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1 Division of Natural Sciences, John Brown University, Siloam Springs, Arkansas 72761
2 Department of Biological Sciences, Auburn University, Alabama 36849-5414
* To whom correspondence should be addressed. E-mail: twakefie{at}jbu.edu.
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Introduction |
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TOPAbstractIntroductionMaterials and MethodsResultsDiscussionLiterature Cited |
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In an attempt to consolidate endosymbiotic research so that homologies in all the systems could be investigated, Roth et al. (1988) introduced the terms "symbiosome" and "symbiosome membrane" to describe the intracellular symbiotic compartment in which symbionts reside. In zooxanthellae–cnidarian symbioses, the symbiosome is commonly recognized to include a continuous outer, or vacuole, membrane underlain by multiple layers of additional membrane that surround the "pellicle," or cell wall, of the internal symbiont (Battey, 1992). The origin and nature of these membranes have been debated since they were first described. Taylor (1968) stated that some of these membranes were formed from host tissue, but others were contributed by the symbiont itself; however, he also stated that determining which membranes came from which source was difficult. Dodge and Crawford (1970), in their investigation of the "periplast" of zooxanthellae, agreed with Taylor, stating that "it is impossible to tell whether these [membranes] belong to the host or alga." Undaunted by this proclamation, many papers have been published suggesting possible origins for these membranes. These suggestions have naturally centered on two possible origins: (1) the host cell (Colley and Trench, 1983; Palincsar et al., 1988; Rands et al., 1993; Tripodi and Santisi, 1982), and (2) the algal cell (Kevin et al., 1969; Taylor, 1971; Trench and Winsor, 1987; Wakefield et al., 1998, 2000). Unfortunately, in all cases, suggested origins are presented without any supporting experimental evidence.
In this study, immunocytochemical methods were employed to obtain evidence needed to support or dispute the claims of symbiont versus host origins of the multiple layers of apparent membrane surrounding the endosymbiont. The marine anthozoan Aiptasia pallida and its endosymbiont Symbiodinium bermudense (Banaszak et al., 1993) were used in confocal and electron microscopic investigations that employed two monoclonal antibodies produced against antigens associated with the membranes surrounding the algal symbiont. The data obtained provide the experimental evidence needed to reject one of the proposed hypotheses for the origin of the multiple layers of membrane. The significance of these findings is discussed.
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Monoclonal antibody production
Antigen preparation. Eight specimens of Aiptasia pallida, each about 5.0–8.0 cm long, were removed from the aquaria. Two specimens were placed in 6.0 ml of a homogenization solution consisting of 1.0 mM phenylmethylsulfonyl fluoride (PMSF) in 0.45-µm Millipore-filtered seasoned aquarium water (MFSAW) pre-cooled to 4 °C. The tissues of the specimens were fully dispersed using a Handishear (Virtis) tissue homogenizer; an ice bath was used to maintain the pre-cooled temperature. This process was repeated for all eight specimens, which were then combined and split into four samples. Each sample was poured into 15-ml glass centrifuge tubes and centrifuged at about 250 x g for 3 min to pellet the extracted symbiosomes. The supernatant was discarded, and the pelleted symbiosomes were resuspended in homogenization solution. This process was repeated three times until a firm pellet of extracted symbiosomes was obtained in each tube. Each symbiosome pellet was resuspended in 1.0 ml of nanopure filtered water (np-H2O) and either remained untreated or was processed in one of the following ways: (1) fixed in 4% paraformaldehyde containing 0.2 M Millonigs phosphate buffer (MPB) for 10 min and then rinsed three times in 0.2 M MPB and finally resuspended in np-H2O, (2) boiled for 1 min, or (3) repeatedly frozen and thawed by placing the sample in an Eppendorf tube, dropping it into liquid nitrogen, removing and warming until it thawed, and then repeating. All four types of samples were then combined to produce about 4.0 ml of "symbiosome slurry." This slurry was used as a source of antigen for the production of antibodies.
Antibody production. The production of monoclonal antibodies followed standard procedures (Kohler and Milstein, 1975). Briefly, BALB\/C mice were injected subcutaneously with 0.1–0.2 ml of the symbiosome slurry in Freund’s complete adjuvent. At 3 weeks and 6 weeks, symbiosome slurry in Freund’s incomplete adjuvent was injected into the peritoneal cavity. Seven days after the last injection, blood was taken from the tail vein of the inoculated mice. Blood sera were titered 1:50 through 1:10,000 and placed in a 96-well plate coated with fixed symbiosomes to undergo an enzyme-linked immunosorbant assay (ELISA) for the presence of symbiosome antibodies.
