INTRODUCTION
Brown algae (Phaeophyceae) are often morphologically complex with considerable anatomical differentiation based on parenchymatous or syntagmatic development (Fritsch 1945; Christensen 1980; Bold and Wynne 1985).
The level of differentiation approaches that in vascular plants in that tissue level differentiation occurs in many groups, but especially in members of Fucales and Laminariales and their relatives. It has long been known that different morphological parts of the same thallus can have very different photosynthetic properties (King and Schramm 1976; Arnold and Manley 1985; Kilar et al.
1989). Thus even the same tissue in different part of the same thallus can be cytologically differentiated (e.g., Davies et al. 1973; Fagerberg et al. 1979; Clayton and Ashburner 1990), suggesting differentiation of function associated with morphology and development.
Specialized cells are morphologically differentiated, and numerous cytological and ultrastructural studies suggest functional differentiation of cells associated with
photosynthesis, storage of photosynthate, transport, reproduction etc. (e.g., Oates 1988; Lobban and Harrison 1994).
Virtually all cells within complex brown algae have plastids. The outer tissues (meristoderm and cortex) have cells that are typically heavily pigmented with well-developed chloroplasts and perform a primary role in photosynthesis. Deeper into the thallus there are fewer chloroplasts and these tend to have different arrangements and be morphologically differentiated (e.g., McCully 1966; Schmitz and Srivastava 1974, 1975, 1976; Rüffer et al. 1978; Katsaros and Galatis 1985;
Clayton and Ashburner 1990). The extent to which these modified chloroplasts have different photosynthetic properties remains to be determined. Thus, given the cytological differentiation, it is of interest to evaluate if the photosynthetic process in these interior tissues is qualitatively or quantitatively different from those on the outside.
Here we use Microscopy-PAM (pulse amplitude modulation) fluorometry of chlorophyll a fluorescence to evaluate photosynthetic physiology within different tissues of the same thallus part. The use of fluorescence analysis for this general application is well established Algae
Volume 20(3): 233-238, 2005
Anatomical Differentiation and Photosynthetic
(Schreiber et al. 1994); however, technology to allow for fluorescence on spatial scales of single cells or tissues is more recent and carried out using Microscopy-PAM (Walz GmbH) (Schreiber 1998). Here we examine photosynthetic processes in adjacent cell and tissue types in a variety of parenchymatous and syntagmatic brown algae.
MATERIALS AND METHODS
Specimen collection and processing
All brown algal species (Table 1) were collected at low tide from Tor Bay, Guysborough Co, Nova Scotia in August 2004. Thalli were removed from rocks or collected from the drift, placed in plastic bags and transported on ice. Upon return to the laboratory (2 h) thalli were placed in seawater (18-20°C) with circulation and aeration under fluorescent lights (10 µmol photons m–2· sec–1) until processing (within 48 h).
Portions of plants were removed with a razor blade and hand sections were cut (ca. 0.5 to 1 mm thick) and placed in a well slide in seawater. Sections were cut from the blades of kelp species (the midrib in Alaria esculenta), or from mature, undamaged axes of mature fronds. A transverse section from each three plants was used for each species.
Fluorescence Measurement
Fluorescence measurements were performed using a Microcopy-PAM facility (Microscopy-PAM, Walz GmbH, Effeltrich, Germany). This consists of a modified epifluorescence microscope, the PAM control unit and a notebook computer with dedicated Windows-software (WinControl) for system operation and fluorescence analysis. Using visual inspection and an iris diaphragm the active field is narrowed so that the fluorescence characteristics of a small area (i.e. 200 µm diameter) can be assessed. The apparatus is equipped with a pin-hole
micro quantum sensor (MC-MQS) to assess quantum flux density of the blue excitation light in the object plane.
First, minimum fluorescence, Fo, was induced by low irradiation pulses in dark-adapted samples. Following a saturating flash, maximal fluorescence, Fm, was detected.
Variable fluorescence, Fv, was calculated as the difference between Foand Fm.
