Identification of unstimulated constitutive immunocytes, by enzyme histochemistry, in the coenenchyme of the octocoral Swiftia exserta.
Aside from theoretical interest in the subject, there have been recent suggestions that disease is a significant factor in widespread coral reef decline (e.g., Rogers, 2009). Thus far, more efforts have been expended in identifying the putative coral pathogens than the coral host response. There exists a need, in particular, for more functional information at the cellular and histological levels to move forward (Rinkevich, 2012). Describing the constitutive state of an animal is crucial before differentiating a response due to stimulation (Sparks, 1972).
The effectors of immune systems traditionally have been categorized into two groups: the humoral and the cellular. In most animals, the cellular effectors circulate and are thus readily apparent and easily isolated. The white blood cells from humans, other mammals, and other vertebrates have been the subject of intense investigations over an extended period (reviewed in Hayhoe and Quaglino, 1994); in the last few decades, many specialized markers, reagents, and techniques have been developed. Unfortunately, many of these have not yet been adapted for use in anthozoan cnidarians. Studies with some invertebrate blood cells, such as echinoderm coelomocytes, have an even longer history (Metchnikoff, 1893), but generally have not been accorded the same dedication of resources. Of the invertebrate hemocytes and coelomocytes (taxa-specific terms for leukocyte-like cells) that have been studied, some cells of arthropod (e.g., Culex pipiens (Wang et al., 2011); Carcinus maenas (e.g., Soderhall and Smith, 1983); Limulus polyphemus (Copeland and Levin, 1985)); mollusc (Mytilus edulis (Pipe, 1990); Crassostrea virginica (Cheng and Downs, 1988); Chamelea gallina (Pampanin et al., 2002)); protochordate (e.g., Ballarin and Cima, 2005); and annelid (e.g., Stein and Cooper, 1978) have been characterized. Those studies focused on cellular functions (Chemotaxis and phagocytosis) and on cellular contents (e.g., hydrolases, (per)oxidases, lysozyme; Soderhall and Smith, 1983).
Since Metchnikoff's (1893) classic demonstration of a foreign-body response in Scyphozoa (translucent jellyfish), we have made little additional progress in identifying the primary immune cells in cnidaria in general--hydrozoa, anthozoa, schyphozoa, and cubozoa. By inserting a carmine-soaked splinter into the mesoglea of a scyphozoan and describing the resultant accumulation of amoebocytes around this large foreign body that phagocytized the carmine, Metchnikoff pioneered the field of cellular immunology.
Although phagocytosis is an innate capability of many cell populations, higher animals have evolved a set of professional phagocytes that circulate throughout the organism to capture and digest, or capture and encapsulate (sequester) invading cells--microbes, conspecific non-self, or parasites.
Phagocytic immunocytes carry a range of enzymes to kill and subsequently digest their ingested targets, including bacteria, yeast, protozoa, and eukaryotic cells. Human neutrophils, in particular (e.g., Hayhoe and Quaglino, 1994; Nauseef and Borregard, 2014), have been studied extensively, and several reviews list granule contents for the main granule types (e.g., Borregard et al., 1993). Colorimetric and heavy-metal deposition reactions have been developed for several of the enzymes involved in killing microbes (e.g., lysozyme and peroxidase) as well as for an array of leukocyte-specific hydrolytic enzymes (Pearse, 1968, 1972; Bancroft and Hand, 1987). The phagolysosome in the phagocytes of coelomate animals contains a variety of hydrolases, including phosphatases (alkaline and acid), lipases and esterases with varying substrate specificity, and an array of substrate-specific glycosidases (reviewed in: Borregard et al., 1993; Hayhoe and Quaglino, 1994). Different phagocyte enzymes are activated or reach optimal activity rates during phagosome maturation as the phagolysosomal pH decreases from slightly alkaline to acidic (7.8 to below 5). These enzymes degrade and digest microbes and other ingested particles. Linking enzyme activity (biochemistry) to a cellular or tissue location (morphology) has a long-established history (e.g., Hardonk and Koudstaal, 1976), and has been named "enzyme histochemistry."
Lysosomal enzyme markers have been used extensively to characterize different blood cell types (hemocytes, coelomocytes, or leukocytes) in vertebrates (e.g., Garavini et al., 1981; Hayhoe and Quaglino, 1994; Nauseef and Borregard, 2014) and many invertebrates (e.g., Yoshino and Cheng, 1976; Stein and Cooper, 1978; Soderhall and Smith, 1983; Copeland and Levin, 1985; Ballarin and Cima, 2005; Wang et al., 2011). It has generally been assumed that all members of a particular cell population will stain for their characteristic enzymes. Most quantitative analyses of cellular lysosomal enzyme content, however, have measured enzymatic activities of cytolysates of pooled cells (or tissues) expressed on a per-cell or per-milligram-of-protein basis (e.g., Papadimitiou and Wyche, 1976; Mydlarz et al., 2008).
In many invertebrates, the cellular immune response includes encapsulation of the foreign body in an attempt to isolate the invading material from the host's tissue. During this process, phagocytic cells wrap themselves around the invader in many layers while secreting enzymes and antimicrobials (peptides and other compounds). The capsule is firmly held together by cross-linked melanin produced by the enzyme phenoloxidase from tyrosine and dihydroxyphe-nylalanine (DOPA). The (pro)phenol oxidase system has been studied extensively in marine invertebrates (e.g., Smith and Soderhall, 1991). Reports by Mydlarz and coworkers have found several phenol oxidase isotypes in tissue extracts of Caribbean octocorals (Mydlarz et al., 2008; Mydlarz and Palmer, 2011), indicating that this enzyme is evolutionarily ancient.
