadically distributed in the cells (Fig. 5D and ​andE).E). Isolated oil bodies were obtained by smearing small pieces of cotyledon onto a microscope slide and staining them with Sudan III (Fig. 5F).

Protein bodies in Cannabis seeds contained electron-dense globoids with a heterogenous protein matrix. The storage proteins were concentrated in the centre of the protein body as a dense homogenous circular mass surrounded by a light heterogeneous area (Fig. 6A and ​andB).B). As the major seed storage organelles in Cannabis, protein bodies and oil bodies within the cotyledon cells underwent unique morphological changes throughout germination (Fig. 6B–D). The protein bodies and oil bodies gradually degenerated in the cells and were used as a primary energy source during germination. During germination, the rER was frequently present in all cotyledon cells (Fig. 7A and ​andD).D). As ribosomes and rER began to increase, dense substances were also concentrated in the outer region of oil bodies (Fig. 7B and ​andC).C). At the early stage of germination, dense substances aggregated adjacent to the oil bodies and associated with them (Fig. 8A and ​andB).B). Later, the border of the oil bodies became a dense cluster and appeared as a sinuous outline. In addition, irregular hyaline areas were distributed throughout the oil bodies, reflecting the destabilized emulsification of oil bodies (Fig. 8C). Finally, the oil bodies fused with one another and had an irregularly contoured surface (Fig. 8D).

Discussion

Our fundamental data on seed germination provides insight into the understanding of the degradation mechanisms controlling the metabolism of storage proteins in the cotyledon of oilseeds. As small subcellular storage organelles, the protein bodies and oil bodies in the cotyledon cells of Cannabis seeds are gradually degenerated and used as a primary energy source during germination. Particularly, the biological function of storage proteins correlated with oil bodies stored in the cotyledon cells appears to be more diverse than simply constituting a source of carbon made available for the germinating seedling.

Conclusions

As the storage organelles, protein bodies and oil bodies were packed in the cotyledon cells of Cannabis seeds. They remarkably changed the morphology at the early stage of germination. The storage proteins were concentrated in the centre of the protein body as a dense homogenous circular mass surrounded by a light heterogeneous area. Some of the storage proteins appear to act as emulsifying agents on the surface region of oil bodies. These proteins maintain the individuality and stability of oil bodies inside or outside the cotyledon cells. After rER appeared near the oil bodies, dense substances considered proteins rapidly aggregated adjacent to the oil bodies, resulting in the coalescence of oil bodies. We concluded that these rER-associated proteins play a key role in the degeneration of oil bodies by weakening the emulsifying agent on their non-polar surfaces and inducing the coalescence of oil bodies during seed germination. Finally, most oil bodies fused with one another and had an irregularly contoured surface at the late stage of germination.

Acknowledgements

Contributor Information

Eun-Soo Kim, Institute of Cannabis Research, Colorado State University-Pueblo, 2200 Bonforte Blvd. Pueblo, CO 81001-4901, USA.

Joon-Hee Han, Institute of Biological Resources, Chuncheon Bioindustry Foundation, 32, Soyanggang-ro, Chuncheon-si, Gangwon-do 24232, Republic of Korea.

Kenneth J Olejar, Department of Chemistry, Colorado State University-Pueblo, 2200 Bonforte Blvd. Pueblo, CO 81001-4901, USA.

Sang-Hyuck Park, Institute of Cannabis Research, Colorado State University-Pueblo, 2200 Bonforte Blvd. Pueblo, CO 81001-4901, USA.

Literature Cited