One-way analysis of variance (ANOVA) tests were used to determine if significant differences were present among the regions of transverse sections of each species for the fluorescence parameter. The non-parametric Kruskal-Wallis test was performed on data from each species, whereas a general linear model was used on pooled data. These analyses were performed using the software SPSS Release 11.0.1 (SPSS Inc., Chicago, IL, USA).
RESULTS
The photosynthetic parameters Fo (minimum fluorescence) and Fv/Fm (maximum quantum yield) were determined in the outer, middle and inner portions of transverse sections for eight species brown algae (Figs 1-8) representing four orders (Table 1). These corresponded to meristoderm-outer cortex, inner cortex and medullary portions of transverse sections, respectively. In all species Fowas always greatest in the outermost cell/tissue layers. For most species Fowas lowest in the innermost cells (medulla) and significantly lower than in the outermost cells (outer cortex or meristoderm) (ANOVA, p < 0.05). The exceptions were Desmarestia aculeata and Laminaria digitata where there was a rise or consistency in Fobetween inner cortical and medullary cells (ANOVA, p = 0.051). Within a given species, mean values for Fo from outer to inner layers ranged from 1:0.7 in D. aculeata to 1:0.2 in Laminaria saccharina. Across all eight species, values for Fofor Table 1. List of brown algal species, their taxonomic affiliation and plant portions examined
Species Order Portion of Plant Used
Alaria esculenta (L.) Grev. Laminariales Midrib of blade
Ascophyllum nodosum (L.) Le Jol. Fucales Main axis of frond
Chordaria flagelliformis (O.F. Müll.) C.Agardh Chordariales Main axis of frond
Desmarestia aculeata (L.) J.V. Lamour. Desmarestiales Main axis of frond
Fucus distichus ssp. distichus L. Fucales Main axis of frond
Fucus vesiculosus L. Fucales Main axis of frond
Laminaria digitata (Huds.) J.V. Lamour. Laminariales Mid portion of blade
Laminaria saccharina (L.) J.V. Lamour. Laminariales Mid portion of blade
medullary cells were 39 ± 16% of outer cells. These corresponded to the conspicuous change in pigmentation visible with the unaided eye across transverse sections of most species.
Levels of Fo showed positive correlations when the three areas of sections were compared across all species (r values from 0.317 to 0.463). Only the highest of these values (between outer and mid portions of sections) was significant (p = 0.023). When the two syntagmatic species were deleted from the analysis, the other two correlations (i.e., between inner and outer and between middle and inner parts of sections) became significant at p < 0.05 (r = 0.503 and 0.588, respectively).
There was less variation in maximum quantum yield between outer and inner cell types than for Fo. Values for medullary cells ranged from 42 to 72 % of outer cells (50
± 9%). When all species were considered together,
maximum quantum yield (Fv/Fm) varied significantly across transverse sections. Average values for the outer, densely pigmented cells fell within a narrow range from 0.513 (Alaria esculenta) to 0.589 (Fucus vesiculosus) (0.559 ± 0.026, X ± s.d.). This declined in inner cortical cells to 0.395 ± 0.063 and again in medullary cells to 0.296 ± 0.067. Values for the two syntagmatic species (C.
flagelliformis and D. aculeata) were not consistent with the overall pattern, and there was no difference between mid and inner portions of sections. When the parenchymatous taxa were considered alone (i.e., with the two syntagmatic species removed) all comparisons of cell-tissue types were significant at p < 0.001.
Correlations of values for maximum quantum yield between cell layers were not significant. Across all species and cell types Fo and Fv/Fm were highly correlated (r = 0.687, p < 0.001).
Garbary & Kim: Anatomy and Photosynthesis in Brown Algae 235
Figs 1-4. Photosynthetic parameters Foand Fv/Fmin meristoderm and outer cortex (outer), inner cortex (mid) and medulla (inner) portions of transverse sections from four species of Phaeophyceae. Values indicate mean ± s.d. (n = 3). Fig. 1. Alaria esculenta. Fig.
