Plant Cell Reports
Plant Cell Reports (1995) 14:571-574
9 Springer-Verlag1995
Protoplast isolation in the marine brown alga Dictyopteris prolifera (Dictyotales) T. Fujimura 1, T. Kawai ~, T. Kajiwara 2, and Y. Ishida 3 1 Laboratory of Marine Bioteehnology, Shiono Koryo Kaisha, Ltd., Niitaka, Yodogawa-ku, Osaka 532, Japan 2 Department of Biological Chemistry, Faculty of Agriculture, Yamaguchi University, Yoshida, Yamaguchi 753, Japan 3 Department of Fisheries, Faculty of Agriculture, Kyoto University, Oiwake-cho, Kitashirakawa, Sakyo-ku, Kyoto 606, Japan Received 7 May 1993/Revised version received 15 November 1994 Communicated by A. Komamine
Abstract Protoplasts
were isolated from thalli of prolifera using a mixture of crude enzymes from vicera of live oysters (Crassostrea gigas) and the following commercial enzymes: an abalone enzyme, cellulase, polygalacturonase and hemicellulase. The enzyme mixtures produced up to 3.3 x 107 cells per 1 g of tissue fresh weight. The conversion to protoplasts of the cells was about 100% using the oyster enzyme or the abalone enzyme alone. The optimum pH for protoplast isolation was 6.0 and 20 hours were required for conversion to protoplasts.
Dictyopteris
Key words:
C m s s o s ~ ' e a ffZ~as -
pro~ra
Marine brown alga -
-
D1VO~o/TtelTir Protoplast-
Single cell Introduction Ever since Cocking work in 1960, plant protoplasts were adapted in various fields of plant science as useful experimental material (Cocking 1972). Protoplasts have been a useful research tool in physiology, morphology, biochemistry, pathology, cell biology, cell fusion, somatic hybridization, and genetic manipulation (Pilet 1985; Fowke and Constabel 1985; Puite et al. 1988). Researchers have been continuously progressing to advance the development of more efficient techniques for plant protoplasts research (Bajaj 1989a,b). On the other hand, in seaweeds protoplast research lagges far behind that of higher plants. Recently, the t e c h niques for isolation of protoplasts from seaweeds have been developing and advancing as described in various reviews (Cheney 1984, 1986; P o l n e Correspondence to: T. Fujimura
FuUer and Gibor 1987; Polne-Fuller 1988). Millner et al. (1979) reported the first isolation of protop-lasts from the marine green alga .Enteromo_rp,ha ~tes/z~ab3- by commercial enzymatic degradation. Since then, protoplasts from many green algae have been isolated using commercial cell wall-degrading enzymes commonly employed for higher plants (Cheney 1986). Brown algal protoplasts were produced only when feeder enzymes from marine animal guts which were previously fed on these specific seaweed were used for isolation procedures. Such enzymes were obtained from sea urchin (Saga and Sakai 1984), abalone (Saga et al. 1986), sea hare (Kloareg et al. 1987), and sea snail (Wu 1988). The induction of digestive enzymes in grazers for the p~oduction of brown algal protoplasts was suggested by P o l n e Fuller and Gibor (1987), since marine invertebrates feeding on benthic algae must contain the necessary digestive enzymes to dissolve brown algal polysaccharides in their diet. Kajiwara et al. (1988) selected enzyme systems consisting of marine feeder enzymes mixed with commercial ones, and further demonstrated the isolation of protoplasts from the brown algae Dictyotaceae. However, their isolation produced highly variable yields of protoplasts, possibly due to the purification levels of these crude biological enzymes. This paper reports the formation of protoplasts from the marine brown alga Z). pl'o/zi~ra via through an intermediate single ceils stage. We also report an effective method to isolate a great number of the protoplasts by using an enzyme mixture of partially purified oyster enzyme with a variety commercial ones. We then discuss some features of cell wall composition as revealed by the enzymatic varied effects of different enzymes on the degradation of cell walls.
