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Adherens junctions modulate diffusion between epithelial cells in Trichoplax adhaerens.

Abstract. Trichoplax adhaerens is the sole named member of Placozoa, an ancient metazoan phylum. This coin-shaped animal glides on ventral cilia to find and digest algae on the substrate. It has only six cell types, all but two of which are incorporated into the epithelium that encloses it. The upper epithelium is thin, composed of a pavement of relatively large polygonal disks, each bearing a cilium. The lower epithelium is thick and composed primarily of narrow ciliated cells that power locomotion. Interspersed among these cells are two different secretory cells: one containing large lipophilic granules that, when released, lyse algae under the animal; the other, less abundant, is replete with smaller secretory granules containing neuropeptides. All cells within both epithelia are joined by adherens junctions that are stabilized by apical actin networks. Cells are held in place during shape changes or under osmotic stress, but dissociate in low calcium. Neither tight, septate, nor gap junctions are evident, leaving only the adherens junction to control the permeability of the epithelium. Small (<4 kDa) fluorescent dextrans introduced into artificial seawater readily penetrate into the animal between the cells. Larger dextrans enter slowly, except in animals treated with reduced calcium, indicating that the adherens junctions form a circumferential belt around each cell that impedes diffusion into the animal. During feeding, the limited permeability of the adherens junctions helps to confine material released from lysed algae within the narrow space under the animal, where it is absorbed by endocytosis.


Placozoans are millimeter-sized, coin-shaped animals living in warm oceans where they locomote on surfaces to find and eat algae and bacteria (Grell and Ruthmann, 1991). They have no gut; instead, they feed by secreting digestive enzymes externally to lyse algae and bacteria trapped below their lower surfaces (Smith et al., 2015). This mode of feeding is unique among extant metazoans, but may have been more prevalent at an earlier era in metazoan evolution (Sperling and Vinther, 2010; Arendt et al., 2016). Recent analyses of the metazoan phylogenetic tree based on genomic sequences of representatives of different phyla place Placozoa as sister to the Eumeta-zoa, and Ctenophora and Porifera as the earliest diverging metazoan phyla (Ryan et al., 2013; Moroz et al, 2014). That ctenophores appear to have diverged earlier than placozoans and, possibly, also poriferans, came as a surprise because ctenophores appear to be much more complex (Ryan et al., 2013; Marlow and Arendt, 2014; Halanych, 2015; Jekely et al., 2015).

Trichoplax adhaerens Schulze, 1883 is the only named member of the phylum and has been the focus of most of the research on placozoans. Like all metazoans, T. adhaerens is enclosed by an epithelium (Grell and Benwitz, 1971; Grell and Ruthmann, 1991; Smith et al., 2014). Its dorsal (upper) epithelium is thin, composed of a pavement of relatively large polygonal disks whose cell bodies lie deeper in the interior (Smith et al., 2014). Around the margins of the animal, the epithelium transitions to a pseudostratified arrangement including many more densely packed cells. The most prevalent cell type is columnar and bears a single cilium and multiple microvilli; this cell type is present in the digestive tracts of many metazoans (Field and Frizzell, 1991). Dispersed among these cells are non-ciliated lipophil cells that contain large granules, including one very large granule closely apposed to the ventral surface. Arrayed in a ring around the circumference and sparsely scattered in the interior are typical gland cells that contain neuropeptides and express neurosecretory proteins. Sandwiched between the dorsal and ventral epithelia are fiber cells whose branching processes contact each other and all other cell types. Crystal cells, each containing a birefringent crystal, are distributed near the edge of the animal. Other than its well-defined top and bottom, the animal displays no axis of symmetry (Grell and Benwitz, 1974).

Trichoplax adhaerens locomotes by gliding, propelled by its cilia, which beat asynchronously and appear to contact the substrate intermittently (Smith et al, 2015). The path of movement is a random walk (Ueda et al, 1999). The animal often undergoes changes in shape from roughly circular to elongated. When it encounters a cluster of algae, it may pause to feed (Smith et al., 2015). First, the cilia cease beating and the animal becomes completely immobile. Then lipophil cells in the vicinity of the algae secrete their large ventral granule, followed rapidly by lysis of the algae underneath. The algal lysate disappears over the course of minutes, presumably due, at least in part, to uptake by the animal (Ruthmann et al., 1986). Cells in central regions of the animal begin to make rhythmic churning motions while the algal contents are endocytosed. Cells around the margin remain stationary. When most of the algal material has disappeared, the animal begins to move again. Neuropeptides secreted by neurosecretory cells may have a role in coordinating the activities of cells during feeding, as do neuropeptides secreted by neurons in more complex animals (Jekely, 2011, 2013).