2.Ascophyllum nodosum. Fig. 3. Chordaria flagelliformis. Fig. 4. Desmarestia aculeata.
DISCUSSION
Our data from brown algae demonstrate the usefulness of Microscopy-PAM for showing changes in photosynthetic parameters on very small spatial scales.
Although minimum fluorescence and maximum quantum yield are quantitative parameters that are indirect measures of actual O2 production, they do provide a perspective on the physiological status of the different thallus tissues pertaining to carbon fixation. In this study we used only a representative portion of each species, and this work could be continued to evaluate developmental changes (e.g., young versus old branch segments in Ascophyllum) or tissue-organ differences (e.g., blade versus stipe in kelps).
The cutting of transverse sections might be considered
a major tissue trauma that would considerably increase stress and reduce quantum yields. This does not appear to be the case in the three fucoids examined here at least over the short-term duration of our measurements. Kim and Garbary (in preparation) used imaging PAM to examine age and organ related changes in photosynthetic parameters in the three fucoids studied here. Those data showed maximum quantum yields in the range of 0.6 to 0.7 that are only slightly higher than the values reported here for meristoderm-outer cortex tissues of the same fucoids.
Of the eight species examined, two stand out as different in terms of photosynthetic differentiation, i.e., Chordaria flagelliformis and Desmarestia aculeata. These species are entirely syntagmatic whereas the remaining fucoid and kelp species, have a parenchymatous organization (Christensen 1980). Chordaria flagelliformis Figs 5-8. Photosynthetic parameters Foand Fv/Fmin meristoderm and outer cortex (outer), inner cortex (mid) and medulla (inner) portions of transverse sections from four species of Phaeophyceae. Values indicate mean ± s.d. (n = 3). Fig. 5. Fucus distichus. Fig.
6.Fucus vesiculosus. Fig. 7. Laminaria digitata. Fig. 8. Laminaria saccharina.
was the only species in which Foand Fv/Fmwere not well correlated, with the former increasing and the latter declining between inner cortex and medulla. Desmarestia aculeata was also anomalous in that maximum quantum yield was the lowest of any species and it showed the smallest changes in both Foand Fv/Fm. This may reflect cytological deterioration as a consequence of damage incurred during its time in the drift and short exposures to air. Additional studies of syntagmatic brown algae, in general, and Desmarestia species, in particular, would be useful to extend our observations.
Despite the apparent lack of pigmentation in the inner cortex and medulla of most brown algae, considerable photosynthetic capacity is retained. Thus the apparently colourless medulla of all these brown algae retain substantial chlorophyll with Fo in medullary cells 20 to 74% of maximum levels (mean of 39%) in meristoderm or outer cortex. This suggests at least the possibility of significant photosynthetic capacity in medullary tissue.
Our data for maximum quantum yield supports this contention. Indeed, for six of the eight species, the ratio of quantum yield in medullary cells to epidermal or outer cortical cells was 42 to 72% (mean of 50%). This suggests that despite clearly lower pigment levels, that photosynthetic capacity may be compensated by higher photosynthetic potential in these cells. Thus medullary cells in brown algae may be physiologically differentiated from meristoderm/outer cortical cells with respect to photosynthetic function. This could be resolved with Imaging-PAM in which greater magnification was possible than with current instrumentation. In Laminaria saccharina Grevby et al.
(1988) concluded that chlorophyll amount and chloroplast area were not necessarily proportional to photosynthetic capacity. This is similar to our conclusion that maximum quantum yield in inner cortical and medullary cells may be elevated relative to chlorophyll amounts in different tissues.
Elsewhere we show organ level differences in photosynthetic fluorescence parameters in fucoid algae (Kim and Garbary, in preparation). This is analogous to previous work on kelp using more traditional O2 evolution and CO2 fixation approaches that demonstrated organ level variation (e.g., Arnold and Manley 1985). Those studies were primarily focused on developmental changes during growth and senescence.