572 Materials and methods M~e matr D. flfofffcra was collected from the coast of Shimonoseki and from the southern Osaka Bay, Japan. Live oysters were obtained from the beach of Yamaguchi, Japan. E~es. Some enzymes normally used for digesting higher plant cell walls were used; an Abalone Acetone Powder (an abalone enzyme) was purchased from Sigma Chemical Co. (St. Louis, USA), and Cellulase Onozuka R - 1 0 and Macerozyme R 10 were from Yakult Honsha Co., Ltd. (Tokyo, Japan), Driselase was from Kyowa Hakko Kogyo Co., Ltd. (Tokyo, Japan), and Bigatase M and Sumizyme X were fTam Kyowa Chemical Products CO., Ltd. (Osaka, Japan).
P~para~'on o f an oyster crude e~zj,z~e. The crude enzyme solution extracted from oysters was prepared by the following method. 100 g of entrails were homogenized with 100 ml of 50 mM sodium phosphate buffer (pH 7.0) containing seawater instead of water and filtered through some layers of gauze, then centrifuged at 12,000 x g for 10 rain. The super'natant was precipitated with 90 % saturation of ammonium sulfate for 3 h with stirring and then centrifuged at 12,000 x g for 15 rain. The precipitate was dissolved in a minimal amount of the same phosphate buffer (30-35 mi) and dialyzed for 12 h against several changes of the same buffer. The dialyzed liquid was brought up to 50 mi with 1.0 M mannitol solution in the same buffer and adjusted to pH 6.0. One ml of the solution (straight oyster enzyme) was used as an oyster crude enzyme for one test to isolate single cells and further to determine the protoplasts. All procedures were carried out at 5 *C. Prepara~,;on o f co~u~erHM e ~ T m c solul~bos. The commercial abalone digestive enzyme was dissolved into 5 % (w/v) solution in the buffer. The commercial enzymes were dissolved at 2 % (w/v) concentrations and then mixed equally into a solution. Ten ml aliquots of mixed enzyme solutions were tested on chopped seaweed tissues. Isolation o f sfngle c~ffs. After being washed three times with sterilized seawater, the thalli were cut into small pieces about 5 mm square. The pieces (0.5 g fresh weight) were moderately shaken (30-35 strokes/rain) at 25 ~ for 10 rain in 0.5 % papain solution containing 100 mM Tris-HC1 buffer (pH 7.4) and 1.0 M mannitol. They were transferred to other 1.0 M mannitoI solution (.pH 6.0) and shaken for 10 rain, and then 10 ml enzyme solution was added onto them to digest the cell walls. Single cells were released from the tissue after incubation in the enzymes for 15-20 h at 25 *C with moderate shaking (2530 strokes/rain). The released cells were collected with two kinds of filters (stainless mesh, 88-; nylon mesh, 40 tun-opening). The filtrates were washed three times in Tris-HC1 buffer (pH 8.0) contzlning 1.0 M mannitol, and centrifuged at 500 x g for 5 rain. The isolated single cells were counted with a hemocytometer.
Isola~bn o[protoplasts. Protoplasts, whose cell walls were completely removed by the enzymes, remained mixed with the single cells. They were microscopically counted on the basis of their morphological difference. FinaUy, the cell wall digestion was confirmed by the absence of fluorescent brightener, Calcofluor White ST (Sigma Chemical Co.) staining as previously described (Fujimura et al. 1989).
Rcsul~
fsola[fon of Siafflc cells When applied to chopped D. prolAFcra tissues, the variou~ enzyme ~ e s resulted in varying yields of single cells, and/or protoplasts. Sphere like single cells which were released from tissues are known as spheroplasts. They are on the way to convert themselves to the protoplasm if additional and suitable enzymes digesting the cell wails should be present- Table 1 showed that the commercial enzymes had no effects on formation of single cells, although those enzymes were shown to digest cell walls of many higher plants (Power and Chapman 1985) and some marine green algae (Fujimura et al. 1989). However, both the oyster, and abalone enzymes, and these two in mixtures, with the commercial enzymes, produced great number of single ceils (nearly 10 7 cells per 1 g of fresh weight as shown in Table 1). It suggested that some enzymes derived from algal grazers are more specialized than the available commercial enzymes in digesting brown algal cell walls, indicating that the brown algal ceil wails are composed of polysaccharides which differed from ceil walls of green algae or higher plants.
Isolation of l~otoplasts All of the isolated cells which were produced by the combined enzyme mixtures, consisting of the oyster enzyme, the abalone enzyme, and the commercial enzymes, were walless protoplasts (Table 1). Oyster enzyme, or abalne enzyme alone,
Table 1. Effect of different enzyme mixtures on isolation of single cells and protoplasts from D. prolifera. Enzyme solution Commercial enzymes Oyster enzyme Abalone powder Oyster enzyme + Comm. enzymes Abalone powder + Comm. enzymes Oyster enzyme + Abalone powder + Comm. enzymes 'a'Cells/g fresh weight.
