Development of protoplasts of Ulva fasciata (Ulvales, Chlorophyta) for algal seed stock

Yean-Chang Chen and Hsiu-Chuan Shih

Department of Aquaculture,

National Taiwan Ocean University,

Keelung, Taiwan.

 

Email: ycchen@mail.ntou.edu.tw

b0232@ind.ntou.edu.tw

ycchen@ntou66.ntou.edu.tw

　

ABSTRACT

The aim of this study was to isolate and cultivate protoplasts of the green alga Ulva fasciata Delile and subsequently induce them to form a micro-thalli suspension for algal ‘seed stock’. The protoplasts were covered with secreted mucilage following 6 h of culture when viewed with SEM. The mucilage fused to form thick layers during day 1 of culture. Microfibrillar cell walls were deposited into the thick layers of mucilage on the 5^th day of culture. An average of ca. 10% of the freshly isolated protoplasts began to divide at 6-14 days. These protoplasts subsequently developed varied morphologies, depending upon the time of collection during the year. Protoplasts isolated from U. fasciata collected in March to June developed frond thalli or micro-thalli when they were cultured in low or high densities (cells/area), respectively. The micro-thalli suspension was cultured for more than two years at 10-40 µmol photons^.m^-2.s^-1. Frond thalli formed when the suspension was cultivated at 100-160 µmol photons^.m^-2.s^-1. Therefore, the micro-thalli suspension can serve as a ‘seed stock’ of U. fasciata.

 

Key index words: algal seed stock; micro-thalli; cell wall; green alga; protoplast; SEM; sporangium; TEM; Ulva fasciata; zoospore

 

INTRODUCTION

Numerous studies have been conducted over the past decade on edible marine algal protoplasts, such as Enteromorpha spp., Ulva spp., Monostroma spp., Laminaria sp., Porphyra spp., Grateloupia spp., (Ar Gall et al. 1993, Chen 1987, Chen 1998, Chen and Chen 1991, 1993, Chen and Chiang 1994a b, Fujimura et al. 1989, Fujita and Migita 1985, Polner-Fuller and Gibor 1984, 1990, Reddy and Fujita 1991, Reddy et al. 1989, 1992, Saga and Sakai 1984, Saga and Kudo 1989, Waaland et al. 1990). Algal protoplasts were cultivated to form complete macro-thalli. Recent developments in techniques for the large-scale release and isolation of protoplasts offer the opportunity to cultivate edible algae in mass. However, macro-thalli cannot be kept in the laboratory for long periods of time. Laboratory conditions often cannot provide a large enough area having the essential high irradiance and nutrient-replete seawater (Chen 1998). Protoplasts of holdfasts of Monostroma latissimum Wittrock can form filamentous thalli (Chen 1998). These filaments were micro-thalli and could be kept in an incubator for longer periods without losing their ability to develop edible frond-thalli. These “filaments” represent a ‘seed stock’. Therefore, the ability to control the differentiation of algal protoplasts to form a ‘seed stock’ would facilitate the practical application of algal protoplasts in commercial-scale culture.

Ulva protoplasts (Chen and Chen 1991, 1993, Reddy et al. 1989) develop into morphologically-variable thalli. However, such protoplast-derived thalli must be transferred quickly to the sea for further cultivation or death occurs. This is a big problem in the application of protoplast technology to algal culture. Therefore, in this study, we tried to induce protoplasts to form micro-thalli. As with the embryo of a dormant seed, growth and differentiation of protoplast-derived micro-thalli can be suppressed and then induced to produce complete thalli when needed. This ‘seed stock’ culture from protoplast-derived thalli is novel among the Ulvales. It provides the most convenient method for algal mass culture in addition to methods for Monostroma (Chen, 1998).