Microscope-PAM has allowed us to examine physiological performance at the cell and tissue level, and to relate anatomy and cell differentiation to
photosynthetic capacity.
ACKNOWLEDGEMENTS
We thank John Garbary for assistance in the field. This work was supported by grants from the Natural Sciences and Engineering Research Council of Canada to DJG and MarineBio21, Ministry of Maritime Affairs and Fisheries, Korea to KYK.
REFERENCES
Arnold K.E. and Manley S.L. 1985. Carbon allocation in Macrocystis pyrifera (Phaeophyta): intrinsic variability in photosynthesis and respiration. J. Phycol. 21: 154-167.
Bold H.C. and Wynne M J. 1985. Introduction to the algae, 2nded.
Prentice-Hall, Englewood Cliffs, New Jersey, USA.
Christensen T. 1980. Algae: a taxonomic survey, Fasc. 1. Aio Tryk as, Odense, Denmark.
Clayton M.N. and Ashburner C.M. 1990. The anatomy and ultrastructure of ‘conducting channels’ in Ascoseira mirabilis (Ascoseirales, Phaeophyceae). Bot. Mar. 33: 63-70.
Davies J.M., Ferrier N.C. and Johnston C.S. 1973. The ultrastructure of the meristoderm of the hapteron of Laminaria. J. Mar. Biol. Ass. U.K. 53: 237-246.
Fagerberg W.R., Moon R. and Truby E. 1979. Studies on Sargassum III. A quantitative ultrastructural and correlated physiological study of the blade and stipe organs of S.
filipendula. Protoplasma 99: 247-261.
Fritsch F.E. 1945. The structure and reproduction of the algae.
Volume II. Cambridge University Press, Cambridge, UK.
Grevby C., Axelsson L. and Sundquist C. 1989. Light-independent plastid differentiation in the brown alga Laminaria saccharina (Phaeophyceae). Phycologia 28: 375-384.
Katsaros C. and Galatis B. 1985. Ultrastructural studies on thallus development in Dictyota dichotoma (Phaeophyta, Dictyotales). Br. Phycol. J. 20: 263-276.
Kilar J.A., Littler M.M. and Littler D.S. 1989. Functional-morphological relationships in Sargassum polyceratium (Phaeophyta): phenotypic and ontogenetic variability in apparent photosynthesis and dark respiration. J. Phycol. 25:
713-720.
King R.J. and Schramm W. 1976. Determination of photosynthetic rates for the marine algae Fucus vesiculosus and Laminaria digitata. Mar. Biol. 37: 209-213.
Lobban C.S. and Harrison P. J. 1994. Seaweed ecology and physiology. Cambridge University Press, Cambridge, UK.
McCully M.E. 1966. Histological studies on the genus Fucus. I.
Light microscopy of the mature vegetative plant.
Protoplasma 62: 287-305.
Oates B.R. 1988. Water relations of the intertidal saccate alga Colpomenia peregrina (Phaeophyta, Scytosiphonales). Bot.
Mar. 31: 57-63.
Rüffer U., Nultsch W. and Pfau J. 1978. Untersuchungen zur lichtinduzierten Chromatophorenverlagerung bei Fucus Garbary & Kim: Anatomy and Photosynthesis in Brown Algae 237
vesiculosus. Helgol. Wissens. Meeresunters. 31: 333-346.
Schreiber U. 1998. Chlorophyll fluorescence: new instruments for special applications. In: Garab G. (ed.), Photosynthesis:
mechanisms and effects. Vol. V. Kluwer, Dortrecht, The Netherlands. pp. 4253-4258.
Schreiber U., Bilger W. and Neubauer C. 1994. Chlorophyll fluorescence as a non-intrusive indicator for rapid assessment of in vivo photosynthesis. In: Schulze H.D. and Caldwell M.M. (eds.), Ecophysiology of photosynthesis.
Ecological studies, Vol. 100. Springer-Verlag, Berlin, Germany. pp. 49-70.
Schmitz K. and Srivastava L.M. 1974. Fine structure and development of sieve tubes in Laminaria groenlandica Rosenv. Cytobiologie 10: 66-87.