Once levels of serum antibodies were sufficient, a mouse was selected and sacrificed. Lymphocytes were removed from the spleen, washed to remove red blood cells, and then suspended in RPMI 1640 tissue culture medium (RPMI). Throughout the fusion procedure, this suspension was maintained at 37 °C by using a dry heating block. A 4:1 mixture of lymphocytes and AG-8.653 myeloma cells was added to a sterile centrifuge tube and pelleted. The supernatant was removed, and a solution of 50% polyethylene glycol in 20 mM phosphate buffered saline (PEGM) was slowly added while the cell suspension was constantly stirred. Additional RPMI was then added while stirring continued. After several minutes the suspension was again centrifuged to pellet the cells, and the PEGM + RPMI medium was removed and replaced with RPMI supplemented with BSA (15% bovine serum albumin), OPI (0.15% oxaloacetate, 0.05% pyruvate, 0.0082% bovine insulin), and HAT (5 x 10-5 M hypoxanthine, 2 x 10-7 M aminopterin, 8 x 10-6 M thymidine). The resulting fusion mixture of cells was aliquoted in 100-µl amounts into the wells of 96-well plates that were then incubated at 37 °C.
Developing cell cultures were periodically fed and checked as they increased in size. Once the 96-well plate cultures had reached one-third confluency they were screened by an immunofluorescent dot-spot test to identify culture wells that were producing antibodies against symbiosome antigens (see below).
Screening. Symbiosomes that had been extracted using the above-outlined procedure were fixed in 4% paraformaldehyde and 0.2 M MPB for 10 min. Fixed symbiosomes were rinsed three times in 0.2 M MPB and then resuspended and diluted in np-H2O. Spots of fixed symbiosomes were dried onto spot slides. Each spot was incubated first in 30 µl of blocking media (BM) consisting of 5% heat-inactivated goat serum in 20 mM phosphate buffered saline solution containing 0.1% Triton X-100 (PBS+). Subsequently, spots were incubated in 30 µl each of RPMI supernatant from cell-culture wells for about 1 h. Spots were then washed three times in PBS+ and incubated in goat anti-mouse fluorescence isothiocyanate (FITC)-conjugated secondary antibody (1:100 in BM) for 1 h, followed by three more rinses in PBS+. Spots were then examined with a Zeiss epifluorescence microscope. Cells in culture wells producing symbiosome antibodies were suspended in RPMI and cloned by limiting dilution. Upon reaching one-third confluency, clone wells were screened and positive wells cloned again. This process of recloning was repeated until all of the clones in a freshly inoculated 96-well plate showed the same degree of antigen recognition. The final outcome was the isolation of two hybridoma cell lines (G12 and PC3) producing antibodies that recognized antigens associated with the symbiosome membranes. The two cell lines were then expanded, allowing collection of antibody containing spent culture medium for immunohistochemistry and providing cells for frozen storage in liquid nitrogen.
The two cell lines were isotyped using an ISO-1 isotyping kit purchased from SIGMA chemical company. The G12 antibody was found to be an IgG, and the PC3 antibody was an IgM.
Immunocytochemistry and confocal microscopy
Cultured symbionts. Axenic cultures of Symbiodinium microadriaticum (Freudenthal, 1962) extracted from Cassiopea xamachana, S. pulchrorum (Banaszak et al., 1993) extracted from Aiptasia pulchella, and S. bermudense extracted from A. tagetes (= A. pallida Muller-Parker et al., 1996) have been continuously maintained in incubation chambers within our laboratory for about 3 years (cultures were initially provided by Dr. William Fitt, University of Georgia). Incubation temperatures were maintained at 28.5 °C and illumination of the cultures was on a 12-h light/12-h dark cycle. Cultures were grown in 10 ml of ASP-8A culture medium (Blank, 1987) in sterile 15-ml culture tubes. New cultures were started from existing cultures roughly every 2 months, using sterile techniques.
Samples from each culture were removed and fixed in 4% paraformaldehyde in 0.2 M MPB. Fixed algal cell samples were rinsed three times in 0.2 M MPB and then incubated in the following solutions: BM for 1 h, PC3 primary antibody (1:100 in BM) or G12 primary antibody (1:100 in BM) for 3 h, rinsed 3 times in PBS+ for 10 min each, FITC-conjugated–IgM-specific secondary antibody (1:100 in BM) for PC3 antibody or FITC-conjugated–IgG-specific secondary antibody (1:1000 in BM) for G12 antibody for 1 h, followed by three rinses in PBS+ for 10 min each. Controls were also run using either BM only or a nonspecific antibody-producing hybridoma cell supernatant in place of the primary antibody solution.
Extracted symbiosomes. Symbiotic Aiptasia pallida specimens were homogenized, and the symbiosomes were extracted and fixed using the previously described procedures. Labeling of the fixed symbiosomes with the same controls was done using the procedures described above for cultured symbionts.