 

MATERIALS AND METHODS

Fronds of the marine macroalga Ulva fasciata Delile were collected at Keelung, Taiwan, in January to December 1997. Freshly collected algae were wrapped in absorbent paper towels moistened with seawater, sealed in plastic bags and packed in an ice-box for transport to the laboratory of the Department of Aquaculture, National Taiwan Ocean University in Keelung, Taiwan.

The preparation, isolation, and purification of the protoplasts were carried out using the same methods as in our previous studies (Chen 1998, Chen and Chen 1991, 1993, Chen and Chiang 1994a).

 

Preparation of axenic algal materials. Selected pieces of vegetative frond (2cm²) were cleaned in filtered seawater. They were then put in an ultrasonic cleaner (Branson 3200) with two changes of autoclaved seawater containing 1 % KI-I₂ for 5 min each, to remove small animals and epiphytes (Reddy et al. 1989). The pieces were then rinsed several times with autoclaved seawater. Finally they were incubated in 100 mL of autoclaved enriched seawater (Provasoli 1968) containing 10 mL of antibiotic mixture (Polne-Fuller and Gibor 1984) for 24 h at 24℃ under 60 µmol photons^.m^-2.s^-1 and 12:12 L:D photoperiod in a culture room.

 

Isolation of protoplasts. Protoplasts of Ulva fasciata were isolated enzymatically with 4% cellulase Onozuka R-10 and 2 % Macerozyme R-10 (Yakult Honsha Co. Ltd. Japan) in 10 mL of 1.2 M sorbitol solution. All isolation procedures were carried out in the dark (Marchant and Fowke 1977, Cheney et al. 1986, Reddy and Fujita 1991) to promote the sinking of isolated protoplasts (Liu et al. 1992) at 24°C. The yield of freshly isolated protoplasts was counted with a hemacytometer (Bright-Line, improved Neubauer, 0.1 mm deep) under a light microscope (Zeiss, Axioskop). The experiments in this study were performed at least three times.

 

Protoplasts culture and formation of micro-thalli. Purified protoplasts were cultured in Provasoli's enriched seawater (PES) medium containing 0.84 M mannitol immediately after isolation (Chen and Chen 1991, Chen and Chiang 1994a). For inducing protoplasts to develop micro-thalli, they were diluted or concentrated into eight groups of culture densities (protoplasts/bottom surface area), such as 10⁵, 5x10⁵, 10⁶, 5x10⁶, 10⁷, 5x10⁷, 10⁸ and 5x10⁸ protoplasts in 10 mL of PES medium in eight flasks (Falcon, 40-mL, 35x65mm² of bottom area) separately for further culture. Since not all protoplasts survived during culture, the final densities were determined at day 14 using an inverted microscope (Zeiss, Axiovert 135). All protoplast cultures were incubated under a photoperiod of 12:12 L:D and 160 µmol photons^.m^-2.s^-1 at 24°C. The regenerated protoplasts were cultured in 0.84 M mannitol-PES medium for 5 days. The medium was then replaced into a pure PES medium for further cultivation under the conditions mentioned above (Chen and Chiang 1994a b). An inverted microscope (Zeiss, Axiovert 135) with camera equipment (MC 80DX) was used at intervals to observe and photograph the regenerating protoplasts at different stages, and to determine the division (survival) rate of protoplasts.

 

Protoplast cell wall formation. Calcofluor White ST (Sigma chemical Co. St. Louis, Missouri), a fluorescent brightener reagent was used to investigate the course of cell wall resynthesis in the cultured protoplasts (Galbraith 1981, Roberts et al. 1982, Chen and Chen 1993). About 0.01% of Calcofluor White (w/v) was added to a culture of freshly isolated protoplasts in 0.84 M mannitol-PES medium. The culture was examined hourly on an inverted microscope with fluorescent equipment (Zeiss, Axiovert 135, Excite filters: Lp510-Kp560; Chromatic beam splitters: Ft580; Barrier filter Lp590) to determine if the protoplasts had regenerated new walls. (For additional details, see Chen and Chen 1993).