Schmitz K. and Srivastava L.M. 1975. On the fine structure of sieve tubes and the physiology of assimilate transport in Alaria marginata. Can. J. Bot. 53: 861-876
Schmitz K. and Srivastava L.M. 1976. The fine structure of sieve elements of Nereocystis lütkeana. Am. J. Bot. 63: 679-693.
Received 23 June 2005 Accepted 16 August 2005
INTRODUCTION
A unique ability to form protoplasts in vitro from the extruded protoplasm has been found in a number of the marine coenocytic green algae (Tatewaki and Nagata 1970; Rietema 1973; Ishizawa and Wada 1979; Kim and Klotchkova 2004). After cell injury, extruded cell organelles in seawater aggregate to form compact protoplasmic masses; a process seems to be mediated by a lectin-carbohydrate complementary system (Kim et al.
2002; Klotchkova et al. 2003).
The agglutinated protoplasmic masses soon change into spheres, i.e., protoplasts, surrounded with an envelope (Kobayashi and Kanaizuka 1985; Pak et al.
1991). Cytochemical and ultrastructural studies revealed that this envelope is not a lipid-based plasma membrane but is a membrane composed of some polysaccharides (Kim et al. 2001; Klotchkova et al. 2003). The envelope
transforms into a plasma membrane within several hours, presumably by incorporation of disintegrated plasma membrane of the precursor cell from the interior of protoplast onto its surface (Kim et al. 2001; Klotchkova et al. 2003). The protoplasts develop a new cell wall in a day or two.
In some species these cells grow into plants, whereas in other species they lose the ability to grow but remain as an enlarged single cell, producing numerous aplanospores or swarmers inside. It was therefore suggested that wound-induced protoplast formation in vitro may be a natural dispersal mechanism (Kim and Klotchkova 2004). Because coenocytic greens often grow in close proximity and their protoplasms may come into physical contact in nature, it is important to investigate if they can intermix while assembling the protoplast.
The dissociated cell components of the coenocytic protoplasm, separated by forcing it through sucrose gradient solution by ultracentrifugation, could be reunited into live cells and thus the formation of new species by mixing cell components from different Algae
Volume 20(3): 239-249, 2005
Experimental Hybridization between Some Marine Coenocytic Green Algae Using Protoplasms Extruded in vitro
Tatyana A. Klochkova, Kang-Sup Yoon
1, John A. West
2and Gwang Hoon Kim*
Department of Biology, Kongju National University, Kongju 314-701, Korea
1Department of Chemistry, Kongju National University, Kongju 314-701, Korea
2School of Botany, University of Melbourne, Parkville VIC 3010, Australia
Some marine coenocytic green algae could form protoplasts from the extruded protoplasm in seawater. The dissociated cell components of the coenocytic protoplasm could be reunited into live cells and, hence, the formation of new species by mixing protoplasms from different coenocytic cells has been predicted. Our results showed that an incompatibility barrier was present during protoplast formation in coenocytic algae to exclude foreign inorganic particles or alien cell components. No inorganic particles or alien cell components were incorporated into protoplast formed spontaneously in seawater. Even when the inorganic particles or alien cell and/or cell component were incorporated into protoplast in some experimental condition, they were expelled from the protoplast or degenerated within several days. A species-specific cytotoxicity was observed during protoplast hybridization between the protoplasms of Bryopsis spp. and Microdictyon umbilicatum. The cell sap of M. umbilicatum could destroy the cell components of Bryopsis spp., but had no effect on Chaetomorpha moniligera. Species C. moniligera and Bryopsis did not affect protoplast generation of either species. The wound-induced protoplast formation in vitro might have evolved in some coenocytic algae as a dispersal method, and the incompatibility barrier to alien particles or cell and/or cell component could serve as a protective mechanism for successful propagation.
