Plant Cell Reports (2000) 19 : 868±874
Springer-Verlag 2000
C E L L BI O L O G Y A N D M O R P HO G E N E S I S
A.V. Patel ´ I. Pusch ´ G. Mix-Wagner ´ K.D. Vorlop
A novel encapsulation technique for the production of artificial seeds
Received: 28 October 1999 / Revision received: 11 February 2000 / Accepted: 22 February 2000
Abstract A novel technique for the encapsulation of plant material in calcium alginate hollow beads was tested. The technique involves suspending plant material (i.e. plant cells, tissues, organs, shoot tips, somatic embryos) in a solution containing carboxymethylcellulose and calcium chloride and then dripping it into a stirred sodium alginate solution. In initial experiments with Daucus carota (carrot), it was found that after 14 days of cultivation, 100 % of seeds encapsulated in calcium alginate hollow beads would germinate in the liquid core and that 13 % would burst the capsules. Embryogenic calli developed inside hollow beads and formed somatic embryos while calli in conventional calcium alginate beads became detached from the beads early in development, and no somatic embryogenesis occurred. With Solanum tuberosum (potato), development of calli was observed in 50 % of hollow beads. Eighty-one percent of shoot tips encapsulated in hollow beads sprouted and grew out of the capsules. Key words Artificial seeds ´ Encapsulation ´ Microcapsule ´ Daucus carota ´ Solanum tuberosum
Communicated by H. Lörz A.V. Patel ()) ´ K.D. Vorlop Institute of Technology and Biosystems Engineering, Federal Agricultural Research Centre (FAL), 38116 Braunschweig, Germany e-mail:
[email protected] Fax: +49-531-596363 I. Pusch ´ G. Mix-Wagner Institute of Crop and Grassland Science, Federal Agricultural Research Centre (FAL), 38116 Braunschweig, Germany
Introduction Encapsulation materials and methods for the production of artificial seeds (Fig. 1) have seldom varied since the concept was introduced by Murashige in 1977. A range of publications on the encapsulation of somatic embryos in conventional calcium alginate beads have been published in the last two decades (reviews: Redenbaugh et al. 1987; Bajaj 1995a, 1995b). In all these cases, the embryos protruded from the beads or were located near the surface due to the process of bead production so that complete protection of the embryo as in a natural seed was not ensured. In a labour-intensive and costly second step, calcium alginate beads were coated with a synthetic, biologically nondegradable polymer (Redenbaugh et al. 1987) Such two-step procedures were also tested using polyethyleneimine (Kersulec et al. 1993) or chitosane (Tay et al. 1993) as the coating material. So-called hollow beads which can be produced in a simple onestep procedure have been reported in one paper only, but the liquid core contained a chitosane so that no embryos survived (Tay et al. 1993). Other encapsulation materials and methods commonly used in classical biotechnology (Vorlop and Klein 1983) should be tested. The use of hollow beads seems to be especially promising in the realization of true artificial seeds. Calcium alginate hollow beads were successfully used to encapsulate highly sensitive animal cell cultures (Spiekermann et al. 1987) or entomopathogenic nematodes (Patel and Vorlop 1994). We now suggest using this type of encapsulation method for the encapsulation of plant cells, shoot tips, or somatic embryos, for conservation purposes and especially for the production of artificial seeds (Fig. 1). The advantages are summarized in Table 1 . Here, we present initial results on the encapsulation of embryogenic calli of Daucus carota (carrot), as well as calli and shoot tips of Solanum tuberosum (potato).