Our model animal, the octocoral Swiftia exserta Duchassaing & Michelotti, 1864, is a sessile colonial octocoral that divides from a single stem into many branches at irregular intervals. Polyps are eight-tentacled and are embedded in a column of fleshy tissue, the coenenchyme, at their base, with the polyp's gastric cavity extending into the coenenchyme. The animal's structural support is a hollow protein skeleton. Polyp gastric cavities, the main sites of digestion, are lined with mesentery filaments that extend from the tentacle bases into the gastric cavity. Mesenteries are attached to the external polyp wall on one side only. Individual gastric cavities are interconnected through the coenenchyme by the solenia, or tube-shaped canals. The gastroderm tissues of the mesenteries and the solenia are separated from the surrounding mesoglea by collagen-rich basal lamina (Menzel et al., 2015), and are thus separate niches from the mesoglea. Commonly, cnidaria are considered diploblastic animals, with a cellular ectoderm and endoderm (gastroderm) separated by an acellular mesoglea (Brusca and Brusca, 2003).
Cnidaria such as Swifiia exserta lack a cellular vascular system but have amoeboid-shaped cells, termed "granular amoebocytes," which have been reported in the mesoglea (reviewed in Bigger and Hildemann, 1982). Granular amoebocytes have been described in anthozoan wound infiltrates (Young, 1974; Meszaros and Bigger, 1999) and graft rejection areas (Olano, 1993). Based on different metachromatic staining, granular amoebocytes are not thought to be a homogeneous population (Bigger and Hildemann, 1982). Some of these cells are phagocytic (Olano and Bigger, 2000), and thus have been viewed as putative "immunocytes" in these cnidarians. The term "granular amoebocyte" in S. exserta is misleading, however; there are cell types that are clearly granular and cell types that have irregular outlines (some even have granules!), but amoeboid motion cannot be observed since living animals are heavily pigmented and the tissues are opaque. A recent report illustrates the histology and ultrastructure of S. exserta's coenenchyme (Menzel et al., 2015). Since Cnidaria lack a circulatory system, they cannot simply be bled to obtain putative immunocytes. Moreover, aside from showing the existence of phagocytic cells in S. exserta, Olano and Bigger (2000) demonstrated that trauma or injury changes the phagocytic characteristics of cell populations in S. exserta.
To study constitutive levels of unstimuated phagocytic cells, we used functional proxies to identify the unstimulated immunocytes present in "resting" animals. Using unstimulated animals, we aimed to identify the cells that would initiate the phagocytic and/or encapsulation response in S. exserta. Since, for many years, hemocytes and leukocytes have been characterized based on their enzyme content (e.g., Janoff and Hawrylko, 1963; Granath and Yoshino, 1984; Gelder and Moore, 1986; Adema et al., 1992; Ballarin et al., 1993; Lopez et al., 1997; Pampanin et al., 2002), we used this approach and the following enzymes due to their colocalization to phagocytic cells: (1) acid phosphatase, (2) alkaline phosphatase, (3) non-specific esterases, (4) [beta]-glucuronidase, (5) peroxidase, and (6) phenoloxidase. The first four enzyme groups are hydrolytic enzymes that aid in intracellular killing and digestion (Borregard et al., 1993; Hayhoe and Quaglino, 1994). Peroxidase is a proxy for the superoxide (respiratory burst)-mediated killing system, and was one of the first enzymes linked to phagocytes (Linossier, 1898; Fischel, 1910). Based on the extensive literature on phenoloxidase function in arthropods (Soderhall and Smith, 1983; Smith and Soderhall, 1991; Sugumaran, 2001; Cerenius and Soderhall, 2004) and the detection of phenoloxidase in cell slurries from a sea fan (Mydlarz et al., 2008; Mydlarz and Palmer, 2011; Pinzon et al., 2014), we set out to determine which cells contain phenoloxidase in our octocoral. We suggest that these cells would also be good immunocyte candidates. The current literature reporting the presence of phenoloxidase in corals is based on tissue slurries (Mydlarz et al., 2008; Mydlarz and Palmer, 2011; Pinzon et al., 2014), and thus does not allow tissue, cellular, or sub-cellular localization within the animals.
Materials and Methods
Maintenance of animals
Colonies of Swiftia exserta were collected by Dr. Henry Feddern, a commercial supplier, at 20-30-m depths off Southeast Florida and maintained in 2080-1 aquaria while at Florida International University. With the relatively low animal tissue mass present and minimal care, basic water conditioning was achieved with sub-gravel and activated charcoal filters, and a protein skimmer. Care was taken to avoid inter-colony contacts between 30 individually tagged colonies. Animals were fed "San Francisco Bay"-brand Artemia nauplii three times weekly (Salter-Cid and Bigger, 1991). Aquarium temperature was maintained between 18-22[degrees]C, and salinity between 35-37 ppt. Animals were acclimated for 2-6 wks before the harvesting of branches for experiments.
This set-up is well suited for long-term use; in other studies with this system (unpubl. data), S. exserta colonies have been maintained for more than a year, with isolated branches adding tissue mass, and, when challenged, engaging in allogeneic tissue rejections without significant change. Accordingly, the tissues used in this study would appear to represent native, normal tissue such as would be present before an immunological challenge.