 

Culture of micro-thalli and formation of frond thalli. The regenerated micro-thalli were moved from the flasks (Falcon, 40-mL) into glass flasks (500 mL) with cotton plugs and with 250 mL of PES medium. They were cultivated under a photoperiod of 12:12 L:D and 10, 20, 30, 40, 60, 80, 100 and 160 µmol photons^.m^-2.s^-1 at 24°C, separately.

 

Plantlet culture (The sowing of the algal seed stock on nylon rope). The micro-thalli suspensions were moved from the flasks into glass aquarium jars (40x30x20 cm) with fine nylon ropes (1 mm in diameter) at the bottoms and with 12 L of PES medium, under a photoperiod of 12:12 L:D and 40, 60, 80, 100 and 160 µmol photons^.m^-2.s^-1 at 24°C for further cultivation.

 

Fixation of the culture materials for electron microscopy studies. Some purified protoplasts were fixed immediately (as 0-h old protoplasts), whereas some culture groups, as described above, such as 6 h-old and 1 to 16 day-old protoplasts were transferred into 15-mL centrifuge tubes followed by separate fixation. Those protoplasts were collected using gentle centrifugation (Hermle Z320) at 120 xg for 20 min at 24 °C. At 0 and 6-h old, the protoplasts were fixed at 4℃ for 1 h in PES medium containing 2 % glutaraldehyde, and 0.5 M sucrose with gentle shaking. Subsequently, these protoplasts were collected by centrifuging at 300 xg for 10 min. The pellets were then re-fixed with 5 % glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 6.8) containing 10 mM CaCl₂ and 0.2 M sucrose for 1 h at 4℃. They were then rinsed with a 0.1 M sodium cacodylate buffer containing 10 mM CaCl₂ four times, with a sucrose concentration successively reduced to 0.05 M. This treatment was followed by two rinses in a pure (sucrose-free) 0.1 M sodium cacodylate buffer containing 10 mM CaCl₂. Protoplasts 24 h or older, were fixed as those 0 h and 6 h, except that the duration of fixation was extended to 2 h and 0.2 M sucrose was used in the first fixative. Post-fixation was done with 2 % OsO₄ in 0.1 M sodium cacodylate buffer containing 10 mM CaCl₂ for 1 h at 4℃.

Thereafter, all materials were rinsed four times with a 0.1 M sodium cacodylate buffer containing 10 mM CaCl₂, 3x with aqueous ethanol (50 %), and gradually dehydrated in ethanol (50, 70, 85, 95 and 100%). During this dehydration, 0, 6, 24, 48, 72, 96 and 144 h-old protoplasts generally formed small clumps. These clumps and others (7-16 days-old protoplasts) were prepared for transmission electron microscopy (TEM) by rinsing in propylene oxide (3x, 30 min each), followed by infiltration in propylene oxide-Spurr's resin at a decreasing ratio from 2:1 (2 parts propylene oxide :1 part Spurr's resin) to 1:1 each for 4 h. Samples were then suspended in pure Spurr's resin for two days at 4℃ in darkness before embedding in Spurr's resin (Spurr 1969). The thin-sections were stained with uranyl acetate and lead citrate according to Smith and Croft (1991).

Protoplast clumps (0, 6, 24, 48, 72, 96, 120 and 144-h old protoplasts) and early stage thalli were also prepared for scanning electron microscopy (SEM). Protoplast clumps were suspended in pure ethanol for two days with gentle shaking (two cycles per min) on a rotary disc (Firstek, Taiwan). This procedure freed protoplasts from clumps. Thereafter, the protoplast- and early stage thalli-ethanol suspension were gently dropped onto specimen holders separately; and then dried with a critical-point-drying machine (Hitachi-HCP-1). Finally, they were coated on an ion coater (Joel, JCF-1100E) for 220s.