Total number of isolated cells a 9.4 1.7 4.9 2.2 3.3
0 X 106 X 107 X 107 x 107 X 107
Protoplasts (%)
Walled single cells (%)
0 0 0 3 91 100
0 100 100 97 9 0
573 produced not protoplasts but walled single cells. The combination of enzymes produced a large number of the protopast (3.3 x 10 7 cells per 1 g of fresh weight) where were almost 100 % of the formation. The isolated protoplasts were completely spherical and 9-20 pin in diameter (Fig. 1). The abalone enzyme in combination with the ordinary enzymes produced much higher protoplasts ( 9 1 % yield) than the oyster enzyme mixed with the commercial enzymes did (3 %). This indicates that the abalone enzymes really contributed to the degradation of the intercellular matrix. However, the final addition of the oyster enzymes to the above enzyme mixtures resulted in a reliable and effective method for isolation of protoplasts of this brown alga.
The OpSmum Incuba~'on Time for the Isolation o f Protoplasm Table 3 presents the optimum incubation time for this method of protoplast isolation. Although, reasonable numbers of protoplasts were released after 3 h of incubation in the enzyme mixture (1.1 x 10 6 ceils per 1 g of fresh weight) the. yield increased in time and peaked at 20 h in enzyme treatment. Table 3. Effect of incubation time in the enzyme mixture on isolation of protoplasts from fresh tissue of D. prolifera. Time (h)
Protoplast number a
0 3 6 10 15 20 25 a Cells/gfresh weight.
0 1.1 X 10 6 4.0• 10 6 7.3 X 10 6 1.5 X 10 7 3.0 X 10 7 1.1 X 10 7
Discussion
Fig.1. Newly isolated protoplasts from
19.
prolifera. Scale bar=20 pm. The O p ~ z o ~ Protoplasm
pH
for the Isola[zbn of
The effect of p H on the protopast isolation was tested using the most effective enzyme mixture presented in Table 1. This enzyme ~ e had the highest cumulative activity for protoplast isolation at p H between 4.0-8.0 with an optimum pH around 6.0, as shown in Table 2. Table 2. Effect ofpH in the enzyme mixture on isolation of protoplasts from fresh tissue of D. prolifera. pH 4.0 5.0 6.0 7.0 8.0 a Cells/gfresh weight.
Protoplast number a 4.9 9.8 2.8 6.5 1.3
• x x • •
10 6 10 6 10 7 l0 6 10 6
The isolation of protoplasts from D. pro/ifera was exnmined by various combinations of different enzyme solutions (Table 1). Crude oyster enzyme solution in combination with the commercial enzymes, Cellulase Onozuka R - 1 0 , Macerozyme R - 1 0 , Driselase, Bigalase M, Sumizyme X, and Abalone Acetone Powder, released pure protoplast suspensions in a high yields. The oyster enzymes and abalone powder were essential for protoplast release from fresh tissues of D. p:olKem. But, none of the individual enzymes tested alone produced protoplast in different concentrations to evaluate their efficiency. This means the single enzymes do not have enough for an ability completely to decompose the cell walls in /9. prolzYera. Therefore, this suggests that the cell walls contain the polysaccharides which are degraded by the various enzymes used to isolate protoplasts. While, these marine herbivore enzymes can specialize in the digestion of characteristic polysaccharides, such as alglnate and fucoidan, which compose the cell walls of marine brown algae (Onishi et al. 1985; Yamaguchi et al. 1989). Thus, they are expected to contain brown algal polysaccharide-degrading enzymes such as alginate lyase and fucoidanase. The abalone powder from crude abalone entrails seem to also contain B-glucuronidase and sulfatase. The numbers of the