Cell and tissue processing
Cryoembedding for enzyme histochemistry. Short, 2-3 cm branches were cut (with scissors) from four animals and fixed in 2% paraformaldehyde (Electron Microscopy Sciences, Hatfield, PA) in 0.22-[micro]m filtered seawater (FSW) at room temperature for 4 h. Tissue was infiltrated with increasing concentrations of sucrose (10%, 20%, and two changes of 30%) in 0.22-[micro]m filtered sterile FSW, followed by 30% sucrose:O.C.T. (2:1) (optimal cutting temperature; Tissue-Tek O.C.T. Compound, Sakura Finetek USA, Inc., Torrance, CA); 30% sucrose:O.C.T. (1:1); and three changes of O.C.T. (the last, overnight). Branches were aligned, two vertically and two horizontally, in fresh O.C.T. in plastic boats and gradually immersed in liquid nitrogen. Serial 6-[micro]m sections cut with a Leica CM 1850 Cryo-Stat (Leica Biosystems, Buffalo Grove, IL) at -25[degrees]C were collected on pre-cooled Fisherbrand SuperFrost Plus glass slides (Thermo Fisher Scientific, Waltham, MA) only after the horizontal branches were reached. This action eliminated the branch areas that were traumatized during collection.
Paraffin embedding for histology
Tissues were fixed in Helly's fixative (Clark, 1981), rinsed in FSW, decalcified in 2% ascorbic acid, dehydrated in a graded series of ethanol, and embedded in Paraplast (Leica Biosystems) at 56[degrees]C. Sectioning was performed on a Leica Reichert Jung 2800 Frigocut N Cryostat (Leica Biosystems) at -20[degrees]C, using a metal knife to obtain 5-6-[micro]m sections.
Single-cell suspensions for enzyme cytochemistry
One short branch each was cut (with scissors) from four different animals and washed for 5-10 min in three changes of calcium and magnesium-free artificial seawater containing 30 mmol [1.sup.-1] ethylenediaminetetraacetic acid (EDTA) (Humphreys, 1963). Tissue was scraped onto 50-[micro]m nylon mesh (folded four times) and washed through the mesh in a collection vessel. The resulting single-cell suspension was either (1) used directly for cytospins (5 min at 800 g) or (2) washed three times in artificial seawater containing calcium and magnesium (Peddie et al., 1995) and allowed to adhere to mechanically cleaned glass slides for 45 minutes. Both cytospin and adherent cells were fixed with 4% paraformaldehyde (in FSW) at room temperature for 15 min.
Mallory's aniline blue collagen stain technique (Clark, 1981) was used for general histological staining.
Quinone and quinoprotein detection. Cytospin preparations, adherent cells, and cryosections were incubated with 0.024 mol [1.sup.-1] nitroblue tetrazolium (NBT) in 2.0 mol [1.sup.-1] glycine-NaOH buffer, pH 10, protected from light (Paz et al., 1991). Positive reactions for quinones and quinoproteins yielded a faint blue precipitate.
Peroxidase staining with diaminobenzidine and hydrogen peroxide, and phenoloxidase staining with DL-dihydroxy-phenylalanine (DL-DOPA) was performed as described in Pearse(1968, 1972).
The azo dye coupling methods were used for acid phosphatase and alkaline phosphatase, with naphthol AS-BI phosphate and [alpha]-naphthol phosphate as substrates, and for non-specific esterases, with the substrates [alpha]-naphthyl acetate, naphthyl AS-D chloroacetate, and Tween-80 (DifCo), as described in Bancroft and Hand (1987). For the esterase and [beta]-glucuronidase reactions, the incubation pH was modified: pH values of 5.0, 6.0, and 7.0 were used.
The coupling dyes, Fast Red ITR and Fast Blue RR (dissolved in N,N'-dimefhylformamide), were used since the animal remains a slightly yellowish-orange color after processing. Positive reactions yield an insoluble precipitate, colored depending on the coupling dye used. Acid phosphatase and the non-specific esterase were coupled to Fast Red ITR; alkaline phosphatase and [beta]-glucuronidase were coupled with both dyes. A positive peroxidase reaction results in a reddish-brown precipitate, and a positive phenoloxidase reaction yields a blue-black deposit of melanin. All reagents were purchased from Sigma (St. Louis, MO), unless otherwise specified.
Enzyme specificity controls
Phosphatase inhibitors included 2 mmol [1.sup.-1] sodium orthovanadate, a tyrosine phosphatase, adenosine triphosphatase (ATPase), and a non-specific alkaline phosphatase inhibitor; 20 mmol [1.sup.-1] sodium fluoride, a non-specific acid phosphatase inhibitor; 4 mmol [1.sup.-1] sodium pyrophosphate, an inhibitor of serine and threonine phosphatases; 50 mmol [1.sup.-1] EDTA, which chelates calcium and magnesium essential for phosphatase function; and 250 mmol [1.sup.-1] tetramisole, a specific inhibitor of alkaline phosphatase.
Non-specific esterase activity was inhibited by 20 mmol [1.sup.-1] sodium fluoride.