Key Words:coenocytic green algae, cytotoxicity, hybridization, protoplasm, protoplast formation
*Corresponding author (ghkim@kongju.ac.kr)
coenocytic cells was predicted by Kobayashi and Kanaizuka (1985). The production of hybrid organisms by means of the induced fusion of somatic animal cells, or plant protoplasts prepared by enzyme digestion of the cell wall has been widely exploited in connection with cellular and genetic manipulation techniques (e.g.
Ephrussy 1972; Ringerts and Savage 1976; Markert and Petters 1978; Gleba and Sytnik 1984; Kalthoff 2001). It is a completely artificial method and bypasses the natural restrictions inherent in sexual crosses. The fusion techniques are known to be quite complex, requiring specific conditions and stabilizing factors in some cases (e.g. Gleba and Sytnik 1984; Kalthoff 2001).
In this paper, we describe the results of protoplasm hybridization between five different coenocytic species to determine whether or not intergeneric and interspecific protoplasm crosses can occur, as well as data on fusion of the protoplasts with unicellular algae or fluorescent beads. This investigation may lead to a better understanding of dispersal by the wound-induced protoplast formation and isolating mechanisms in this group of algae.
MATERIALS AND METHODS
Plant materials
The algae used in this research, their source and conditions for laboratory culture are given in Table 1. All isolates were maintained in IMR medium (Klochkova et
al. in press). The isolate of Chlorococcum sp. used in protoplast hybridization experiment was grown in a liquid ATCC Medium 625 (www-cyanosite.bio.purdue.
edu).
Protoplast preparation
For preparation of protoplasts of the coenocytes, the algal thalli were cut with a razor blade, squeezed out and the protoplasmic clumps were transferred to 90 x 15-mm plastic Petri dishes containing IMR medium. The protoplasts that formed were checked under an inverted microscope and all remnants of the disrupted cell wall were removed from the dish. The newly generated protoplasts were moved to the culture chamber and kept at the same conditions as specified in Table 1 for the original plants. Thereafter, the cells were checked daily.
All treatments described were repeated a minimum of 10 times. Numerical data present the means of multiple comparisons (n ≥10).
Protoplast hybridization experiment
Species combinations used in the hybridization experiment are given in Fig. 1.
Ability of protoplasm to take up inorganic particles To determine the ability of the protoplasm to take up inert particles spontaneously during formation, the algal thalli were cut in IMR medium containing 1-2 µL of FluoSpheres of 0.02-2 µm in diameter (yellow-green Table 1. List of algae used in this study, their collection site or source and conditions for laboratory culture
Conditions for laboratory culture
Species Temperature, Irradiance, µmol Light: Dark Collection site or source 0°C photons m–2· s–1 photoperiod, h
Bryopsis hypnoides (Lamouroux) 23, 25 30 12 : 12 Peter the Great Bay, East Sea, Russia, September 2003, col. T.A. Klochkova Bryopsis plumosa (Hudson) Agardh 23, 25 30 12 : 12 South Sea, Kachon, Korea, May 2000,
col. T.A. Klochkova
Chaetomorpha moniligera Kjellman 15, 20 < 15 12 : 12 East Sea, Kangneung, Korea, June 2002, col. T.A. Klochkova
Microdictyon boergesenii Setchell 23 30 12 : 12 Bahamas, June 1975, col. S. Brawley
(No. 1557)
Microdictyon umbilicatum (Velley) 23 30 12 : 12 Burill Lake, NSW, Australia, August
Zanardini 1996, col. J.A. West (No. 3674)
Chlamydomonas sp.1 12, 23 < 2-4, < 15-30 12 : 12 Daecheon (West coast), Korea, 2003
Porphyridium purpureum (Bory) 23 30 12 : 12 Daecheon (West coast), Korea, 1998
Drew et Ross1
Chlorococcum sp. 20, 25 15 12 : 12 Miruksazi stupa, Iksan, Korea,
April 2002, col. T.A. Klochkova
1Epiphytic microalgae found during culture of other isolates.
fluorescing carboxylate-modified microspheres, Molecular Probes). These FluoSpheres are inert particles used for observations of intracellular transportation (www.probes.com). Protoplasts that formed were checked under the blue filter (filter set BA510IF and BA510-540) to detect if the beads were taken up.