 

RESULTS

Protoplast yields varied from 1x10⁶ to 3.13x10⁸ protoplasts ^. g^-1 (FW). As shown in Table 1, the varied yields were due to collection of wild materials in different months. However, the average yield was 3.23x10⁷ protoplasts ^. g^-1. It was found that 80.5% of the freshly isolated protoplasts had secreted a cell wall at day 4 of culture in 0.84M of mannitol-PES at 24℃, regardless of the irradiance.

The ultrastructure of freshly isolated protoplasts (Fig. 1·a) showed that chloroplasts were the predominant organelles. The simple internally-suspended pyrenoids (Fig. 1·a) were normally closely surrounded by a cap of reserve material. Typically, the cell membrane of the protoplast was smooth (Fig. 1·a). With SEM it was found that the freshly isolated protoplasts were spherical and had already secreted a little amorphous mucilage on the surface. Mucilage was initially secreted onto the protoplast’s surface in ordered, serrated rows, and it covered the entire protoplast (Fig. 1·b) by 6 h. Subsequently the mucilage fused to form thick, bulge-like layers on the surface of the protoplast on the 24^th h of cultivation. As the protoplasts aged, the mucilage layers thickened. The protoplast expanded its appearance from a spherical to an oval shape at day 6 of the culture.

Protoplasts secreted a microfibrillar cell wall (ca. 1.7-2μm in thickness) (Fig. 1·c) within these thick layers of mucilage by day 4-6 of cultivation. The new regenerated walls were composed of a thin compact layer of primary wall and thick loose layers of secondary walls beneath the former (Fig. 1·c). About 5-30 % (the average was ca. 10 %) of freshly isolated protoplasts that formed new walls (Figs. 1·d, 1·e,1·f) had divided by 6-14 days in culture. Subsequently, they developed various morphologies, or developed into frondose thalli (Table 1), in PES at 100-160 µmol photons^.m^-2.s^-1 and a photoperiod of 12:12 L:D at 24ºC at further cultivation.

Protoplasts from the material collected in December-February developed into tubular thalli and then into normal fronds. Such protoplasts formed neither micro-thalli nor sporangia. Their developmental patterns were the simplest of all the collections during the year (Table 1). Protoplasts from material collected in late August-November  followed two different developmental patterns. About 60 % of the viable protoplasts developed into tubular thalli and then into frondose thalli. The other 40 % of the viable protoplasts developed sporangia (Figs. 2·a, 2·b) at day 14-20 of culture. Zoospores (Figs. 2·a, 2·c, 2·d) were released from the sporangia within 24-30 days of culture in PES under 100-160 µmol photons^.m^-2.s^-1 at 24ºC. The biflagellate zoospores (Fig. 2·c) were initially suspended in the sporangium, and using amoeboid movement (Fig. 2·d), they escaped through pores of sporangium in a few seconds. However, the release was not synchronous. It took about 3-7 days for all of the sporangia to release their zoospores.

Protoplasts isolated from material collected in late August-November, never developed into micro-thalli regardless of the culture densities.

In contrast to protoplasts from material collected in late August-November, those that had been collected in March-June showed other developmental patterns. They did not develop sporangia; rather, they developed into micro-thalli and/or frondose thalli depending on the culture densities but regardless of the irradiance. When incubated at densities of less than 25^.mm^-2 (total viable protoplasts were ca. 10⁶ protoplasts in 10 mL of PES medium in 40-mL flask), they developed into frondose thalli. Protoplasts cultured at densities of 25 to 125^.mm^-2 (10⁶-5x10⁶ protoplasts in 10 mL of PES medium in 40-mL flask) developed into either frondose thalli or micro-thalli. Protoplasts cultivated at densities above 125^.mm^-2 (i.e. more than 5x10⁶ protoplasts in 10 mL of PES), grew on the flask bottom. They subsequently developed micro-thalli (Fig. 2·e). However, the development of micro-thalli was affected by irradiance. When they were incubated at 10-40 µmol photons^.m^-2.s^-1, they remained micro-thalli in form. At 40-80 µmol photons^.m^-2.s^-1, the micro-thalli grew larger, and then fragmented into numerous smaller micro-thalli, subsequently developing into a ‘micro-thalli suspension’ (Fig. 2·e) in the culture flasks. When incubated above 100 µmol photons^.m^-2.s^-1, they expanded in volume to form tubular (or spindle), saccate (or spherical) and irregular thalli (Fig. 2·f) that were monostromatic. However, they did not fragment into smaller structures. Some of these further developed into frondose thalli. Others became rather huge saccate thalli (Fig. 3·a) with sizes approximate of 1.4 mm². Subsequently, several tubular and frondose thalli (Fig. 3·b) developed from these saccate thalli. Finally, tufted fronds formed (Fig. 3·c).