574 protoplast yielded successfully in this study were greater than those harvested from the marine green alga Ulva pen~sa (Fujimura et al. 1989). While, the sizes of 19]ct_-fopteliS protoplasts were smaller than that of the green alga. The pH was also important in effecting the yields of protoplasts. Generally, the pH optimum of the commercial cell wall-degrading enzymes, namely cellulase, polygalacturonase and h e m i cellulase, is known to be 4.0-7.0, whereas that of alginate lyases produced from marine herbivores is 7.0-8.0 (Franssen and Jeuniaux 1965). Fitzsimons and Weyers (1985) described the effect of pH on protease activity present in Cellulase Onozuka R I0, through the range of 6.0-10.0 (pH optimum ca. 8.0). From these reports, it is suggested that the commercial enzymes rather than the alginate-lyase greatly contributed to the degradation of the brown algal cell wall at the pH optimum, and that the proteases in the enzymes did not function. Furthermore, time was an important factor in increasing the yields. However, the viability of the isolated protoplasts decreased with the longer incubation in the crude marine herbivore enzymes. This can be attributed to various toxic substances and contaminants which may be present in the enzyme mixtures such as proteases, lipases, and other enzymes, phenolics, and salts (Evans and Bravo 1983). Our results show that the cell wails of D. p:off:c:a are composed of complex different constituted system, cellulose and non-cellulosic polysaccharides. The cell wall composition o f / 9 . pla~iog~-amma has been reported to contain fucoidan and alginate (Percival et al. 1981). The non-cellulosic polysaccharides such as alghaate and fucoidan are characteristic to cell walls of brown algae. The detailed complexity of these wails is not known. It is thus not surprising that mixtures of a large number of enzymes were required to completely remove the cell wall. Therefore, the oyster and abalone digestive enzymes are most likely more complex than just alginases and fucanases. AcA~7owledgemenls. We wish to thank Prof. A. Cribor and Dr. M. Polne-Fuller, University of California at Santa Barbara, USA for valuable discussion and critical reading of the manuscript. We are also grateful to Prof. H. Fukui, Kagawa University, Japan and Prof. J. Sekiya, Kyoto University, Japan for helpful advice.
References Bajaj YPS (1989a) Plant protoplasts (Biotechnology in agriculture and Berlin Bajaj YPS (1989b) Plant protoplasts (Biotechnology in agriculture and
and genetic engineering I. forestry 8.) Springer-Verlag, and genetic engineering II. forestry 9.) Springer-Verlag,
B erlin Cocking EC (1960) A method for the isolation of plant protoplasm and vacuoles. Nature 187:962-963 Cocking EC (1972) Plant cell protoplasts-isolation and development. Ann Rev Plant Physiol 23:29-50 Cheney DP (1984) Genetic modification in seaweeds: applications to commercial utilization and cultivation. In: ColweU RR, Pariser ER, Sinskey AJ (eds) Bioteclmology in the marine sciences. John Wiley & Sons, New York, pp 161-175 Cheney DP (1986) Genetic engineering in seaweeds: applications and current status. Beih Nova Hedwigia 83:22-29 Evans DA, Bravo JE (1983) Protoplast isolation and culture. In: Evans DA, Sharp W-R, Arnmlrato PV, Yamada Y (eds) Handbook of plant cell culture. Vol. 1 Techniques for propagation and breeding. Macmillan Publishing Co., New York, pp 124-176 Fitzsimons PJ~ Weyers JDB (1985) Properties of some enzymes used for protoplast isolation. In: Pilet P - E (ed) The physiological properties of plant protoplasts. (Proceedings in, life sciences.) Springer-Verlag, Berlin, pp 12-23 Fowke LC, Constabel F (1985) Plant protoplasts. CRC Press, Inc., Boca Raton, Florida Franssen J, Jetmiaux Ch (1965) Digestion de l'acide alginique ehez les inverttbrts. Cab Biol Mar 6:1-21 (In French with English and German summary) Fujimura T, Kawal T, Shiga M, Kajiwara T, Hatanaka