Different types of junctions in metazoan epithelia are specialized to adhere cells together, to mediate communication between them, and to impede passage of solutes between cells (Farquhar and Palade, 1963; Miller et al, 2013). Epithelial cells in fresh water, but not marine sponges, can be joined by tight junctions, conferring high transepithelial resistence (Adams et al., 2010), while cells in cnidarians are joined by septate junctions that impede movements of proteins between cells (Tucker and Adams, 2014). Epithelial cells of T. adhaerens are interconnected by apical junctions that resemble the adherens junctions of other metazoans (Ruthmann et al., 1986; Smith et al., 2014). Dense periodic material suggestive of a second type of junction occupies the clefts between cells subjacent to the apical adhering junctions. The T. adhaerens genome contains a classical-type cadherin as well as the cadherin-binding partners, [beta]-catenin and pl20ctn (Srivastava et al, 2008; Hulpiau and Van Roy, 2010). Core constituents of septate junctions also are present (Ganot et al, 2015). No gap junctions are reported (Smith et al., 2014), nor are genes for pannexins present in the genome (Srivastava et al, 2008).

Our goal was to better characterize the apical junctions in the epithelium surrounding the animal, particularly to determine whether they might control passage of molecules between epithelial cells into its interior during feeding. We first confirmed that cells in the epithelium retain their topographic relationships while the animal changes shape, indicating that the cells are, indeed, held together by stable junctions. We probed the junctions that hold the cells together by examining the effects on the epithelium of osmotic stress (Brightman and Reese, 1969). We then examined the potential role of the junctions in regulating the passage of materials between epithelial cells into the animal by testing the permeability of the epithelium with fluorescent dextrans of varying sizes. Finally, we monitored the diffusion of fluorescent material released from lysed algae to test the possibility that it diffuses through the junctions during feeding, and to find out how effectively it is trapped in the space under the animal. The picture that emerges is of an epithelium held together and partially sealed by a single belt of zonulae adherens surrounding each epithelial cell. During feeding, the limited permeability of the adherens junctions helps to confine materials released from lysed algae to the space under the animal, where they are taken up into epithelial cells by endocytosis.

Materials and Methods


Trichoplax adhaerens of the Grell and Benwitz (1971) strain, a gift of Leo Buss of Yale University, were maintained in artificial seawater (ASW; Instant Ocean, Blacksburg, VA) with Rhodamonas salina algae (also called Pyrenomonas salina', Bigelow National Center for Culture of Marine Algae and Microbiota, East Boothbay, ME), as described previously (Jackson and Buss, 2009). Animals selected for experiments ranged in diameter from ~0.5 to 1.5 mm.

Light microscopy

To visualize actin filaments, lipid granules, and nuclei simultaneously, animals were concurrently fixed, permeabilized, and stained in 4% paraformaldehyde and 0.125% glutaraldehyde in 0.1 mol [l.sup.-1] sodium cacodylate buffer with 0.4 mol [l.sup.-1] NaCl, 0.1% Triton X, Alexa Fluor 488 phalloidin (Thermo Fisher Scientific, Waltham, MA), Nile red, and Hoechst dye (Life Technologies, Carlsbad, CA) for 15 min. Specimens were rinsed in ASW, mounted in VectaShield (Vector Laboratories, Burlingame, CA), and imaged on a LSM510 confocal microscope (Carl Zeiss Microscopy, LLC, Thornwood, NY) with a 405, 488, or 543-nm excitation and a 40X1.3 numerical aperture (NA) objective. For staining actin filaments, animals were fixed in 4% paraformaldehyde in ASW with 20 mmol [l.sup.-1] HEPES for 30 min, permeabilized with 0.1 % Triton X (5 min), stained with Alexa Fluor 488 phalloidin (30 min), and mounted in Prolong Gold (Life Technologies). Images were collected using a SP5 confocal microscope (Leica Microsystems, Wetzlar, Germany) with a 40X1.3 NA objective and 488-nm excitation for Alexa Fluor 488 and 561-nm excitation for autofluorescence of intracellular granules.