Cryostat samples. Small (0.5–1.0 cm) symbiotic and aposymbiotic specimens of Aiptasia pallida were fixed and rinsed following the same procedure as used on the extracted symbiosomes. After rinsing, these specimens were placed in a 1:1 mixture of 0.2 M MPB and Fisherbrand tissue freezing medium (TFM) and left overnight. Specimens were then removed from the 1:1 mixture and placed in 100% TFM. Specimens were frozen, and thin sections were cut using a Reichert-Jung 2800 Frigocut N cryostat. The sections were collected and placed on warm, 4% gelatin-coated slides. The tissues were labeled using the same procedures as those used to label extracted symbiosomes. Sections of symbiotic tissue and aposymbiotic tissue were tested for positive labeling using both PC3 and G12 primary antibodies. The same controls as those used on the extracted symbiosomes were used on the cryosectioned tissue.
Whole-tissue samples. Tentacles (2.0–3.0 cm long) from A. pallida were removed from symbiotic specimens. Small (0.5–1.0 cm) whole, aposymbiotic A. pallida specimens were also collected. Both types of tissue were fixed and rinsed following the same procedure as that used for extracted symbiosomes and sectioned tissue. After the final rinse, the aposymbiotic specimens were cut into small pieces (2.0–3.0 mm). Symbiotic tentacles were also cut into small pieces (2.0–3.0 mm), and additional cuts was made longitudinally through the tentacle so that the symbiotic endodermal layer was fully exposed to the incubating solutions. Single- and double-labeling experiments using G12 and PC3 primary antibody were performed using the same procedures and controls as described above for extracted symbiosomes and sections.
Imaging. Labeled symbiosomes, sections, and whole-tissue samples were placed on glass slides and infiltrated with either glycerol mounting medium containing 4% n-propyl gallate (pH 9.5, Giloh and Sadat, 1982) or Permafluor aqueous mounting medium (Beckman Coulter). Fluorescent images were captured using a BioRad MRC-1000 laser scanning confocal microscope equipped with a krypton-argon laser. For those tissues that were labeled only with FITC-conjugated antibodies, the FITC-labeled tissue was visualized first with 488-nm laser excitation and a 500-nm fluorescent emission filter. A second image was then acquired of the autofluorescence of the algal chloroplast by using 514-nm laser excitation and a 660-nm fluorescent emission filter. When the filter sets provided with the confocal microscope are used, autofluorescence due to chlorophyll is very dim; thus it was necessary to maximize the gain to capture images of chloroplast fluorescence. Each image was saved, pseudocolored (green for FITC; red for autofluorescence), and merged using BioRad’s confocal assistant. For double-labeled tissues the same procedures were employed as for single-labeled tissues, except that maximizing the gain for the RITC-labeled tissue was unnecessary because its fluorescence was much brighter than the apparent autofluorescence of the chloroplast.
To ensure that the labeling could be compared accurately, the positively labeled tissue was imaged first. The laser intensity, iris diameter, gain, black level, and enhancement were all optimized for the FITC label. After switching the filter set, each parameter (i.e., laser intensity, iris diameter, etc.) was again optimized to capture either the autofluorescent algal chloroplast or the RITC label. Once both sets of parameters were established (one for FITC and the other for either autofluorescence or RITC label), they were not changed during the image acquisition of other experimental tissues.
Electron microscopy
Standard fixation and embedding. Small pieces (2–4 mm) of symbiotic tentacle were removed from a live anemone and fixed in 0.5% glutaraldehyde + 4% paraformaldehyde + 0.2 M sucrose in 20 mM phosphate buffered saline (PBS) solution at 37 °C for 4 h. Tissues were rinsed in 20 mM PBS + 0.2 M sucrose three times for 15 min each time and dehydrated in an ethanol series (30%, 50%, 70%, 85%, 90%, 95%, 100%) for 30 min in each, at room temperature, with constant rotation. Tissues were then incubated in a 1:1 mixture of 100% ethanol and LR White and left overnight with constant rotation. The next day, tissues were placed in 100% LR White, incubated for 6 h with constant rotation, and then placed in fresh LR White in an aluminum pan; the pan was placed inside an acrylic plastic chamber and sealed. Intake and outflow valves were used to flood the chamber with N2 and exclude oxygen. Finally, the sealed chamber was incubated at 70 °C overnight to polymerize the plastic.