A (!989) Regeneration of protoplasm into complete thalli in the marine green alga D T r - a ] ~ s a . Nippon Snisan Gakkaishi (Formerly Bull Japan Soc Sci Fish) 55:1353--1359 Kajiwara T, Hatanaka A, Fujimura T, Kawai T, hie M (1988) Isolation of protoplasts from marine brown algae Dictyotaceae plants. Nippon Suisan Gzkkalshi (Formerly Bull Japan Soc SCI Fish) 54:1255 Kloareg B, Polne--Fniler M, Gibor A (1987) Preparation of protoplasm from brown macroalgae. In: Indergaard M, Ostgaard K, Guiry MD (eds) Seaweed protoplast and tissue culture. COST 48, Brussels, pp 44-45 Millner PA, Callow ME, Evans LV (1979) Preparation of protoplasts from the green alga Enteromoljoha ~esz~aIAr (L.) Link. Planta 147:174--177 Onishi T, Suzuki M, Kikuchi R (1985) The distribution of polysaceharide hydrolase activity in gastropods and bivalves. Bull Japan Soc Sci Fish 51:301-308 Percival E, Rzhman MA, Weigel H (1981) Chemistry of the polysaccharides of the brown seaweed D J c / y o p t e ~ plaff/o~a. Phytoehemistry 20:1579-1582 Pilet P - E (1985) The physiological properties of plant protoplasts. (Proceedings in life sciences.) Springer-Verlag, Berlin Poine-Fuller M, Gibor A (1987) Tissue culture of seaweeds. In: Bird KT, Benson PH (eds) Seaweed cultivation for renewable resottrces. Elsevier Science, Amsterdam, pp 219-239 Pohae-Fuller M (1988) The past, present, and future, of tissue culmre and bioteehnology of seaweeds. In: Stadler T, Mollion J, Verdus M - C , Karamanos Y, Morvan H, Christiaen D (eds) Algal biotechnology. Elsevier Applied Science, London, pp 1731 Power JB, Chapman JV (1985) Isolation, culture and genetic manipulation of plant protoplasts. In: Dixon R A (ed) Plant cell culture: a practical approach. IRL Press, Oxford, pp 37-66 Pulte KJ, Dons JJM, Hnizing I/J, Kool AJ, Koornneef M, Krens F A (1988) Progress in plant protoplast research. Kluwer Academic Publishers, Dordrecht Saga N, Sakal Y (1984) Isolation of protoplasts from Z-.~,w/n,arfa and _Porpbja'a. Bull Japan Soc Sci Fish 50:1085 Saga N, Polne-Fuller M, Gibor A (1986) Protoplasts from seaweeds: production and fusion. Beih Nova Hedwigia 83:3743 Wu S (1988) Isolation and culture of protoplasts from Una'atia pd~taaffffda (Harv.) Suringar. J Ocean Univ Qingdao 18(21): 57--65 (in Chinese with English abstract) Yamaguchi K, Araki T, Aoki T, Tseng C - H , Kitamikado M (1989) Algal ceIlwaU-degrading enzymes from viscera of marine animals. Nippon Snisan Gakkaishi (Formerly Bull Japan Soc Sci Fish) 55:105-110 (In Japanese with English snmmary)
Plant Cell Reports (1995) 14:571-574
9 Springer-Verlag1995
Protoplast isolation in the marine brown alga Dictyopteris prolifera (Dictyotales) T. Fujimura 1, T. Kawai ~, T. Kajiwara 2, and Y. Ishida 3 1 Laboratory of Marine Bioteehnology, Shiono Koryo Kaisha, Ltd., Niitaka, Yodogawa-ku, Osaka 532, Japan 2 Department of Biological Chemistry, Faculty of Agriculture, Yamaguchi University, Yoshida, Yamaguchi 753, Japan 3 Department of Fisheries, Faculty of Agriculture, Kyoto University, Oiwake-cho, Kitashirakawa, Sakyo-ku, Kyoto 606, Japan Received 7 May 1993/Revised version received 15 November 1994 Communicated by A. Komamine
Abstract Protoplasts
were isolated from thalli of prolifera using a mixture of crude enzymes from vicera of live oysters (Crassostrea gigas) and the following commercial enzymes: an abalone enzyme, cellulase, polygalacturonase and hemicellulase. The enzyme mixtures produced up to 3.3 x 107 cells per 1 g of tissue fresh weight. The conversion to protoplasts of the cells was about 100% using the oyster enzyme or the abalone enzyme alone. The optimum pH for protoplast isolation was 6.0 and 20 hours were required for conversion to protoplasts.