869 Fig. 1 Production of artificial seeds
Table 1 Advantages of somatic embryos encapsulated in hollow beads Advantages compared with natural seeds
Advantages compared with naked somatic embryos
Advantages compared with somatic embryos encapsulated in conventional beads
Quick clonal propagation
Easy handling (storage, sowing)
Economical propagation of plants which can not be easily propagated via natural seeds (i.e. hybrids, transgenic plants, trees) Economical production of virus-free plants Easy maintenance of the genetic resources
Protection from extreme environmental conditions (temperature, dryness)
Closer to natural seeds because of complete protection of somatic embryo in sterile core Continued development of somatic embryo within the hollow bead
Protection from pathogens
No early emergence
Enhanced shelf life
Entrapment and coating realised in one step
Materials and methods
Table 2 Phytohormone concentrations (mg/l) used for the induction of embryogenic callus of Daucus carota
Chemicals
Hormones
Variant 1a
Variant 2a
Variant 3b
Sodium alginate (Protanal LF20/60) was obtained from Protan (Norderstedt, Germany) and carboxymethylcellulose (Blanose 7MXF) from Aqualon (Düsseldorf, Germany). Gelrite and agaragar were from Roth (Karlsruhe, Germany). The basal medium used in these experiments comprised the inorganic and organic salts of a Murashige and Skoog (1962) (MS) medium at a pH of 5.8 and was autoclaved for 10 min at 121 C. Phytohormones were then added by sterile filtration.
2,4-D IAA BAP K
0.1 0.2 ± 0.2
0.25 ± 0.25 ±
0.5 ± ± ±
Production of embryogenic callus of Daucus carota
Propagation of S. tuberosum plants
Mature roots of D. carota cv. `Nantaise' were washed in tap water, cut into 5-cm-long pieces, submersed in 96 % ethanol for 15 s and then washed with deionized water. The pieces were then sterilized for 30 min in a solution of sodium hypochloride (Domestos, Unilever, Hamburg; diluted 1:10) and washed with sterile deionized water three times 3±4 min. After the root tips and 2±3 mm of the cortex had been cut away, the pieces were cut into 2±3 mm thin discs which again were divided into quarters. These explants were put onto MS medium (two to six explants per petri dish) and incubated at 26 C. The MS medium contained 30 g/l sucrose, 10 g/l agar, 0.1 mg/l pyridoxine hydrochloride and phytohormones (Table 2). Every 4 weeks, the explants were transferred onto fresh medium.
In a preliminary screening, the health and growth speed of sterile in vitro plants of the cultivars `Priwal', `Kromak', `DesirØe', `Profijt', `Sieglinde' and `Simson' of S. tuberosum L. ssp. tuberosum of the culture collection BGRC (BAZ, Germany) were monitored. The plants were raised in 6- to 8-cm-high baby-food jars (sealed with Magenta B Caps) on MS medium at 22 C and a 16-h photoperiod provided by warm white fluorescent lights (36 mmol ´ m ± 2 ´ s ± 1, overlight system). The MS medium (25 ml) contained 10 g/l sucrose, 3.4 g/l Gelrite and was autoclaved for 10 min at 121 C. Plants were propagated by cutting off sterile shoot buds. The shoot buds were placed into MS medium. The rest of the shoot was cut into pieces containing two nodes each with the lower
a b
Cultivation in the dark Cultivation with 12 h day length (96 mmol ´ m ± 2 ´ s ± 1)
870 leaf removed and was also placed in MS medium. One propagation cycle lasted 3 weeks. Of the cultivars tested, `Priwal', `Kromak' and `DesirØe' showed the fastest growth during propagation (four- to fivefold increase of shoot length within 14 days). Of these, `Priwal' was used for the production of callus and the preparation of shoot tips. Preparation of shoot tips Under a stereo-microscope placed on a clean bench, the shoot buds of the main shoots of 6-cm-high in vitro plants of S. tuberosum cv. `Priwal' were dissected according to Schäfer-Menuhr et al. (1998). Production of callus of S. tuberosum For induction of callus in S. tuberosum cv. `Priwal', half-strength MS medium containing 20 g/l sucrose and 3.4 g/l Gelrite was supplemented with the phytohormones kinetin (0.3 mg/l), indole-3-acetic acid (0.2 mg/l) and 2,4-dichlorophenoxyacetic acid (2.4 mg/l). After autoclaving, 25 ml of this callus induction medium was poured into each glass petri dish. Sterile shoots were cut into 3-mm pieces, and leaves were cut at least twice (depending