The peroxidase inhibitors used were 20 mmol [1.sup.-1] sodium orthovanadate, a non-specific peroxidase inhibitor; 20 mmol [1.sup.-1] sodium azide, an irreversible myeloperoxidase inhibitor, which also inhibits superoxide dismutase and catalase; 20 mmol [1.sup.-1] sodium cyanide, a reversible inhibitor of cytochrome c oxidase; and a complete inhibition of peroxidase activity with 3% methanolic hydrogen peroxide.
Reactions were repeated three times independently with freshly prepared solutions each time (technical replicates), on four different animals each (n = 12).
Histological staining in paraffin sections
Cytochemical staining of paraffin sections with Periodic acid-Schiff (PAS) reagent stained every part of the animal, including the mesoglea, in varying shades of purple. Feulgen's stain similarly colored the sections completely (less intensely than PAS). Data are not shown.
Mallory's aniline-blue collagen stain (Clark, 1981) dyes paraffin sections intensely (Fig. 1B). In Mallory-stained sections, cell nuclei are colored orange, collagenous fibers are intensely blue, mucus-containing granules appear in shades of blue, and other fibrous material, in red (see also Menzel et al., 2015).
To introduce the reader to Swiftia exserta, Figure 1A shows a (live) branch; Figure 1B illustrates a section through coenenchymal tissue stained with Mallory's aniline blue connective tissue stain (to show the overall tissue organization of the animal); and Figure 1D shows an unstained cryosection (as the basis for interpreting the sections stained by enzyme histochemistry). The coenenchyme in 5. exserta is loosely organized into a richly cellular mesoglea, in which the mesogleal fibers stain intensely blue, indicating their collagenous nature; a very thin external ectoderm; and a layer of gastroderm that is only a single columnar cell layer thick. This gastroderm is supported with an external (blue) collagenous layer and an internal (red), fibrin-like layer (similar to the gastroderm of the solenia (Menzel et al., 2015)). It has a few intensely acidophilic globular gland cells randomly interspersed with poorly staining, heavily granular cells. Many cells in the highly folded mesentery filaments of the gastric cavity stain intensely acidophilic, and a series of parallel myonemes is visible. Open spaces in the mesoglea are from (partially) decalcified spicules.
Even though the animal is highly pigmented in life (Fig. 1 A), the processing for cryosectioning removed much of the pigment (Fig. 1D). In the cryosection (Fig. 1D), parts of the mesentery filaments and the gastric cavity are visible towards the top of the section. A lining of solenia-type gastroderm delimits the mesoglea from the gastric cavity, while a thin ectoderm (at the bottom of Fig. 1D) separates the mesoglea from the external environment. The cryosections retain only a small amount of the animal's natural orange-brown pigment after the processing for cryosectioning.
The NBT-quinone reaction is strongly positive in cells with few and very large round granules. Weakly positive quinoprotein (blue) staining is evident in amorphous hyaline cells with granules interspersed in the cytoplasm (see Table 1). In cryosections, quinones and quinoproteins stained blue in the outermost ectodermal epithelium cells and in oblong cells with large granules in the mesogleal cell cords (Fig. 1C). The positive cells in the mesoglea often stained very intensely, indicating a high concentration of quinones and quinoproteins.
Phosphatases. The alkaline phosphatase reactions were negative for freshly prepared cells and cryosections, with two different substrates (naphthol AS-BI phosphate and [alpha]-naphthol phosphate). However, the acid phosphatase reaction stained several cell types in both adherent and cytospin cell preparations (see Table 1) and several cell types in cryosections (Fig. 2A): irregularly shaped cells with large granules and amorphous hyaline cells with granules interspersed in the cytoplasm (Fig. 2A, arrowheads). Figure 3 shows several inhibitor controls specific for acid phosphatases in cryosections. Sodium orthovanadate (Fig. 3A), which inhibits tyrosine phosphatase, ATPase, a non-specific alkaline phosphatase, and the non-specific acid phosphatase inhibitor sodium fluoride (Fig. 3B), inhibited all acid phosphatase activity. The serine and threonine phosphatase inhibitor sodium pyrophosphate (Fig. 3C), and the calcium and magnesium chelator EDTA (Fig. 3D) restricted positive staining to the solenia gastroderm cells containing large round granules and many gland cells in the mesentery gastroderm. Reactions performed without substrate convey only the residual pigment in the animal (as in Fig. 1D).
Esterases. In the fresh cell preparations, the heavily granular cells stained positive with [alpha]-naphthyl acetate in both cytospin and adherent cell populations (see Table 1); sodium fluoride inhibited this reaction completely. In cryosections, naphthyl AS-D chloroacetate and Tween-80 did not yield any colorimetric reactions at any of the three pHs, whereas [alpha]-naphthyl acetate yielded strongly positive results at pH 7.0 only (Fig. 2B). The inhibitor, sodium fluoride, significantly reduced staining in the sub-epithelial layer and inhibited the enzyme isotype in the gastroderm (both solenia and mesentery gastroderm) (Fig. 4A).
[beta]-glucuronidase. Neither cryosections, paraffin sections, nor cell suspensions stained for the enzyme [beta]-glucuronidase under any of the three pH conditions tested: pH 5.0, 6.0, and 7.0.