Dictyopteris
Key words:
C m s s o s ~ ' e a ffZ~as -
pro~ra
Marine brown alga -
-
D1VO~o/TtelTir Protoplast-
Single cell Introduction Ever since Cocking work in 1960, plant protoplasts were adapted in various fields of plant science as useful experimental material (Cocking 1972). Protoplasts have been a useful research tool in physiology, morphology, biochemistry, pathology, cell biology, cell fusion, somatic hybridization, and genetic manipulation (Pilet 1985; Fowke and Constabel 1985; Puite et al. 1988). Researchers have been continuously progressing to advance the development of more efficient techniques for plant protoplasts research (Bajaj 1989a,b). On the other hand, in seaweeds protoplast research lagges far behind that of higher plants. Recently, the t e c h niques for isolation of protoplasts from seaweeds have been developing and advancing as described in various reviews (Cheney 1984, 1986; P o l n e Correspondence to: T. Fujimura
FuUer and Gibor 1987; Polne-Fuller 1988). Millner et al. (1979) reported the first isolation of protop-lasts from the marine green alga .Enteromo_rp,ha ~tes/z~ab3- by commercial enzymatic degradation. Since then, protoplasts from many green algae have been isolated using commercial cell wall-degrading enzymes commonly employed for higher plants (Cheney 1986). Brown algal protoplasts were produced only when feeder enzymes from marine animal guts which were previously fed on these specific seaweed were used for isolation procedures. Such enzymes were obtained from sea urchin (Saga and Sakai 1984), abalone (Saga et al. 1986), sea hare (Kloareg et al. 1987), and sea snail (Wu 1988). The induction of digestive enzymes in grazers for the p~oduction of brown algal protoplasts was suggested by P o l n e Fuller and Gibor (1987), since marine invertebrates feeding on benthic algae must contain the necessary digestive enzymes to dissolve brown algal polysaccharides in their diet. Kajiwara et al. (1988) selected enzyme systems consisting of marine feeder enzymes mixed with commercial ones, and further demonstrated the isolation of protoplasts from the brown algae Dictyotaceae. However, their isolation produced highly variable yields of protoplasts, possibly due to the purification levels of these crude biological enzymes. This paper reports the formation of protoplasts from the marine brown alga Z). pl'o/zi~ra via through an intermediate single ceils stage. We also report an effective method to isolate a great number of the protoplasts by using an enzyme mixture of partially purified oyster enzyme with a variety commercial ones. We then discuss some features of cell wall composition as revealed by the enzymatic varied effects of different enzymes on the degradation of cell walls.
572 Materials and methods M~e matr D. flfofffcra was collected from the coast of Shimonoseki and from the southern Osaka Bay, Japan. Live oysters were obtained from the beach of Yamaguchi, Japan. E~es. Some enzymes normally used for digesting higher plant cell walls were used; an Abalone Acetone Powder (an abalone enzyme) was purchased from Sigma Chemical Co. (St. Louis, USA), and Cellulase Onozuka R - 1 0 and Macerozyme R 10 were from Yakult Honsha Co., Ltd. (Tokyo, Japan), Driselase was from Kyowa Hakko Kogyo Co., Ltd. (Tokyo, Japan), and Bigatase M and Sumizyme X were fTam Kyowa Chemical Products CO., Ltd. (Osaka, Japan).
P~para~'on o f an oyster crude e~zj,z~e. The crude enzyme solution extracted from oysters was prepared by the following method. 100 g of entrails were homogenized with 100 ml of 50 mM sodium phosphate buffer (pH 7.0) containing seawater instead of water and filtered through some layers of gauze, then centrifuged at 12,000 x g for 10 rain. The super'natant was precipitated with 90 % saturation of ammonium sulfate for 3 h with stirring and then centrifuged at 12,000 x g for 15 rain. The precipitate was dissolved in a minimal amount of the same phosphate buffer (30-35 mi) and dialyzed for 12 h against several changes of the same buffer. The dialyzed liquid was brought up to 50 mi with 1.0 M mannitol solution in the same buffer and adjusted to pH 6.0. One ml of the solution (straight oyster enzyme) was used as an oyster crude enzyme for one test to isolate single cells and further to determine the protoplasts. All procedures were carried out at 5 *C. Prepara~,;on o f co~u~erHM e ~ T m c solul~bos. The commercial abalone digestive enzyme was dissolved into 5 % (w/v) solution in the buffer. The commercial enzymes were dissolved at 2 % (w/v) concentrations and then mixed equally into a solution. Ten ml aliquots of mixed enzyme solutions were tested on chopped seaweed tissues. Isolation o f sfngle c~ffs. After being washed three times with sterilized seawater, the thalli were cut into small pieces about 5 mm square. The pieces (0.5 g fresh weight) were moderately shaken (30-35 strokes/rain) at 25 ~ for 10 rain in 0.5 % papain solution containing 100 mM Tris-HC1 buffer (pH 7.4) and 1.0 M mannitol. They were transferred to other 1.0 M mannitoI solution (.pH 6.0) and shaken for 10 rain, and then 10 ml enzyme solution was added onto them to digest the cell walls. Single cells were released from the tissue after incubation in the enzymes for 15-20 h at 25 *C with moderate shaking (2530 strokes/rain). The released cells were collected with two kinds of filters (stainless mesh, 88-; nylon mesh, 40 tun-opening). The filtrates were washed three times in Tris-HC1 buffer (pH 8.0) contzlning 1.0 M mannitol, and centrifuged at 500 x g for 5 rain. The isolated single cells were counted with a hemocytometer.