For imaging epithelial cell membranes in living T. adhaerens, animals were transferred to Lab-Tek II coverslip chambers (Thermo Fisher Scientific, Waltham, MA) containing filtered ASW with FM1-43 (green) and FM4-64 (red) membrane dyes (Life Technologies). Stained cell membranes were visualized with a 40 X 1.2 NA objective on a SP5 laser-scanning confocal microscope (Leica Microsystems) with 488 and 561-nm excitation and detections windows at 500-550 nm (green) and 570-650 nm (red). A rectangular region was irradiated with high-intensity, 488-nm excitation to photobleach the green fluorescence while enhancing the red fluorescence of cell membranes there.

The permeability of epithelial junctions was probed with small (<1 kDa) fluorescent dyes (Bodipy, Life Technologies; and CF 488, Biotium, Fremont, CA) and fluorescein-conjugated dextrans (4, 10, 40, and 70 kDa, Life Technologies; 147 kDa, Sigma-Aldrich, St. Louis, MO). The fluorescent dextrans were dialyzed against ASW before use. Trichoplax adhaerens animals were transferred to a Warner RC-40LP chamber with a #1.5 cover glass (Warner Instruments, Hamden, CT) that contained 0.5-ml filtered ASW. Animals were incubated in FM4-64 to label cell membranes before the fluorescent probe (~0.25 mg/ml) was added. Image stacks (2-[micro]m interval) were collected with a 40X1.2 NA C-Apochromat objective on a LSM510 confocal microscope with 488 and 543-nm illumination.

Merged transmitted light and fluorescence images of T. adhaerens feeding on Rhodamonas salina algae were captured on a LSM 510 confocal microscope with 543-nm illumination and a 5X0.25 or 10X0.45 NA objective. For higher-magnification views of the diffusion of phycobilin fluorescence released from lysed algae during feeding, T. adhaerens were transferred to Lab-Tek II coverslip chambers (Thermo Fisher Scientific) containing filtered ASW, Rhodamonas salina algae, and a fluorescent fatty acid (Cl-Bodipy 500/510-C12; Life Technologies) to label granules of lipophil cells. Image stacks (2-[micro]m interval; 720 ms per stack) were collected with a 40X 1.2 NA objective on a SP5 laser-scanning confocal microscope (Leica) with 488 and 561-nm excitation and 500-550 and 570-700-nm detection windows. Orthogonal views of the image stacks were created with Volocity 3D Image Analysis software (Perkin-Elmer, Waltham, MA).

Freeze-substitution and transmission electron microscopy

Trichoplax adhaerens were prepared by freeze-substitution, as previously described (Smith et al., 2014). Briefly, the animals were transferred to gold specimen carriers designed for high-pressure freezing (Leica, Bannockburn, IL) and high-pressure frozen in a Baltec 010 high-pressure freezing machine (TechnoTrade Inti., Manchester, NH). Specimens were gradually warmed over 3 days in a Leica AFS1 freeze-substitution machine in uranyl acetate, osmium tetroxide, and hafnium chloride dissolved in dry acetone. Epon resin was used for embedding; sections ~70-nm thick were stained with uranyl acetate and lead citrate, then photographed in a JEOL 200-CX electron microscope (JEOL USA, Inc., Waterford, VA) with a bottom-mounted AMT camera (AMT [Advanced Microscopy Techniques], Woburn, MA). Tomography of thin sections of animals prepared by freeze-substitution was performed in a FEI Tecnai TF-30 transmission electron microscope (Thermo Fisher Scientific) operated at 300 kV with a field emission electron gun. Images were collected in darkfield mode with 1.2-nm pixel resolution, as previously described (Chen et al., 2008).


Staining Trichoplax adhaerens with fluorescent phalloidin highlights actin networks at the apical junctions of epithelial cells, as evident by confocal microscopy in cross-sectional (Fig. 1A) and en face (facing) views (Fig. 1B). The irregular polygons of fluorescent actin networks surrounding epithelial cells are much smaller in the ventral epithelium than in the dorsal epithelium (Fig. 1B), reflecting the myriad narrow necks of epithelial cells reaching the ventral surface. The polygonal actin networks in the dorsal epithelium may vary in eccentricity in different regions of an animal or between animals, likely because they were stretched to different degrees and in different directions by movements of the animal prior to fixation (Fig. 1B).