Oxidases. Both oxidases showed distinct positive reactions in cytospin and adherent cell preparations (see Table 1), both oxidases labeling similar cell types: spherical cells filled with small, round granules, and amorphous hyaline cells with granules interspersed in the cytoplasm. Peroxidase-positive cells in cryosections (Fig. 2C) were localized mainly in the granular cell layer immediately below the ectodermal epithelium, with some cells in the mesogleal cell cords also staining positive (Fig. 2C, arrowheads). These mesogleal cells closely resemble the granular amoebocytes described in Menzel et al. (2015). Occasional mesentery gland cells also stained. Several control reactions for peroxidase were included. A non-specific inhibitor, sodium orthovanadate, only allowed staining in the occasional mesentery gastroderm gland cells (data not shown), but eliminated staining in the mesogleal cells. Sodium azide (a cytochrome c inhibitor) had little effect on the peroxidase staining pattern (Fig. 4B). However, sodium cyanide, an inhibitor for myeloperoxidase, catalase, and superoxide dismutase, abrogated the peroxidase staining (Fig. 4C); a complete inhibition of peroxidase activity with 3% methanolic hydrogen peroxide was seen (complete inhibition, as expected; Fig. 4D). Cells staining positively for phenoloxidase in cryosections (Fig. 2D) were restricted to the cells within the mesoglea (Fig. 2D, arrowheads); neither the ectoderm nor cells of the gastroderm reacted for phenoloxidase. Though they were not as numerous as the oblong granular cell population described, these phenoloxidase-positive cells resembled oblong granular cells, as reported by Menzel et al. (2015)--perhaps they define a subpopulation within oblong granular cells.
Some cells of the cnidarian mesoglea historically have been reported to include putative immunocytes (Chapman, 1974; reviewed in Bigger and Hildemann, 1982) in the form of "granular amoebocytes." Because these immunocytes have been described as phagocytic, they should be involved in any foreign body response and wound healing, and thus carry antimicrobial substances (enzymes, peptides, and oxidative burst capacity). Here we localize a number of enzymes associated with white blood cells (leukocytes) (e.g., Janoff and Hawrylko, 1963; Granath and Yoshino, 1984; Gelder and Moore, 1986; Adema et al., 1992; Ballarin et al., 1993; Lopez et al., 1997; Pampanin et al., 2002) to the layer of amorphous, highly granular cells just beneath the squamous ectodermal epithelium, and in similar cells dispersed within the deeper mesoglea in Swiftia exserta. The mesoglea and ectoderm are physically separated from the gastroderm (both solenia and mesentery) by a basal membrane (Menzel et al., 2015).
This is the first report to localize any enzymes in an octocoral, and the first to detect enzymes in the clonal tissue between polyps in any cnidarian (hard coral, soft coral, or hydroid). Hydrolytic enzymes, such as acid phosphatase and several esterases, are concentrated in granule-packed cells immediately below the thin ectoderm epithelial cells. In contrast, the oxidases and quinones (readily available reactive oxygen) are found primarily in the granular cells deeper in the mesoglea. This observation augments the findings of Olano and Bigger (2000), who described two populations of phagocytic cells: a smaller population determined 2 h post-injury and a large population of cells 24 h post-injury (cut or thread insertion). These authors noted that injury dramatically altered the cell types capable of phagocytosis--histology on 2-h, post-injury sections showed that "granular amoebocytes" were the sole phagocytic cell type, while at 24 h post-injury, internalized particles were detected in many cell types (ectoderm, gastroderm, sclerocytes, "granular amoebocytes," and mesogleal cells). Cells previously labeled "mesogleal cells," "globular granular cells," and "granular amoebocytes" in that earlier report (Olano and Bigger, 2000), were reclassified in Menzel et al. (2015); they are labeled "oblong granular cells" here in accordance with the latter report.
The solenia-type gastroderm in the coenenchyme has many acid phosphatase-positive cells, indicating the likely presence of acidic vesicles (possible lysosomes). Some cells in the solenia also stain for non-specific esterases; however, the lack of phenoloxidase staining in the solenia-type gastroderm and in the mesenteries suggests a digestive function for these gastroderm tissues, since phenoloxidase is associated with immune reactions in invertebrates. In addition, we must stress the physical barrier separating these gastroderm cells from the rest of the animal by a clear basal lamina, visible by light microscopy in Figure 1B and transmission electron microscopy (Menzel et al., 2015). As a result of this robust barrier to cell migration between the gastoderm and the mesoglea, we assign distinct roles to these tissues. We conclude that the cells of the gastroderm (solenia and mesentery) are the sole nutritive cells of the animal (several phagocytic vesicles are documented by Menzel et al. (2015) in the solenia gastroderm, but none in the mesoglea or the ectoderm). However, we consider the cells of the mesoglea and the ectoderm to include potential immunocytes. In additional support of this conclusion, we note that the cell populations of the gastroderm are also morphologically very distinct from the cell populations of the mesoglea and ectoderm (Menzel et al., 2015), and are thus serving different functions.
Many of the oblong granular cells in the mesoglea stain positive for acid phosphatase, acetate esterase, and oxidases, all of which have served as markers for "professional phagocytes" in mammals (neutrophils; Borregard et al., 1993) and in many invertebrate hemocytes (e.g., Janoff and Hawrylko, 1963; Granath and Yoshino, 1984; Gelder and Moore, 1986; Adema et al., 1992; Ballarin et al., 1993; Lopez et al., 1997; Pampanin et al., 2002). The scattered, non-specific esterase, peroxidase, phenoloxidase, and quinone-positive cells infused throughout the cell cords in the mesoglea seem ideally situated to migrate to sites of need. We also observed that many of the acid phosphatase, nonspecific esterase, peroxidase, and phenoloxidase-positive cells adhere to mechanically cleaned glass slides, suggesting that these cells are capable of phagocytosis (Papadimitriou and Wyche, 1976).