Cryofixation and embedding. As above, small pieces (2–4 mm) of symbiotic tentacle were removed from a live anemone. Two or three pieces were placed in a brass planchet, inserted into a Balzers 010 HPM high pressure freezing machine, and quick-frozen. Frozen tissues were transferred to liquid N2 and held there briefly before being transferred into a Leica EM AFS automatic freeze-substitution system. The tissues were placed in a freeze-substitution medium of methanol (MeOH) + 0.5% uranyl acetate. The freeze-substitution medium and tissues were maintained at -90 °C for 2 days before the temperature of the system was slowly ramped up 5 °C/h until it was stabilized at -45 °C. The freeze-substitution medium was removed and the tissues were rinsed twice, for 30 min each time, in pure MeOH. The tissues were then incubated at -45 °C in the following: a 1:1 mixture of MeOH:Lowicryl HM20 resin for 2 h, a 1:2 mixture of MeOH:HM20 for 2 h, pure HM20 for 2 h, and finally pure HM20 overnight. Tissues were placed into molds filled with fresh HM20 and polymerized at -45 °C by ultraviolet light over a 3-day period.
Sectioning, labeling, and staining. Blocks of the standard-fixed and cryofixed tissues were trimmed, and ultrathin sections were cut using a Reichert-Jung Ultracut E ultramicrotome. Sections were collected on carbon-coated, Formvar-covered slot and 100-mesh grids. Grid-mounted sections were incubated on drops of the following solutions: 0.05 M glycine in incubation buffer (10 mM phosphate buffer + 150 mM NaCl + 0.1–0.2% Aurion BSA-c [acetylated bovine serum albumin] + 20 mM NaN3) for 15 min, blocking medium (incubation buffer + 5% BSA + 0.1% cold water fish skin gelatin + 5% heat inactivated goat serum) for 30 min, incubation buffer 3 x 5 min, diluted primary antibody (see below) in incubation buffer overnight, incubation buffer 6 x 5 min, immunogold-conjugated secondary antibodies (see below) diluted in incubation buffer for 2 h, incubation buffer 6 x 5 min, 10 mM PBS 3 x 5 min, postfixed in 2.5% glutaraldehyde in 10 mM PBS for 5 min and then rinsed in 10 mM PBS for 5 min, nanopure H2O 5 x 2 min, Aurion silver enhancement for 20 min, nanopure H2O (np-H2O) 3 x 5 min, saturated uranyl acetate in 50% MeOH for 20 min, and nanopure H2O 3 x 5 min. Grids were then carefully blotted by touching the edge of the grid to filter paper and allowing most of the water to be wicked away. Grids were then placed on filter paper and allowed to dry. Labeled tissues were viewed and photographed using a Zeiss EM-10 transmission electron microscope.
Primary antibody dilutions used in the procedure above were either PC3 1:100, G12 1:100, or both 1:100 in incubation buffer. Secondary antibodies used were goat anti-mouse (GAM) IgM-specific antibody conjugated to an ultrasmall (
1.0 nm) gold particle (Aurion), GAM IgG-specific antibody conjugated to an ultrasmall gold particle (Aurion), or a combination of the GAM IgM-specific antibody–ultrasmall gold with a GAM IgG-specific antibody conjugated to a 20-nm gold particle (ICN). Some sections were treated with protein A conjugated to a 15-nm gold particle (Aurion). Sections were incubated in both 1:10 or 1:100 dilutions of secondary antibody or protein A in incubation buffer.
Single-label controls included incubations in BM or nonspecific primary antibodies in place of PC3 or G12. Controls for nonspecific cross reaction of the secondary antibodies were also done. This included incubation in PC3 followed by the IgG-specific secondary antibody and incubation in G12 followed by the IgM-specific secondary antibody. Double-label controls included incubations in BM, in nonspecific primary antibodies, in PC3 only, or in G12 only; each was followed by incubations in both secondary antibodies.
Ultra-cryomicrotomy. Small pieces (2–4 mm) of symbiotic A. pallida tentacle were removed from a live specimen. Tentacle pieces were fixed using the same protocol as that used in the standard fixation and embedding procedure described above. However, after the tissue had been rinsed it was placed in an 80% sucrose solution at 4 °C and left overnight. The following day the tentacle pieces were removed and placed on small metal stubs. Care was taken to remove excess sucrose solution, and then the stub and tissue were plunged into liquid nitrogen (LN2) until completely frozen. Following this initial freezing the tissues remained frozen at all times, either in LN2 or in the cold chamber of the ultra-cryomicrotomy apparatus.
The frozen tissue was trimmed and sectioned on the Reichert-Jung Ultracut E ultramicrotome fitted with an FC4E ultra-cryomicrotomy unit. Trimming was done at -70 °C with glass knives; sections were cut at -100 °C with a Diatome 45° angle ultra-cryomicrotomy diamond knife. A Diatome Static Line I was also used to reduce the build-up of static charge on the knife surface, allowing a fine ribbon of 75- to 85-nm sections to be produced. Each ribbon was pulled from the knife’s edge with an eyelash brush and picked up by using a tiny drop of 80% sucrose suspended in a wire loop. (The sucrose drop begins to freeze as it is brought near the knife surface, and the tissue was picked up just before the drop was completely frozen. Thus the tissue was frozen onto the sucrose drop and removed from the knife.)