on size), and all explants were placed onto the medium. These were then cultivated at 26 C in the dark for 5±7 weeks. Encapsulation methods Calcium alginate hollow beads Plant material (i.e. plant cells or shoot tips) was mixed with 10 ml of 1.5 % carboxymethylcellulose solution containing 1 % calcium chloride. Then the suspension was dropped with a 10-ml syringe into 400 ml 0.8 % sodium alginate solution stirred in a 1 l beaker. At the surface of each drop, a calcium alginate layer formed from the inside to the outside (Fig. 2), while the core remained liquid. After 10 min of gelation, the alginate solution containing the hollow beads was diluted with 400 ml deionized water. Then the diluted solution was decanted. This washing procedure was repeated twice and prevented the capsules from sticking together. Finally, the hollow beads were transferred into Fig. 2 Encapsulation of somatic embryos in beads compared with hollow beads
a stirred 1 % calcium chloride solution and left to harden for another 20 min. The resulting capsules consisted of a liquid core surrounded by a calcium alginate membrane. The membrane thickness can be adjusted by the gelation time and the concentration of the chemicals (0.1±2 mm). In this paper, the thickness ranged from 0.3 to 0.5 mm. This variation results from the fact that the dropping time adds to the gelation time (the gelation time for the the first droplets is longer than for the last ones). Further details regarding the principles of the gelation process are mentioned in Vorlop and Klein (1983) and in Vorlop et al. (1987). Calcium alginate beads Plant material (i.e. plant cells) was mixed with 10 ml of 1.5 % sodium alginate solution and dropped with a 10-ml syringe into a 1 % calcium chloride solution in a 1-l beaker. The beads were formed by gelation from the outside to the inside (Fig. 2). Gelation time was 20 min. Encapsulation of D. carota seeds in calcium alginate hollow beads Seeds of D. carota cv. `Nantaise' were encapsulated to investigate the growth of plants in the liquid core of the hollow beads. After 24 h steeping in tap water, seeds were sterilized in 2 % calcium hypochloride for 15 min. Then, they were thoroughly washed with sterile deionized water and encapsulated. Twenty hollow beads, each containing only one seed, were transferred into 100 ml MS medium (10 g/l sucrose) in 200 ml Erlenmeyer flasks sealed with aluminium caps and were cultivated at 26 C in the dark at 120 rpm. After 4 days, germination started and cultivation took place at 75 rpm and a 12-h photoperiod provided by warm white fluorescent lights (10 mmol ´ m ± 2 ´ s ± 1). As a control, soaked and sterilized seeds were cultivated likewise. Encapsulation of embryogenic callus of D. carota in calcium alginate beads and calcium alginate hollow beads and induction of somatic embryogenesis We experimented to see whether plant cells survive the encapsulation process and grow in calcium alginate hollow beads. For comparison, plant cells were also encapsulated in conventional
871 calcium alginate beads. Therefore, 0.05±0.8 g of minced callus (Table 2, variant 1±3) was mixed with 10 ml of dropping solution (either carboxymethylcellulose/calcium chloride or sodium alginate) and then dropped into the corresponding crosslinking solution. The diameter of the resulting capsules was 3 mm (beads) or 5 mm (hollow beads). Cultivation of encapsulated callus Fifty to sixty beads and 50±60 hollow beads were placed in 250-ml cell culture flasks with 100 ml liquid MS medium supplemented with 30 g/l sucrose, 0.1 mg/l thiaminium dichloride, and the following phytohormones: 0.5 mg/l indole-3-acetic acid, 0.2 mg/l gibberellic acid and 0.5 mg/l zeatinriboside. As a control, calli were cultivated in Erlenmeyer flasks sealed with aluminum caps and parafilm. Cultivation took place at 75 rpm and a 12-h photoperiod provided by warm white fluorescent lights (10 mmol ´ m ± 2 ´ s ± 1) at 26 C. Cultures were subcultured to fresh medium every week. After 3 weeks, all capsules were checked for development of calli. In order to test if the induction of somatic embryogenesis and the development of somatic embryos in the capsules is possible, encapsulated embryogenic calli grown in beads and hollow beads were transferred into hormone-free MS medium (same culture conditions as