The micro-thalli suspensions were stored in an incubator at 10-40 µmol photons^.m^-2.s^-1 and a photoperiod of 12:12 L:D at 24℃. Without renewing the medium in two years, the micro-thalli retained their ability to develop into  frondose thalli. After 7 days of sowing, the micro-thalli would attach firmly onto the nylon rope, and they developed into spherical and tubular thalli (Fig. 3·d), with many thalli (Fig. 3·e) growing from the rope. Thus, this suspension can function as a ‘seed stock’ for commercial culture.

 

DISCUSSION

The variation in the yield of protoplasts with the month of collection may be due to the different physiological conditions of the materials. However, the average yield (3.23x10⁷ cells ^.g^-1) from this study was consistent with our previous observations (Chen and Chen 1991, 1993) and better than the yields (1-4x10⁶ cells ^.g^-1) obtained by Reddy et al. (1989). This study also confirmed that green algal protoplasts secreted mucilage on the surface before depositing the structured microfibrillar walls, as in the red algal protoplasts (Chen and Chiang 1995). The organelles involved in the cell wall resynthesis of the green algal protoplasts require further study.

Various species of Ulva protoplasts developed different morphologies. Protoplasts of U. fasciata and U. conglobata Kjellm developed into spindle and frondose thalli and sporangia. Protoplasts of U. pertusa Kjellm formed spherical and saccate thalli and sporangia (Reddy et al. 1989). Protoplasts of U. lactuca L. regenerated to filamentous and discoid thalli (Huang et al. 1996). In this study, we also found that the protoplasts of U. fasciata developed into more numerous patterns of algal thalli, such as sporangia, micro-thalli, saccate (or spherical) thalli, tubular (or spindle), irregular and fronds. Furthermore, we found that the protoplast-derived sporangia subsequently released zoospores with the characteristic flagella. We could not observe the fusion of those biflagellate-cells and therefore could not ascertain if they were zoospores or gametes. However, in this study, we think that the motile biflagellate-cells were zoospores, as with the study of Reddy et al. (1989).

An axenic laboratory strain of Ulva lactuca L. showed polymorphism while incubated with various bacterial floras. U. lactuca never grew normally without microflora (Provasoli and Pintner 1980). Fujimura et al. (1989) also reported that protoplasts of Ulva pertusa incubated in an environment free of accompanying microflora, regenerated into irregular long, tubular and tufted uniseriate thalli. In this study, the protoplasts of U. fasciata grew to various morphologies in early stages, and subsequently developed into normal frondose thalli. This may be due to a response to axenic culture or absence of morphogenetic substance in the early stages

Polymorphism is also widespread in habitats with similar physical condition (Provasoli and Pintner 1980). In such case, many factors may underlie variability, and the culture density may be one of them. In this study, U. fasciata protoplasts form micro-thalli in order to adapt to the crowded environment in such higher culture density. Protoplasts of Ulva sp. regenerated into calli when the protoplasts were cultured in agar media (Polne-Fuller and Gibor 1987). The agar medium may be other type of crowded condition to the algal protoplasts. Mori and Mizuta (1988) also reported that the higher protoplast density reduced the rate of complete development of Enteromorpha.