Isola~bn o[protoplasts. Protoplasts, whose cell walls were completely removed by the enzymes, remained mixed with the single cells. They were microscopically counted on the basis of their morphological difference. FinaUy, the cell wall digestion was confirmed by the absence of fluorescent brightener, Calcofluor White ST (Sigma Chemical Co.) staining as previously described (Fujimura et al. 1989).
Rcsul~
fsola[fon of Siafflc cells When applied to chopped D. prolAFcra tissues, the variou~ enzyme ~ e s resulted in varying yields of single cells, and/or protoplasts. Sphere like single cells which were released from tissues are known as spheroplasts. They are on the way to convert themselves to the protoplasm if additional and suitable enzymes digesting the cell wails should be present- Table 1 showed that the commercial enzymes had no effects on formation of single cells, although those enzymes were shown to digest cell walls of many higher plants (Power and Chapman 1985) and some marine green algae (Fujimura et al. 1989). However, both the oyster, and abalone enzymes, and these two in mixtures, with the commercial enzymes, produced great number of single ceils (nearly 10 7 cells per 1 g of fresh weight as shown in Table 1). It suggested that some enzymes derived from algal grazers are more specialized than the available commercial enzymes in digesting brown algal cell walls, indicating that the brown algal ceil wails are composed of polysaccharides which differed from ceil walls of green algae or higher plants.
Isolation of l~otoplasts All of the isolated cells which were produced by the combined enzyme mixtures, consisting of the oyster enzyme, the abalone enzyme, and the commercial enzymes, were walless protoplasts (Table 1). Oyster enzyme, or abalne enzyme alone,
Table 1. Effect of different enzyme mixtures on isolation of single cells and protoplasts from D. prolifera. Enzyme solution Commercial enzymes Oyster enzyme Abalone powder Oyster enzyme + Comm. enzymes Abalone powder + Comm. enzymes Oyster enzyme + Abalone powder + Comm. enzymes 'a'Cells/g fresh weight.
Total number of isolated cells a 9.4 1.7 4.9 2.2 3.3
0 X 106 X 107 X 107 x 107 X 107
Protoplasts (%)
Walled single cells (%)
0 0 0 3 91 100
0 100 100 97 9 0
573 produced not protoplasts but walled single cells. The combination of enzymes produced a large number of the protopast (3.3 x 10 7 cells per 1 g of fresh weight) where were almost 100 % of the formation. The isolated protoplasts were completely spherical and 9-20 pin in diameter (Fig. 1). The abalone enzyme in combination with the ordinary enzymes produced much higher protoplasts ( 9 1 % yield) than the oyster enzyme mixed with the commercial enzymes did (3 %). This indicates that the abalone enzymes really contributed to the degradation of the intercellular matrix. However, the final addition of the oyster enzymes to the above enzyme mixtures resulted in a reliable and effective method for isolation of protoplasts of this brown alga.
The OpSmum Incuba~'on Time for the Isolation o f Protoplasm Table 3 presents the optimum incubation time for this method of protoplast isolation. Although, reasonable numbers of protoplasts were released after 3 h of incubation in the enzyme mixture (1.1 x 10 6 ceils per 1 g of fresh weight) the. yield increased in time and peaked at 20 h in enzyme treatment. Table 3. Effect of incubation time in the enzyme mixture on isolation of protoplasts from fresh tissue of D. prolifera. Time (h)
Protoplast number a
0 3 6 10 15 20 25 a Cells/gfresh weight.