before). Encapsulation of callus of S. tuberosum in calcium alginate hollow beads Callus cells of the widely used model system D. carota grew very well in calcium alginate hollow beads (see Results and discussion). We tested to see if this was also true for S. tuberosum. Therefore, 0.05 ± 0.8 g minced callus was mixed with 10 ml 1.5 % carboxymethylcellulose solution containing 1 % calcium chloride and then dropped into a solution containing 0.8 % sodium alginate. The capsule diameter was 5 mm 0.64 mm and the wall thickness was 0.5 0.07 mm. The capsules were then cultivated as the encapsulated D. carota calli (see above). Encapsulation of shoot tips of S. tuberosum in calcium alginate hollow beads The shoot tips were encapsulated as described above for seeds and calli. Then, the encapsulated shoot tips (5 mm capsule diameter, 0.3 mm capsule wall thickness) were cultivated in 6-cmhigh glass culture vessels on solid MS medium supplemented with 2.5 g/l Gelrite. Cultures were kept at a 12-h photoperiod provided by warm white fluorescent lights (10 mmol ´ m ± 2 ´ s ± 1). As a control, unencapsulated shoot tips were similarly cultivated.
Results and discussion Daucus carota Growth of D. carota plants from seeds encapsulated in calcium alginate hollow beads The development of plantlets growing from seeds encapsulated in calcium alginate hollow beads was monitored. The seeds germinated after 4 days in the dark; after 14 days of cultivation in fluorescent light, plantlets had developed inside all of the hollow beads (Fig. 3). Of these, 13 % had burst the capsule. In the
Fig. 3 Daucus carota plant grown from a seed in a hollow bead. 1 Shoot, 2 root, 3 seed coat
control, only 70 % of the seeds germinated. The reason is not known. Encapsulated plantlets grew more slowly inside the capsules than the controls, which grew into full plants in 6 days. This effect is possibly due to internal diffusional limitations which have been investigated for many immobilized biocatalysts (Klein et al. 1984). Growth and somatic embryogenesis of embryogenic callus encapsulated in calcium alginate beads and calcium alginate hollow beads In these experiments the growth and development of embryogenic callus in calcium alginate beads and calcium alginate hollow beads was compared. In both bead forms, the initial cell concentration was so low after encapsulation that no cell aggregates could be visually detected. Growth of visible calli could be observed after 3 weeks. It was also observed that calli encapsulated in the centre of calcium alginate beads exhibited poor growth, and calli emerging from the surface of these beads were friable and hard (Fig. 4). When growing in liquid culture, the cells growing out of the beads resulted in a turbid medium. Furthermore, it was found that more visible calli developed in calcium alginate hollow beads than in calcium alginate beads (Table 3). Calli could grow more easily inside hollow beads because they were protected from shear forces. Also, a toxic effect of the calcium alginate matrix used cannot be ruled out. This could be more pronounced in beads where the calli are in direct contact with the matrix than in hollow beads where the calli grow in the liquid core without direct contact with the alginate. It was observed that controls which could grow without diffusional or spatial limitations developed faster than the encapsulated calli. In general, it can be concluded that plant cells of the widely used model system D. carota are able to
872 Table 3 Development of calli of D. carota encapsulated in calcium alginate beads and calcium alginate hollow beads cultivated in liquid culture Developed calli ( %)
Beads Hollow beads
Variant 1
Variant 2
Variant 3
12 71
31 39
39 69
Regarding calcium alginate hollow beads, after 5 weeks of cultivation some calli started to form dark structures (Fig. 4A) in the core of the hollow bead and in one case (variant 3), plantlets were formed (Fig. 4B). As regards calcium alginate beads, after 5 weeks of cultivation calli had not developed inside the beads but had grown outwardly (Fig. 5). This happened because the calli are located near the surface of the bead. Calli of a size of about 5 mm became detached from the beads. Although dark structures on the calli were observed, no somatic embryogenesis occurred in any of the media.