The overlap in cellular shape, structure, and location strongly supports the notion that many mesogleal cells can function as immunocytes, but that the thin ectoderm layer does not harbor phagocytic immunocytes (Menzel, 2013; Menzel et al., 2015), since no phagosomes were detected in the ectoderm. Based on these experiments, we can only speculate that other immunocyte types may be present. Hutton and Smith (1996) reported the isolation of "amoebocytes" from the mesenteric filaments (tissue including gastroderm and mesoglea) of the sea anemone Actinia equina that are capable of phagocytosis and killing gram-negative bacteria.
In addition to the proposed immunocytes in the mesoglea, we propose that quinones, potent redox-capable molecules, in the outermost ectodermal epithelium may provide a first line of defense against invading pathogens susceptible to oxidative damage.
Due to the activation cascade required for phenoloxidase (e.g., Sugumaran, 2001; Cerenius and Soderhall, 2004), the phenoloxidase reaction is much stronger in the single-cell suspensions than in the cryosections. During the separation procedure, these cells were shocked by calcium and magnesium withdrawal, and by mechanical forces (pressure and shear) before being tested for phenoloxidase activity.
A clustering of enzyme colocalization (acid phosphatase and esterases vs. hydrolytic plus peroxidase and phenoloxidase) suggests the presence of at least two cell populations involved in cellular innate immune responses: a layer in the immediate exterior area where likely incursions will have penetrated the thin ectoderm, and a wandering cell type responding to deeper injury, which may encapsulate offending material. In addition, the thin ectoderm may use quinones or quinoproteins as a first line of defense. Research on the morphology, histology, and ultrastructure of Swiftia exserta have recently been published from our laboratory (Menzel et al., 2015).
As noted by Metchnikoff (1893), and more recently by Bosch's group (Augustin and Bosch, 2010), hydrozoa are morphologically and--in their immune responses--quite different from anthozoa. Augustin and Bosch (2010) reviewed the potential immune responses in Hydra spp. and concluded that in this simple cnidarian, the immune reactions are limited to focal responses in the ectoderm or the endoderm (also known as gastroderm). In Hydra spp. (and hydrozoa), these tissue layers are only one cell thick and are separated by a very thin acellular mesoglea. S. exserta and perhaps other anthozoa seem to have at least three cell populations equipped to defend tissue integrity: initial-encounter cells in the ectoderm epithelium, which can chemically oxidize invading organisms; the focal response cells immediately sub-epithelial; and the migrating DOPA-oxidase-containing cells deeper in the mesoglea. These findings do not preclude the existence of other cells that may serve defensive or immunological functions.
Thus, the cells immediately below the ectoderm epithelium contain at least two cell types by function: 1) immediate-responder defensive cells, and 2) secondary-responder defensive cells. Immediate-responder cells consist of the granular cells beneath the ectodermal epithelium that contain acid phosphatase and esterase, but not the oxidases. This staining pattern suggests that these cells could serve in the uptake of dissolved organics (Mariscal and Bigger, 1977) and/or in host defense. The secondary-responder cells are represented by the oblong granular cells deeper in the mesoglea, which contain acid phosphatase, esterase, peroxidase, and phenoloxidase. Colocalization of all of these enzymes indicates that an immune function takes precedence in these cells.
Identification of the cells that are constitutively ready for an initial immune response is an important first step towards understanding the "resting" state of the anthozoan immune system. It also provides a firmer foundation on which to begin investigations into the changes that occur during responses to immunological challenges and injuries. Based on earlier studies (Bigger, 1984; Salter-Cid and Bigger, 1991; Meszaros and Bigger, 1999; Olano and Bigger, 2000), the changes potentially involve multiple cell types, cell signaling, gene activation, and changes in cellular processes leading to the effector phases. Many new techniques are being adapted for use in studies with these anthozoans, but a proper identification of anatomy and resting cell states is critical, as pointed out by Albert Sparks (1972) and Baruch Rinkevich (2012).
LPM thanks Cecile Tondo (Olano) for excellent paraffin sections, and the FIU Tissue Culture Core Facility for use of the Leica CM 1850 Cryo-Stat.
Adema, C. M., R. A. Harris, and E. C. van Deutekom-Mulder. 1992. A comparative study of hemocytes from six different snails: morphology and functional aspects. J. Invertebr. Pathol. 59: 24-32.
Augustin, R., and T. C. G. Bosch. 2010. Cnidarian immunity: a tale of two barriers. Adv. Exp. Med. Biol. 708: 1-16.
Ballarin, L., and F. Cima. 2005. Cytochemical properties of Botryllus schlossen haemocytes: indications of morpho-functional characterization. Eur. J. Histochem. 49: 255-264.
Ballarin, L., F. Cima, and A. Sabbadin. 1993. Histoenzymatic staining and characterization of the colonial ascidian Botryllus schlossen hemocytes. Boll. Zool. 60: 19-24.
Bancroft, J. D., and N. M. Hand. 1987. Enzyme Histochemistry. Oxford University Press, Oxford.
Bigger, C. H. 1984. Immunorecognition among invertebrates. Dev. Comp. Immunol. 3: 29-34.