The drop was removed from the cryo-chamber and allowed to thaw. The drop was then applied, tissue side down, to a 100-mesh, Formvar + carbon-coated nickel grid. The grid adhered to the drop so that it could be flipped over and floated off the wire loop onto a plate of 4 °C solidified 2% gelatin in np-H2O. Grids could be stored on the gelatin for several days before being used in the immunogold labeling procedure.
In preparation for labeling the grids, a small amount of the solidified gelatin was removed from around and under each grid. The grids and gelatin were placed on a small piece of Parafilm attached to a glass petri dish and placed in a 37 °C oven for about 30 min. When the plate was removed, the grids could be easily picked out of the now-liquid gelatin with fine forceps. The grids were immediately placed into the 0.05% glycine solution and were incubated in all of the same solutions used on the plastic-embedded tissues. After the final rinse in np-H2O, the grids were floated onto a solution of 2% methyl cellulose in np-H2O + 2% uranyl acetate (MU). The grids were then picked up with a wire loop that was just slightly larger than the grid itself, and the excess MU was wicked away by touching the edge of the loop to a piece of filter paper. Care was taken to ensure that only a thin layer of MU was left over the tissue. The MU was then allowed to dry completely, and the grids were removed from the loops and stored in grid boxes until they were viewed and photographed using the Zeiss EM-10 transmission electron microscope.
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Controls for cryosectioned symbiotic and aposymbiotic tissues and whole pieces of tissue that were incubated in either BM or a nonspecific primary antibody showed either no fluorescence in aposymbiotic tissues (Fig. 2b) or only the autofluorescence of symbiont chloroplasts (Fig. 2e).
Immunofluorescence double label
Double-labeled tissues (Fig. 2f) revealed the PC3 antibody labeling (green) to be confined to a very narrow band immediately surrounding the symbionts. Alternatively, G12 labeling (red) was distributed evenly throughout the entire endodermal layer of the host and also appeared as a thin yellow line (the overlap color of red and green) that was immediately adjacent and external to the PC3 label (Fig. 2f). [Note that since the autofluorescence of the algal cell chloroplast is emitted at the same wavelength as the much brighter RITC label used for G12 identification, it was not possible to boost the gain and capture it. This resulted in the symbionts appearing as "black holes" (e.g., Fig. 2f), in contrast to what is seen in the single-labeling experiments (e.g., Fig. 2c).]
Immunogold single label
Table 2 summarizes the results of the immunogold labeling in these experiments. Standard fixed tissue and rapid-freeze–freeze-substituted–low-temperature-embedded tissue labeled with G12 and examined with the electron microscope showed positive labeling of the host cell plasmalemma and internal membranes. In particular, the outermost symbiosome membrane was clearly labeled (Fig. 3a). There was also a strong labeling within the mesoglea of the host tissue (Fig. 3b). However, there was no labeling of the symbiont. A close examination revealed that the multiple layers of membrane that are immediately adjacent to the symbiont were not labeled by the G12 antibody (Fig. 3c); the ectodermal layer of the host tissue also lacked label. In fact, long extensions of the mesoglea could be traced into the ectodermal layer due to the strong label found therein and the absence of label within the ectodermal cells (Fig. 3b).
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The tissue sectioned by ultra-cryomicrotomy confirmed our findings that the G12 antigen indicated specific binding in the host membranes and mesoglea (Fig. 4a, b). It also revealed the specificity of the PC3 antibody to the algal symbiont (Fig. 4c, d). The labeling with the PC3 antibody was significant because our early attempts to label standard-fixed, LR-White-embedded tissue with PC3 had totally failed. Thus, the ultra-cryomicrotomy technique provided our first glimpse of the specificity of the PC3 antibody to the algal symbiont. Unfortunately, the preservation of ultrastructure in the ultra-cryomicrotomy technique was less than favorable, necessitating the use of the freeze-substituted–Lowicryl-embedding technique.