In another experimental set, protoplasm was extruded in artificial seawater of pH 6 to loosen the binding between cell organelles (Kim et al. 2002), and centrifuged at 2,000g for 2 min in 1-mL plastic tubes. The cell organelles in the pellet were re-suspended in new artificial seawater of pH 6, and 1-2 µL of ddH2O containing FluoSpheres was added. The tube was shaken vigorously for several minutes to incorporate the fluorescent beads into protoplasmic masses and then centrifuged at 2,000g for 2 min. The pellet containing protoplasmic masses and fluorescent beads was re-suspended in IMR medium. Then, the protoplasts that formed were checked under the blue filter to detect if the beads were taken up.
Hybridization of protoplasts from coenocytic algae with unicellular algae
Unicellular microalgae Chlamydomonas sp., Chlorococcum sp., and Porphyridium purpureum were used to determine the ability of the protoplasts to include other living organisms during formation. Chlorococcum sp. from soil was used in hybridization experiment because of its ability to tolerate salinity stress (Klochkova et al. in press).
As the cells of P. purpureum were surrounded with a mucilaginous sheath, they were first centrifuged at 1,500g for 1 min and washed for 3-5 min with 0.5%
Triton X-100 diluted in IMR medium. The procedures for protoplasm extrusion and fusion with the unicellular algae were performed as described above for FluoSpheres, using IMR medium and artificial seawater of pH 6 for protoplasm extrusion. Protoplasts formed were checked under the light microscope according to time.
Hybridization of protoplasts of different coenocytic algae
To hybridize protoplasm between different coenocytic green algae, the thalli of each species were placed together on a slide glass and cut with a razor blade in several drops of IMR medium. Protoplasm extruded from the thalli was transferred to 90 x 15-mm Petri dishes containing IMR medium and examined under the inverted microscope over time intervals (5, 10, 20, 30, 40, 50, and 60 min).
To observe the interactions between vacuolar-sap-free cell organelles of different algae, protoplasm of each species was extruded separately in artificial seawater of pH 6, centrifuged at 2,000g for 2 min and washed with the same solution twice. The cell organelles were aggregated very loosely or detached from each other after the treatment. Thereafter, artificial seawater was removed and the cell organelles of different species were re-suspended and mixed together in a small volume of IMR medium and observed under the light microscope.
Hybridization of protoplasm with foreign cell sap The effect of the cell sap of one coenocytic cell on the cell organelles of another species was studied as follows.
The cell sap of Microdictyon umbilicatum was obtained by centrifuging the protoplasm extruded in IMR medium at 2,000 g for 2 min and removing the pellet. The cell sap was mixed with the chloroplasts of Bryopsis spp. and Chaetomorpha moniligera. The mixture was directly transferred into 90 x 15-mm Petri dishes containing IMR medium and examined under the inverted microscope over time intervals (5, 10, 20, 30, 40, 50, and 60 min).
Scanning electron microscopic observations
Dense suspensions of protoplasts were placed on the cover slips coated with 3% 3-aminopropyltriethoxy-sylane (Sigma) diluted in acetone and were then fixed with 3.7% formaldehyde (Junsei Chemical Co., Japan) in Klochkova et al.: Hybridization between Marine Coenocytic Green Algae Using Protoplasms 241
Fig. 1. Species combinations used for the hybridization experiments and the results obtained. Symbols ‘-’ and ‘+’
mean negative and positive hybridization, respectively;
symbol ‘++’ implies that hybridization was very successful.
Abbreviations, B. hyp Bryopsis hypnoides; M. boer -Microdictyon boergesenii; C. mon - Chaetomorpha moniligera;
M. umb - Microdictyon umbilicatum; Chlamy - Chlamydomonas sp.; Chloroc - Chlorococcum sp.; Porphy - Porphyridium purpureum; Fluosph - FluoSpheres.