Solanum tuberosum Development of calli of S. tuberosum in calcium alginate hollow beads Fig. 4 A Embryogenic D. carota callus with dark structures grown out of a calcium alginate hollow bead. B D. carota plantlets formed via somatic embryogenesis from embryogenic callus grown out of a hollow bead
Experiments with the model system D. carota had shown that calli could grow inside hollow beads. To check whether this was also true for S. tuberosum, calli were encapsulated in calcium alginate hollow beads. After 14 days of cultivation in MS medium, calli had grown inside hollow beads. Out of 240 capsules, 124 (50 %) showed callus growth. Sixteen (13 %) contained calli which filled the capsule to more than three quarters (Fig. 6). It should be noted that the initial cell concentration might have been too low in those hollow beads not showing growth of calli. In any case, this experiment showed that cells of S. tuberosum could survive encapsulation in calcium alginate hollow beads and grow within the liquid core.
Development of shoot tips of S. tuberosum in calcium alginate hollow beads Fig. 5 Callus of D. carota attached to a calcium alginate bead
grow not only in calcium alginate beads but also in calcium alginate hollow beads. Embryogenic calli (variant 1±3, Table 2) grown in calcium alginate beads and hollow beads were transferred to hormone-free MS medium in order to induce somatic embryogenesis.
Since shoot tips can produce new plants and are used in classical micropropagation, we investigated whether they can grow inside calcium alginate hollow beads and break the capsule wall to form whole plants. It was found that all shoot tips developed in the liquid core of the hollow beads into whole plantlets. After 4 weeks, 13 out of 16 (81 %) of the shoot tips had developed whole plants (Fig. 7), while two plant-
873
Fig. 6 Growth of calli of S. tuberosum in calcium alginate hollow beads after 14 days of cultivation. 1 Callus filling the capsule to more than 3/4, 2 poor callus growth
Fig. 8 A,B Development of shoot tips of S. tuberosum encapsulated in calcium alginate hollow beads. A Shoot tips developing inside the hollow beads after 2 weeks. B Plantlets growing from hollow beads after 4 weeks
Fig. 7 Growth of shoot tips of S. tuberosum from calcium alginate hollow beads on MS medium
lets (13 %) had remained in the capsule. One shoot tip (6 %) had died, possibly due to damage during the shoot tip preparation process. The development of encapsulated shoot tips is shown in Fig. 8. In some cases, malformed shoot tips were observed and some shoot tips formed two to three shoots. One shoot grew out of the apical meristem and the other(s) out of the leaf primordia. Furthermore, some shoot tips started rooting within the capsules. Similar results were obtained by cultivation of encapsulated shoot tips in liquid MS medium (data not shown). Controls developed within 2 weeks into 1- to 2-cmhigh plantlets. Some of the regenerants also showed two shoots.
The results demonstrate that hollow beads based on ionotropic gels can be used for the encapsulation of plant cells and shoot tips. In special cases the hollow beads could be used for the conservation or distribution of in vitro cultures as an alternative to the tested calcium alginate beads (Maruyama et al. 1997). Furthermore, the encapsulation method is also applicable for the construction of artificial seeds as an alternative to conventional calcium alginate beads. In further experiments, the influence of capsule characteristics such as capsule size, wall thickness and density of encapsulated material on germination and plant quality will be investigated. It should also be mentionend that the method of hollow bead production described in this paper yields capsules with varying numbers of propagules (for instance somatic embryos). For several applications, it would be advisable to produce hollow beads containing only one propagule (for instance only one somatic embryo). This can be achieved by optimizing the concentration of propagules in the dropping solution so that most droplets contain only one propagule, and by further research into encapsulation technology.
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