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The finding that cultured S. microadriaticum is not labeled by the PC3 antibody but both S. pulchrorum and S. bermudense are labeled may reflect phylogenetic differences between the three tested algal species. In past studies, evidence based on karyotyping (Blank and Trench, 1985), morphology (Schoenberg and Trench, 1980b; Trench and Blank, 1987), isoenzyme patterns (Schoenberg and Trench, 1980a), physiology (Fitt and Trench, 1981; Chang et al., 1983; Fitt and Trench, 1983), and biochemistry (Chang and Trench, 1982; Govind et al., 1990) has indicated that the genus Symbiodinium consists of many different species. Also, an examination of similarities and differences in the genes that encode for the small ribosomal subunit of algal symbionts has revealed that S. bermudense and S. pulchrorum are closely related but different species, and both have been placed into clade B of the "zooxanthellae" phylogenetic tree (Rowan, 1998). S. microadriaticum, on the other hand, is more distantly related and has been identified as belonging to clade A (McNally et al., 1994; Rowan, 1991, 1998; Rowan and Powers, 1991). The results of our immunofluorescent studies support the concept of phylogenetic diversity among the symbionts and may be added to the list of biochemical differences that exist among some Symbiodinium species.
The double-labeled immunofluorescent and immunogold results clearly demonstrate that the algal symbionts are surrounded by layers of material that originate from the different symbiotic partners. The narrow region of PC3 labeling that is peripheral to the symbionts and is surrounded by a single, host-derived "symbiosome membrane" and also by a large ubiquitous area of G12 labeling illustrates that a limited layer of this symbiont-derived antigen is maintained within a vast amount of host tissue. The immunogold results indicate that the PC3 antigen is present in the algal cell wall, but only in areas directly adjacent to high cytoplasmic levels of this antigen. This suggests that these may be areas of secretion for a molecule or molecules containing the PC3 antigen. The fact that the PC3 antigen is found both within the algal cell cytoplasm and also closely associated with the multiple layers of membrane, beneath the G12-labeled host symbiosome membrane, supports the hypothesis that these membranous layers are of symbiont rather than host origin.
The results of this research should help clarify some confusing terminology currently used in the field of cnidarian–dinoflagellate endosymbiosis. The term "symbiosome," as introduced by Roth et al. (1988), was defined as "a membrane-bound compartment containing one or more symbionts and certain metabolic components located in the cytoplasm of eukaryotic cells." They further describe the "symbiosome membrane" as being a host-derived membrane that surrounds the microsymbiont, creating a "symbiosome space" (Roth et al., 1988). This terminology has been adopted by researchers in various fields of endosymbiotic research, including plant–bacteria interactions (Day and Udvardi, 1992; Udvardi and Day, 1997), protozoan–bacteria interactions (Choi and Jeon, 1992; Jeon, 1992), and invertebrate–algal interactions (Rands et al., 1993; Wakefield et al., 1998; Wakefield et al., 2000). For plant–bacterial and protozoan–bacterial interactions, these terms are adequate and clearly define the endosymbiotic situation of these partners. However, for invertebrate–algal symbioses, particularly cnidarian–dinoflagellate symbioses, use of the term "symbiosome membrane" has been problematic.
In situ dinoflagellate symbionts have been identified as existing predominantly in a coccoid, or vegetative, state within the host cells (Trench and Blank, 1987). The coccoid state is defined by the presence of the spherical "pellicle," or cell wall. However, exterior to the cell wall exist multiple layers of apparent membranes. Historically, these layers have been referred to by a variety of terms, including the periplast (Dodge and Crawford, 1970; Kevin et al., 1969; Palincsar et al., 1988; Taylor, 1968; Trench and Blank, 1987), the amphiesma (Schoenberg and Trench, 1980a; Thinh et al., 1986), the theca (Battey, 1992; Taylor, 1971), the vacuolar membranes (Colley and Trench, 1983), or simply the cell covering (Tripodi and Santisi, 1982). The origin of these membranes has also been debated for quite some time.
Tripodi and Santisi (1982), working with the symbiotic octocoral Eunicella stricta, assigned the multiple membranes to the host cell. They stated that the location of recognizable host organelles on the "inner side" of the membranes indicated that, in this host species, the membranes were composed of host plasmalemma. They also stated that the "many times folded" plasmalemma would have an increased surface area and thus could have a functional significance; however, they did not relate what this might be, nor did they indicate how the plasmalemma would accomplish this folding process (Tripodi and Santisi, 1982). Palinscar et al. (1988), working with Aiptasia pallida, also assigned the multiple membranes to the host but suggested that they were formed from internal host cell membranes rather than from the plasmalemma. They hypothesized that membrane-bound cytoplasmic extensions of host tissue could envelope the symbiont, followed by the extrusion of the cytoplasm. This would produce the multiple layers of membrane with no cytoplasmic separations (Palincsar et al., 1988). Other authors, including Colley and Trench (1983) and Rands et al. (1993), have also assigned the origin of the multiple membranes to the host cell.