Topographic stability of epithelial cells

Trichoplax adhaerens can stretch markedly while crawling or undergoing fission (Schulze, 1891) and, during feeding, typically make churning movements that stretch cells in different directions (Smith et al, 2015). To determine whether attachments between epithelial cells remain stable during stretching, patterns of contact between epithelial cells were examined in living animals by double-staining their epithelia with red and green membrane dyes (FM4-64 and FM1-43). When a rectangular region was exposed to intense, 488-nm illumination (a wavelength that excites both dyes), the red emission from within the region became slightly more intense while the green emission photo-bleached. The highlighted subset of cells could then be followed for up to 60 min as the animal moved. These epithelial cells maintained their spatial relationships and rectangular pattern even as the animals stretched them, showing that the cells are mechanically joined by stable attachments (Fig. 2).


Apical junctions by electron microscopy (EM)

Thin-section EM of freeze-substituted epithelia (Fig. 3) revealed apical junctions between epithelial cells in every plane of section. Confirming previous reports (Ruthmann et al, 1986; Smith et al. 2014), these junctions manifested features typical of adherens junctions in eumetazoans, so that here they will be referred to as adherens junctions. The gap separating the membrane of adjacent cells at the adherens junctions was ~20 nm. The intercellular spaces immediately adjacent to the junctions varied in width but typically narrowed to 8-10 nm, where the gaps were bridged by loosely periodic material, as previously reported (Ruthmann et al., 1986). Tufts of material were periodically distributed along membrane surfaces near adherens junctions, and these interdigitated where the adjacent membranes come close together, appearing to account for the periodic material in the clefts near adherens junctions. The numbers of tighter appositions containing periodic material varied between and within animals, and were more frequent on dorsal than ventral surfaces.


Adhering property of apical junctions

Adherence of junctions in the ventral epithelium was tested by exposing living animals to artificial seawater (ASW) containing 0.5 mol [l.sup.-1] sucrose for 2 min prior to freezing them. As expected for cells exposed to hypertonic conditions (Brightman and Reese, 1969), ventral epithelial cells shrank and their plasma membranes pulled away from each other, leaving the apical adherens junctions as the only inter-epithelial contacts (Fig. 3D). The dense periodic material near the adherens junctions came apart under the same conditions, making it unlikely that it represents adhesive intercellular junctions. Adhering junctions between dorsal epithelial cells also remained intact. However, many of the membrane appositions containing dense periodic material remained intact in the dorsal epithelium, unlike those in the ventral epithelium, which separated. As is also typical of adherens junctions (Takeichi, 1977; Hyafil et al, 1981), the junctions separated when exposed to calcium- and magnesium-free seawater (Fig. 3E, inset).


Barrier function of apical junctions

The extent to which the apical junctions serve as a barrier to diffusion of molecules through the intercellular spaces was measured by adding fluorescent dyes and dextrans of differing molecular weights to the ambient seawater, then visualizing their penetration into living animals with confocal microscopy (Figs. 4, 5). Animals were stained with the red membrane dye, FM4-64, to outline cell membranes before the dextran was added. Stacks of confocal images extending though the entire thickness of the animal were captured at different time points between 2 min and 24 h after the addition of the fluorescent dextran. The extent of penetration of the different molecular weight (<1-147 kDa) fluorescent dyes and dextran (Fig. 5, graph) was measured as the ratio of the intensity of fluorescent dextran inside to outside the animal in an optical section ~4 [micro]m inside the ventral surface (Fig. 4A, C), because signals from further in (Fig. 4B, D) were diminished due to light scattering and more subject to variability due to differences in the flatness of the animals. At 2 to 30 min, green fluorescent dyes and dextran <4 kDa already had diffused into the spaces between cells, outlining the densely packed epithelial cell bodies near the ventral surface (Fig. 4A) and filling the interior space surrounding fiber cells (Fig. 4C, arrow), consistent with a previous report that the epithelium in T. adhaerens is permeable to small molecules (Thiemann and Ruthmann, 1990). Because cells occupied spaces that the dextran does not penetrate, the ratio of fluorescence inside to outside never exceeded 0.4, even for low molecular weight (4 kDa and smaller) dyes and dextran (Fig. 5, graph). In contrast, little fluorescent dextran, 70 kDa or larger, was detected inside the interior spaces of animals after 30 min of exposure (Fig. 4B, D; Fig. 5). Even after 24-h incubation with 147-kDa fluorescent dextran, very little fluorescence was detected in the interior spaces (not shown). After adding fluorescent dextrans and FM4-64 to the ASW, vesicles stained with both dyes appeared inside ventral epithelial cells within 10 min (not shown), and increased numbers were present after 30 to 45 min (Fig. 2B; Fig. 4B).