Bigger, C. H., and W. H. Hildemann. 1982. Cellular defense systems of the Coelenterata. Pp. 59-87 in The Reticuloendothelial System, N. Cohen and M. Sigel, eds. Plenum Press, New York.
Borregard, N., K. Lollike, L. Kjeldsen, H. Sengel0v, L. Bastholm, M. H. Nielsen, and D. F. Bainton. 1993. Human neutrophil granules and secretory vesicles. Eur. J. Hematol. 51: 187-198.
Brusca, R. C, and G. J. Brusca. 2003. Invertebrates. Sinauer, Sunderland, MA.
Cerenius, L., and K. Soderhall. 2004. The prophenoloxidase-activating system in invertebrates. Immunol. Rev. 198: 116-126.
Chapman, D. M. 1974. Cnidarian histology. Pp. 2-92 in Coelenterate Biology: Reviews and New Perspectives, L. Muscatine and H. M. Lenhoff, eds. Academic Press, New York.
Cheng, T. C, and J. C. Downs. 1988. Intracellular acid phosphatase and lysozyme levels in subpopulations of oyster, Crassostrea virginica, hemocytes. J. Invertebr. Pathol. 52: 163-167.
Clark, G. 1981. Staining Procedures. Williams and Wilkins, Baltimore.
Copeland, D. E., and J. Levin. 1985. The fine structure of the amoebocyte in the blood of Limulus polyphemus. I. Morphology of the normal cell. Biol. Bull. 169: 449-457.
Fischel, R. 1910. Der histochemische Nachweis der Peroxydase. Wien. Klin. Wochenschr. 44: 1557.
Garavini, C, P. Martelli, and B. Borelli. 1981. Alkaline phosphatase and peroxidase in neutrophils of the catfish Ictalurus melas (Rafinesque) (Siluriformes, Ictaluridae). Histochemistry 72: 75-81.
Gelder, S. R., and C. A. Moore. 1986. Cytochemical demonstration of several enzymes associated with phagosomal processing of foreign material within hemocytes of Mercenaria mercenaria. Trans. Am. Microsc. Soc: 105: 51-58.
Granath, W. O., and T. P. Yoshino. 1984. Intracellular distribution of lysosomal enzymes within the hemocytes of Biophalaria glabrata. Trans Am. Microsc. Soc. 103: 38-43.
Hardonk, M. J., and J. Koudstaal. 1976. Enzyme histochemistry as a link between biochemistry and morphology. Prog. Histochem. Cytochem. 8: 1-66.
Hayhoe, F. G. J., and D. Quaglino. 1994. Haematological Cytochemistry, 3rd ed. Churchill Livingstone, London.
Humphreys, T. 1963. Chemical dissolution and in vitro reconstruction of sponge cell adhesions. I. Isolation and functional demonstration of the components involved. Dev. Biol. 53: 27-47.
Hutton, D. M. C, and V. J. Smith. 1996. Antibacterial properties of isolated amoebocytes from the sea anemone Actinia equina. Biol. Bull. 191: 441-451.
Janoff, A., and E. Hawrylko. 1963. Lysosomal enzymes in leucocytes of Mercenaria mercenaria and Asterias forhesi. Biol. Bull. 125: 381.
Linossier, M. G. 1898. Contribution a l'etude des ferments oxydants, sur la peroxydase du pus. C. R. Hedb. Soc. Biol. (Paris) 50: 373-375.
Lopez, C, M. J. Carballal, C. Azevedo, and A. Villalba. 1997. Enzyme characterisation of the circulating haemocytes of the carpet shell clam, Ruditapes decussatus (Mollusca: Bivalvia). Fish Shellfish Immunol. 7: 595-608.
Mariscal, R. N., and C. H. Bigger. 1977. Possible ecological significance of octocoral epithelia ultrastructure. Pp. 127-133 in Proceedings of the Third International Coral Reef Symposium, D. L. Taylor, ed. Rosenstiel School of Marine and Atmospheric Science, Miami, FL.
Menzel, L. P. 2013. Aspects of the innate immune system of the Caribbean octocoral Swiftia exserta. Ph.D. dissertation, Florida International University, Miami. FL.
Menzel, L. P., C. Tondo, B. Stein, and C. H. Bigger. 2015. Histology and ultrastructure of the coenenchyme of the octocoral Swftia exserta, a model organism for innate immunity/graft rejection. Zoology 118: 115-124.
Meszaros, A., and C. H. Bigger. 1999. Qualitative and quantitative study of wound healing processes in the coelenterate, Plexaurella fusifera: spatial, temporal, and environmental (light attenuation) influences. J. Invertebr. Pathol. 73: 321-331.
Metchnikoff, E. 1893. Lectures on the Comparative Pathology of Inflammation: Delivered at the Pasteur Institute in 1891. Kegan Paul, Trench, Trubner & Co., London.
Mydlarz, L., and C. V. Palmer. 2011. The presence of multiple phenol oxidases in Caribbean reef-building corals. Comp. Biochem. Physiol. A Mol. Integr. Physiol. 159: 372-378.
Mydlarz, L., S. F. Holthouse, E. C. Peters, and C. D. Harvell. 2008. Cellular responses in sea fan corals: granular amoebocytes react to pathogen and climate stressors. PLoS One 3: el811.
Nauseef, W. M., and N. Borregard. 2014. Neutrophils at work. Nat. Immunol. 15: 602-611.