Our previous study (Chen 1998) isolated protoplasts from holdfasts of Monostroma latissimum, and subsequently induced the protoplasts to form microscopic filaments for ‘seed stock’. However, this study failed to obtain the protoplasts from holdfasts of Ulva fasciata. It seems that the walls of the holdfast-cells of U. fasciata resisted degradation with the enzyme mixture used in this study. Therefore, the composition of the walls of holdfast-cells of U. fasciata require clarification.

Although no filaments were induced from the protoplasts of U. fasciata, the microscopic protoplast-derived micro-thalli of U. fasciata can be induced to form and differentiate again by simply manipulating the culture conditions. This is potentially a very convenient means of storage and micropropagation for algal commercial culture.

 

ACKNOWLEDGEMENT

Financial support from NSC grants 88-2313-B-019-04; 89-2313-B-019-032 and COA grants 88-AST-1.4-FID-02(12-2); 89-AST-1.2-FID-05(06) of the Republic of China is greatly appreciated.

 

REFERENCES

Ar Gall, E., Chiang, Y. M., & Kloareg, B. 1993. Isolation and regeneration of protoplasts from Porphyra dentata and Porphyra crispata. Eur. J. Phycol. 28:277-283.

Chen, C. S. & Chen, Y. C. 1991. Isolation and regeneration of protoplasts from the green alga Ulva fasciata Delile. Proceedings of the National Science Council, ROC Part B: Life Science 15:244-250.

Chen, L.C-M. 1987. Protoplast morphogenesis of Porphyra leucostica in culture. Bot. Mar. 30:399-403.

Chen, Y. C. 1998. Development of protoplasts from holdfasts and vegetative thalli of Monostroma latissimum (Chlorophyta, Monostromataceae) for algal seed stock. J. Phycol. 34:1075-1081.

Chen, Y. C. & Chen, C. S. 1993. Use of fluorescent staining to monitor the temporal pattern of cell wall resynthesis in Ulva fasciata (Chlorophyta: Ulvales, Ulvaceae) protoplasts. Jp. J. Phycol. 41:237-242.

Chen, Y. C. & Chiang, Y. M. 1994a. Isolation and regeneration of protoplasts of Monostroma latissimum Wittrock (Monostromataceae, Chlorophyta). Bot. Bull. Acad. Sin. 35:45-51.

─── 1994b. Development of protoplasts from Grateloupia sparsa and G. filicina (Halymeniaceae, Rhodophyta). Bot. Mar. 37:361-366.

─── 1995. Ultrastructure of cell wall regeneration from isolated protoplasts of Grateloupia sparsa (Halymeniaceae, Rhodophyta). Bot. Mar. 38:393-399.

Cheney, D. P., Mar, E., Saga, N. & van der Meer J. (1986) Protoplast isolation and cell division in the agar-producing seaweed Gracilaria (Rhodophyta). J. Phycol. 22: 238-243.

Fujimura, T., Kawai, T., Shiga, M., Kajiwara, T. & Hatanaka, A. 1989. Regeneration of protoplasts into complete thalli in the marine green alga Ulva pertusa. Nippon Suisan Gakkaishi 55(8): 1353-1359.

Fujita, Y. & Migita, S. 1985. Isolation and culture of protoplasts from some seaweeds. Bull. Fac. Fish. Nagasaki Univ. 57:39-45.

Galbraith, D. W. 1981. Microfluorimetric quantitation of cellulose biosynthesis by plant protoplasts using Calcofluor White. Physiol. Plant. 53: 111-116.

Huang, X., Weber, J. C., Hinson, T. K., Mathieson, A. C. & Minocha, S. C. 1996. Transient expression of the GUS reporter gene in the protoplasts and partially digested cells of Ulva lactuca L. (Chlorophyta). Bot. Mar. 39:467-474.