0 1.1 X 10 6 4.0• 10 6 7.3 X 10 6 1.5 X 10 7 3.0 X 10 7 1.1 X 10 7
Discussion
Fig.1. Newly isolated protoplasts from
19.
prolifera. Scale bar=20 pm. The O p ~ z o ~ Protoplasm
pH
for the Isola[zbn of
The effect of p H on the protopast isolation was tested using the most effective enzyme mixture presented in Table 1. This enzyme ~ e had the highest cumulative activity for protoplast isolation at p H between 4.0-8.0 with an optimum pH around 6.0, as shown in Table 2. Table 2. Effect ofpH in the enzyme mixture on isolation of protoplasts from fresh tissue of D. prolifera. pH 4.0 5.0 6.0 7.0 8.0 a Cells/gfresh weight.
Protoplast number a 4.9 9.8 2.8 6.5 1.3
• x x • •
10 6 10 6 10 7 l0 6 10 6
The isolation of protoplasts from D. pro/ifera was exnmined by various combinations of different enzyme solutions (Table 1). Crude oyster enzyme solution in combination with the commercial enzymes, Cellulase Onozuka R - 1 0 , Macerozyme R - 1 0 , Driselase, Bigalase M, Sumizyme X, and Abalone Acetone Powder, released pure protoplast suspensions in a high yields. The oyster enzymes and abalone powder were essential for protoplast release from fresh tissues of D. p:olKem. But, none of the individual enzymes tested alone produced protoplast in different concentrations to evaluate their efficiency. This means the single enzymes do not have enough for an ability completely to decompose the cell walls in /9. prolzYera. Therefore, this suggests that the cell walls contain the polysaccharides which are degraded by the various enzymes used to isolate protoplasts. While, these marine herbivore enzymes can specialize in the digestion of characteristic polysaccharides, such as alglnate and fucoidan, which compose the cell walls of marine brown algae (Onishi et al. 1985; Yamaguchi et al. 1989). Thus, they are expected to contain brown algal polysaccharide-degrading enzymes such as alginate lyase and fucoidanase. The abalone powder from crude abalone entrails seem to also contain B-glucuronidase and sulfatase. The numbers of the
574 protoplast yielded successfully in this study were greater than those harvested from the marine green alga Ulva pen~sa (Fujimura et al. 1989). While, the sizes of 19]ct_-fopteliS protoplasts were smaller than that of the green alga. The pH was also important in effecting the yields of protoplasts. Generally, the pH optimum of the commercial cell wall-degrading enzymes, namely cellulase, polygalacturonase and h e m i cellulase, is known to be 4.0-7.0, whereas that of alginate lyases produced from marine herbivores is 7.0-8.0 (Franssen and Jeuniaux 1965). Fitzsimons and Weyers (1985) described the effect of pH on protease activity present in Cellulase Onozuka R I0, through the range of 6.0-10.0 (pH optimum ca. 8.0). From these reports, it is suggested that the commercial enzymes rather than the alginate-lyase greatly contributed to the degradation of the brown algal cell wall at the pH optimum, and that the proteases in the enzymes did not function. Furthermore, time was an important factor in increasing the yields. However, the viability of the isolated protoplasts decreased with the longer incubation in the crude marine herbivore enzymes. This can be attributed to various toxic substances and contaminants which may be present in the enzyme mixtures such as proteases, lipases, and other enzymes, phenolics, and salts (Evans and Bravo 1983). Our results show that the cell wails of D. p:off:c:a are composed of complex different constituted system, cellulose and non-cellulosic polysaccharides. The cell wall composition o f / 9 . pla~iog~-amma has been reported to contain fucoidan and alginate (Percival et al. 1981). The non-cellulosic polysaccharides such as alghaate and fucoidan are characteristic to cell walls of brown algae. The detailed complexity of these wails is not known. It is thus not surprising that mixtures of a large number of enzymes were required to completely remove the cell wall. Therefore, the oyster and abalone digestive enzymes are most likely more complex than just alginases and fucanases. AcA~7owledgemenls. We wish to thank Prof. A. Cribor and Dr. M. Polne-Fuller, University of California at Santa Barbara, USA for valuable discussion and critical reading of the manuscript. We are also grateful to Prof. H. Fukui, Kagawa University, Japan and Prof. J. Sekiya, Kyoto University, Japan for helpful advice.
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