Olano, C. T. 1993. Cellular aspects of alloimmunity and other responses in the gorgonian Swiftia exserta. M. S. thesis, Florida International University, Miami, FL.
Olano, C. T., and C. H. Bigger. 2000. Phagocytic activities of the gorgonian coral Swiftia exserta. J. Invertebr. Pathol. 76: 176-184.
Pampanin, D. M., M. G. Martin, and L. Ballarin. 2002. Morphological and cytoenzymatic characterization of the haemocytes of the venus clam Chamelea gallina. Dis. Aquat. Organ. 49: 227-234.
Papadimitriou, J. M., and P. A. Wyche. 1976. Biochemical profile of glass-adherent cell populations containing multinucleated foreign-body giant cells. J. Pathol. 119: 239-254.
Paz, M. A., R. Fliickinger, A. Boak, H. M. Kagan, and P. M. Gallop. 1991. Specific detection of quinoproteins by redox-cycling staining. J. Biol. Chem. 266: 689-692.
Pearse, A. G. E. 1968. Histochemistry, Theoretical and Applied, Vol. 1, 2nd ed. Churchill Press, London.
Pearse, A. G. E. 1972. Histochemistry, Theoretical and Applied, Vol. 2, 2nd ed. Churchill Press, London.
Peddie, C. M., A. C. Riches, and V. J. Smith. 1995. Proliferation of undifferentiated blood cells from the solitary ascidian, Ciona intestinalis, in vitro. Dev. Comp. Immunol. 19: 377-387.
Pinzon, J. H., J. Beach-Letendre, E. Weil, and L. D. Mydlarz. 2014. Relationship between phylogeny and immunity suggests older Caribbean coral lineages are more resistant to disease. PLoS One 9: e 104787.
Pipe, R. K. 1990. Hydrolytic enzymes associated with granular haemocytes of the marine mussel Mytilus edulis. Histochem. J. 22: 595-603.
Rinkevich, B. 2012. Neglected biological features in cnidarian self-nonself recognition. Adv. Exp. Med. Biol. 738: 46-59.
Rogers, C. 2009. Coral bleaching and disease should not be underestimated as causes of Caribbean coral reef decline. Proc. R. Soc. B 276: 197-198.
Salter-Cid, L., and C. H. Bigger. 1991. Alloimmunity in the gorgonian coral Swiftia exserta. Biol. Bull. 181: 127-134.
Smith, V. J., and K. Soderhall. 1991. A comparison of phenol oxidase activity in marine invertebrates. Dev. Comp. Immunol. 15: 251-261.
Soderhall, K., and V. J. Smith. 1983. Separation of the haemocyte populations of Carcinus maenas and other marine decapods, and prophenoloxidase distribution. Dev. Comp. Immunol. 7: 229-239.
Sparks, A. K. 1972. Invertebrate Pathology. Academic Press, New York.
Stein, E. A., and E. L. Cooper. 1978. Cytochemical observations from the earthworm, Lumbricus terrestris. Histochem. J. 10: 657-678.
Sugumaran, M. 2001. Control mechanisms of the prophenoloxidase cascade. Adv. Exp. Med. Biol. 484: 289-298.
Wang, Z., A. Lu, A. Li, Q. Shao, B. T. Beerntsen, C. Liu, Y. Ma, Y. Huang, H. Zhu, and E. Ling. 2011. A systematic study on hemocyte identification and plasma prophenoloxidase from Culex pipiens quinquefasciatus at different developmental stages. Exp. Parasitol. 127: 135-141.
Yoshino, T. P., and T. C. Cheng. 1976. Fine-structural localization of acid phosphatase in granulocytes of the pelecypod Mercenaria mercenaria. Trans. Am. Microsc. Soc. 95: 215-220.
Young, J. A. 1974. The nature of tissue regeneration after wounding in the sea anemone Calliactis parasitica (Couch). J. Mar. Biol. Assoc. UK 54: 599-617.
LORENZO P. MENZEL (*) AND CHARLES H. BIGGER
Department of Biological Sciences, Florida International University, 11200 S.W. 8th Street, Miami, Florida 33199
Received 3 August 2014; accepted 12 June 2015.
(*) To whom correspondence should be addressed. E-mail: email@example.com
Table 1 Summary of adherence and enzyme-positive staining in the coenenchyme cells of Swiftia exserta Cell type Adherent Acid phosphatase Acetate esterase Ectoderm epithelia - - - Oblong granular cells + + + Granular amoebocytes + - + Morula-like cells ? ? + Mesogleal cells ? - - Cell type Quinone Peroxidase Phenoloxidase Ectoderm epithelia + + - Oblong granular cells + + + Granular amoebocytes + + - Morula-like cells + ? ? Mesogleal cells - + + ?, insufficient data for this cell type due to low cell numbers or morphological ambiguity; +, a positive staining reaction of this cell type for this enzyme reaction, in cryosections or in cytospin preparations; -, no staining of this cell type for this enzyme reaction, in cryosections or in cytospin preparations.
|Printer friendly Cite/link Email Feedback|
|Author:||Menzel, Lorenzo P.; Bigger, Charles H.|
|Publication:||The Biological Bulletin|
|Date:||Oct 1, 2015|
|Previous Article:||Flexibility of crab chemosensory sensilla enables flicking antennules to sniff.|
|Next Article:||A new species of Protodrilus (Annelida, Protodrilidae), covering bone surfaces bright red, in whale-fall ecosystems in the northwest Pacific.|