Liu, Q.Y., Chen, L. C-M, & Taylor, A. R. A. 1992. Ultrastructure of cell wall regeneration by isolated protoplasts of Palmaria palmata (Rhodophyta). Bot. Mar. 35:21-33.

Marchant, H. J. & Fowke, L.C. 1977 Preparation, culture, and regeneration of protoplasts from filamentous green algae. Can. J. Bot. 55: 3080-3086.

Mori, T. & Mizuta, S. 1988. Development of the marine Ulvalean alga Enteromorpha intestinalis from its protoplasts. Rep. Usa mar. Biol. Inst., Kochi Univ. 10:93-98.

Polne-Fuller, M. & Gibor, A. 1984. Developmental studies in Porphyra I. blade differentiation in Porphyra perforata as expressed by morphology, enzymatic digestion, and protoplast regeneration. J. Phycol. 20:609-616.

─── 1987. Calluses and callus-like growth in seaweeds: Induction and culture. Hydrobiologia 151/152: 131-138.

─── 1990. Developmental studied in Porphyra (Rhodophyta) III. Effect of culture conditions on wall regeneration and differentiation of protoplasts. J. Phycol. 20:674-682.

Provasoli, L. 1968. Media and products for the cultivation of marine algae. In Watanabe, A. & Hattori, A. [Eds.] Culture and collections of algae. Jap. Soc. Plant Physiol., Tokyo: 63-75.

Provasoli, L. & Pintner, I. J. 1980. Bacteria induced polymorphism in an axenic laboratory strain of Ulva lactuca (Chlorophyceae). J. Phycol. 16:196-201.

Reddy, C. R. K., Migita, S. & Fujita, Y. 1989. Protoplast isolation and regeneration of three species of Ulva in axenic culture. Bot. Mar. 32: 483-490.

Reddy, C. R. K. & Fujita, Y. 1991. Regeneration of plantlets from Enteromorpha (Ulvales, Chlorophyta) protoplasts in axenic culture. J. Appl. Phycol. 3:265-275.

Reddy, C. R. K., Iima, M. & Fujita, Y. 1992. Isolation of fast-growing and morphologically different strains through intergeneric protoplast fusion of Ulva and Enteromorpha. (Ulvales, Chlorophyta). J. Phycol. 4:57-65.

Roberts, E., Seagull, R.W., Haigler, C. H. & Brown, R. M. JR. 1982. Alteration of cellulose microfibril formation in eukaryotic cells: Calcofluor White interferes with microfibril assembly and orientation in Oocystis apiculata. Protoplasma 113: 1-9.

Saga, N. & Sakai, Y. 1984. Isolation of protoplasts from Laminaria and Porphyra. Bull. Jap. Soc. Sci. Fish. 50(6):10-85.

Saga, N. & Kudo, T. 1989. Isolation and culture of protoplasts from the marine green alga Monostroma angicava. J. Appl. Phycol. 1: 25-30.

Smith, M. &. Croft. S. 1991. Embedding and thin section preparation. In: Harris, J. R. [Ed.] Electron Microscopy in Biology. IRL press, Oxford New York Tokyo. pp. 17-37.

Spurr, A. R. 1969. A low viscosity epoxy resin embedding medium for electron microscopy. J. Ultrastruct. Res. 26:31-43.

Waaland, J. R., Dickson, L. G. & Watson, B. A. 1990. Protoplast isolation and regeneration in the marine red alga Porphyra nereocystis. Planta 181:522-528.

FIGURE LEGENDS

Figs. 1·a-c: Ultrastructure of the protoplasts and the regenerated cell walls.

Fig. 1·a. TEM section of a freshly isolated protoplast of Ulva fasciata. Ch: chloroplasts, P: pyrenoid, Mi: mitochondrion, cell membrane (arrowheads). Fig. 1·b. SEM observation of a 6-h old spherical protoplast that had secreted plenty of mucilage vertically and horizontally arranged in serrate rows on its surface. Fig. 1·c. TEM observation of a 5-day-old protoplast with its complete thin primary wall (Pw) and a thick secondary wall (Sw) beneath the former. The new walls are ca. 1.7-2μm in thickness. Nu: nucleus.