On the other hand, Kevin et al. (1969), Schoenberg and Trench (1980b), Trench and Blank (1987), and Muller-Parker et al. (1996) have identified a single host-derived membrane and assigned the origin of the multiple layers of membrane to the algal symbiont. Most do so without postulating a mechanism for how these membranes might be produced; however, Trench and Blank (1987) have suggested one possibility. They stated that the cell wall of symbionts is covered by an acidic polysaccharide or glycoprotein layer that resembles a membrane at the ultrastructural level, but in fact contains no lipids. They further suggested that this layer is occasionally sloughed off the outer surface of the cell wall. As these layers accumulate, they appear as multiple membranes exterior to the cell wall and produce the peripheral scroll-like layers of apparent membrane often seen in ultrastructural sections (Trench and Blank, 1987).
In earlier papers (Wakefield et al., 1998, 2000), we also suggested that the membranes were derived from algal cells, but we proposed a very different mechanism for the production of these membranes than that put forth by Trench and Blank (1987). Briefly, we hypothesized that the algal symbionts were continuing through their normal ecdysis cycle within the host cell vacuole. By cycling through the coccoid-to-thecate stage and back again, a layer of apparent multiple membranes would accumulate between the host cell symbiosome membrane and the symbiont. The membranous layers would be composed of the sloughed plasma membrane, thecal vesicles, and plates from each ecdysis event. Therefore, we feel that any assignment of the multiple layers of membrane to the host (with the exception of the single outermost symbiosome membrane) is unsubstantiated and probably in error.
If the host-derived origin for the membranes were correct, then each and every membrane in the layer could be defined as the "symbiosome membrane," as defined by Roth et al. (1988). However, the results of labeling with our PC3 antibody indicate that the multiple layers of membrane lying between the host vacuole membrane and the symbiont cell wall originate from the algal symbiont and are held in place by the presence of a single host-derived membrane. Therefore, the host-derived membrane is the definitive symbiosome membrane, while the other apparent membranes could be defined as symbiont-derived "metabolic components" within the symbiosome space (see Roth’s [1988] original definition of the symbiosome). However, designating these membranes as mere "metabolic components" may belie their potential importance to the symbiotic system. In subsequent text, the term "symbiont membranes" is used to describe these structures.
On the basis of the presence of the PC3 antigen within the symbiont membranes and the cytoplasm, it may be hypothesized that, during the ecdysis process, cytoplasmic material containing the PC3 antigen is also shed and becomes trapped between the layers of plasmalemma and thecal vesicles described by Wakefield et al. (2000). Another possibility is that the PC3 antigen is simply a substance exuded through the cell wall during the coccoid stage of the life cycle. This material could accumulate and become trapped between the layers of shed plasmalemma and thecal vesicles during ecdysis. Alternatively, if Trench and Blank (1987) are correct, exuded PC3 antigen would become trapped between the layers of polysaccharide or glycoprotein material that is occasionally sloughed from the outer cell wall.
Although the identification of the PC3 and G12 antigens is still under investigation in our laboratory, recent data indicate that the PC3 antigen is, in fact, an exuded material (Wakefield and Kempf, unpubl. data). Markell and Trench (1993) have demonstrated that cultured symbionts do exude large water-soluble glycoconjugates. If this exudate is produced in situ (in the symbiotic state), then these molecules could provide essential nutrients to the host (provided the host could digest them), and might also serve as molecular signals between the host and the symbiont (Markell and Trench, 1993).
The potential of the symbiont membranes (or trapped exudates) to act as molecular signals is suggested by the findings of Colley and Trench (1983). They performed a series of experiments in which freshly isolated symbiosomes from Cassiopeia xamachana, Aiptasia pallida, A. pulchella, Anthopleura elegantissima, and Zoanthus sp. were introduced into the gastrovascular cavity of aposymbiotic scyphistomae of C. xamachana. They also introduced cultured algae isolated from the cnidarian hosts C. xamachana, A. pallida, A. tagetes, and Zoanthus sociatus. They found that all cultured algae, even those that were specific for C. xamachana, were either completely rejected or phagocytosed at almost immeasurably low rates. By contrast, the freshly isolated symbiosomes from each host species were all phagocytosed at relatively rapid rates. When viewed with the transmission electron microscope, all of the freshly isolated symbionts were surrounded by multiple layers of apparent membrane. These membranes were completely absent from the cultured algal cells. After several experiments, the authors concluded that a recognition process must be occurring at the surface between the host endodermal cell and the specific membrane layers that surround the algal symbiont (Colley and Trench, 1983).
Believing that there were no algal cell membranes exterior to the cell wall, Colley and Trench (1983) assigned the origin of these membranes to the host and called them "vacuolar membranes." They hypothesized that the enhanced uptake of freshly isolated symbiosomes was a result of the presence of these host membranes, and that there must be a specific molecular factor involved. Since the host endodermal cell was making its initial contact with various "host membranes" of injected symbiosomes, they concluded that there was no actual intercellular (host–symbiont) recognition occurring, and that the uptake of symbiosomes might be indistinguishable from the phagocytosis of food particles.