Figs. 1·d-f: The division of protoplasts.

Fig. 1·d. TEM observation of a 2 dividing cell protoplast at day 10 of cultivation. Cw: new cell wall. Fig. 1·e. Light microscope observation of 2- and 4-cell-stage protoplast at day 14. Fig. 1·f. TEM observation of a 4-cell-stage protoplast at day 14.

Figs. 2·a-d: Structure of the protoplast-derived sporangia and their spores.

Fig. 2·a. TEM observation of the zoospores (Z) with flagella (FL) suspended inside the sporangium (Sp). The rim (arrowheads) of the sporangium was clearly defined by the electron dense lines. Fig. 2·b. SEM observation, an oval sporangium (Sp) had produced plenty of mucilage (arrowhead) for attachment to the substrate. A pore (arrow) was at one end of the sporangium for the zoospores being released. Fig. 2·c. TEM observation, the zoospores were characterized by the ellipse shape and the two flagellates (arrows). Fig. 2·d. Light microscope observation, a zoospore (double arrowheads) moved through the pore of the sporangium (Sp). E: an empty sporangium.

 

Figs. 2·e-f: Development of the micro-thalli.

Fig. 2·e. Micro-thalli suspension observed under the light microscope. Fig. 2·f. SEM observation, different forms of early developing tubular (T) (or spindle), saccate (S) (or spherical) and irregular thalli (I).

 

Figs. 3·a-e: Development of the macro-thalli, observed under the light microscope.

Fig. 3·a. A saccate thallus with rough and uneven surface, one cell layer thick. Fig. 3·b. A saccate thallus (S) producing tubular thallus (T) and a frond (F). Fig. 3·c. Formation of multiple fronds and tubular thalli from a saccate thallus. Fig. 3·d. Spherical (St) and tubular thalli (T) attached to the nylon rope (N) 7 days after sowing. Fig. 3·e. Many thalli growing on the nylon rope.

Table 1: The yields and developmental patterns of protoplasts of Ulva fasciata. Algal materials for the protoplast isolation were collected at different times during the year. Protoplasts were cultured in 0.84 M mannitol-PES for 5 days, then in pure PES for growth at 100-160 µmol photons^.m^-2.s^-1 and a photoperiod of 12:12 L:D at 24°C.

 

Months	Yields (protoplasts .g-1 FW)	Developmental patterns
January	2±1x107   (4)*	Tubular and frondose thalli
February	2±1x107   (4)*	Tubular and frondose thalli
March	1.17x108±4.08 x106   (6)*	Micro-thalli**, saccate, tubular, irregular and frondose thalli
April	5.6±4.6x106   (6)*	Micro-thalli**, saccate, tubular, irregular and frondose thalli
May	1.235±1.025x108   (6)*	Micro-thalli**, saccate, tubular, irregular and frondose thalli
June	1.025x107±4.55x106   (3)*	Micro-thalli**, saccate, tubular, irregular frondose thalli
July-middle August	Not tested or collected***	-----------------------------
Late August	2.845x108±2.85x107   (3)*	Tubular and frondose thalli, sporangia
September	6±3.54x106   (4)*	Tubular and frondose thalli, sporangia
October	2.9±1.27x106   (4)*	Tubular and frondose thalli, sporangia
November	1x106±9x105   (3)*	Tubular and frondose thalli, sporangia
December	2.9±2.1x106   (3)*	Tubular and frondose thalli

*The sample size.

**The micro-thalli were formed by culturing protoplasts at a high density of 125 protoplasts^.mm^-2 surface area of flask bottom.

***Mostly the algae died during the hot summer days, therefore, no material was collected during these months.