As we have clearly demonstrated in this study, in Aiptasia pallida there are a multitude of symbiont-produced membranes or membrane-like layers that surround the algal cell wall and are associated with high concentrations of the symbiont-derived PC3 antigen. We postulate that the rapid uptake of isolated symbiosomes compared to the low uptake of cultured algae observed by Colley and Trench (1983) may have resulted from the absence of the correct molecular signal at the external surface of most cultured algal cells. Since our results reveal the PC3 antigen to be absent from most (but not all) suspended, cultured algal symbionts from A. pallida and A. pulchella and present in all symbiosomes in situ, it is possible that such an antigen may be the actual signal that stimulated rapid phagocytosis in the experiments of Colley and Trench (1983). Similarly, since cultured symbionts lack the multiple layers of symbiont membranes found in situ, it is also possible that these membranes may contain the molecular signal that stimulates phagocytosis. We feel that it would be useful to reexamine the question of host recognition of algal symbionts with attention to these possibilities.
It is unknown how aposymbiotic cnidarian offspring that must recruit their symbionts from the environment achieve an initial infection (Berner et al., 1993; Davy et al., 1997; Schwarz et al., 1999); however, there is some support for the hypothesis that the motile thecate stage of the symbiont rather than the sessile coccoid stage is capable of infecting (Kinzie, 1974). If the plasmalemma of the thecate stage contains the appropriate molecular signal, these algal cells would be readily phagocytosed, thus resulting in the initial infection. If such signal molecules can stimulate phagocytosis, then other host- or symbiont-specific molecules must be involved in maintaining the symbiosis once it is established (Trench, 1993). Although Colley and Trench (1983) found that all of the freshly isolated symbionts were phagocytosed by Cassiopeia xamachana scyphistomae, only Symbiodinium microadriaticum and S. bermudense were ultimately maintained in the endodermal cells and could stimulate strobilation. Therefore, the signals that stimulate phagocytosis could not be the same as those that maintain the symbiosis.
Host–symbiont-specific signaling has been shown to occur in many other symbiotic associations such as the Rhizobium–legume association (Lerouge et al., 1990; Sanchez et al., 1991), pathogenic-bacteria–plant interactions (Dixon and Lamb, 1990; Regensburg-Tuink and Hooykaas, 1993), Mycobacterium–macrophage invasions (Ferrari et al., 1999), X-bacteria–amoeba symbiosis (Jeon, 1992, 1997), parasite–host infections (Sibley et al., 1986; Sibley and Krahenbuhl, 1988; Perkins, 1989; Sam-Yellowe, 1992; Collins et al., 1997), and diatom–foraminifera interactions (Lee et al., 1988, 1997; Lee, 1998; Chai and Lee, 1999, 2000). In many of these associations, the specific signals leading to successful symbiotic interactions have been elucidated. Agrobacterium tumefaciens, a bacterium that induces tumors in many types of plants, is dependent upon the VirF protein for tumor formation (Regensburg-Tuink and Hooykaas, 1993). Plasmodium falciparum uses several proteins, including the 155-kDa ring-infected surface antigen and the 140/130/110/-kDa rhoptry complex, to bind to appropriate erythrocyte membranes (Sam-Yellowe, 1992). Rhizobium meliloti uses a ß-1,4-tetrasaccharide of D-glucosamine to induce root nodules in alfalfa (Lerouge et al., 1990). Mycobacterium species use the TACO protein to inhibit lysosomal fusion in macrophages (Ferrari et al., 1999). The frustules of all endosymbiotic diatoms of large foraminifera display a 104-kDa polypeptide that prevents their digestion by the host (Chai and Lee, 1999).
Attempts to locate and characterize similar types of molecular signals required for establishing and maintaining zooxanthella–cnidarian symbioses have met with little success; however, the two reported attempts to identify these signals concentrated on the algal cell wall (Markell et al., 1992) and on water-soluble glycoconjugates released by algal cells in culture (Markell and Trench, 1993). Because we have identified the symbiont membranes to be of algal-cell origin and intimately associated with a symbiont-derived antigen (PC3), and because an earlier investigation (Kinzie, 1974) supports the hypothesis that molecular signals associated with the thecate motile stage must stimulate phagocytosis, it may be hypothesized that these symbiont membranes are involved in this signal transduction event. It is likely that knowledge of the identity and function of such molecular signals will be critical to our understanding of the cellular mechanisms orchestrating events such as coral bleaching (Bunkley-Williams and Williams, 1990; Ghiold, 1990; Gleason and Wellington, 1993; Goreau and Hayes, 1994; Brown et al., 1995), where such signaling capabilities have apparently been lost.
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