Cells joined by adherens junctions separate from each other when calcium is removed from the medium (Takeichi, 1977; Flyafil et al., 1981), so the same permeation tests were run in calcium-free ASW. Trichop lax adhaerens that were incubated in calcium-free ASW for up to 90 min remained intact and continued to move. Nevertheless, electron microscopy showed that the adherens junctions separated as expected (Fig. 3). Dextrans of all sizes penetrated rapidly (Fig. 5), with little dependence on time of exposure and molecular weight. In contrast, dextrans did not more readily penetrate the epithelium after the treatment with hypertonic seawater (not shown), which separated most contacts between ventral epithelial cells except for the adherens junctions (Fig. 3E). A closer look at the structure of the adhering junction afforded by high-resolution (1.4-nm pixel size) tomograms collected by darkfield electron microscopy, showed thickets of minute filaments filling the intercellular spaces at the junctions (Fig. 3D, inset). The 10-nm gold used as fiduciary markers roughly matches the estimated hydration diameter of 40 to 70-kDa dextran molecules, and there appear to be pathways through the thickets of filaments crossing the junction that could admit 10-nm molecules, which might account for the slower passage of 40 kDa and larger dextran (Fig. 3D, inset). That penetration of the higher molecular weight dextran was impeded when the apical adherens junctions, but not other membrane appositions, were intact, suggests that the adherens junctions form circumferential belts, or zonulae, around epithelial cells and are the principal barrier to diffusion across the epithelium.

Limited transepithelial diffusion of materials released from lysed algae

Before the animals feed on algae, lipophil cells in the ventral epithelium secrete granules, including contents that lyse algae (Smith et al., 2015). The material released by the lysed algae includes phycobilins, which are intensely fluorescent. We followed the fate of the released phycobilins by confocal microscopy to find out whether they immediately diffuse into the interior of the animal or remain trapped below it. When the algae were uniformly distributed on the substrate, T. adhaerens crawling over them typically remained flat (Fig. 6A-C), trapping the algae in the narrow space between the ventral surface of the animal and the substrate (~5 [micro]m, as estimated from the focal plane difference measured with an objective providing ~1.0-[micro]m optical sections).

Animals crawling over a clump of algae typically maintained contact with the substrate at their edges (Fig. 6D), forming a pocket around the clump. Lysis of algae was evident as an abrupt swelling and an increase in fluorescent intensity of the algal cells. Released phycobilin fluorescence reached maximum intensity less than a minute after lysis, then declined over 2 to 6 min. Large clouds of fluorescence released by lysing of multiple algae persisted under animals longer than did smaller clouds from lysing of fewer algae. Very little phycobilin fluorescence penetrated under the surrounding flattened margins of the animal, although occasionally a small puff of fluorescence did escape from under the edge and diffused into the ambient seawater (not shown). The fluorescence also extended into shallow infoldings of the ventral epithelium that often developed following algae lysis (Fig. 6D). Examination of thin (~1-[micro]m) optical sections collected from the interior of the ventral epithelium in the vicinity of lysed algae showed that very little phycobilin fluorescence diffused between epithelial cells to reach the animal's interior during the time that fluorescence remained visible (Fig. 7). Although fluorescent endocytic vesicles were visible in ventral epithelial cells of animals stained with fluorescent membrane dyes or bathed in fluorescent dextrans (Fig. 2B; Fig. 4B), phycobilin fluorescence from lysed algae was not observed inside vesicles, possibly because the fluorescence was quenched following uptake into the interiors of endocytic vesicles.



Adherens junctions are ubiquitous in epithelia of metazoans, except for some poriferans (Dickinson et al., 2011; Leys and Riesgo, 2011; Miller et al., 2013). Cells making up the epithelium enclosing Trichoplax adhaerens are reported to be joined by adherens junctions (Grell and Ruthmann, 1991). We confirm that the apical epithelial junctions in T. adhaerens have in common with all adherens junctions a core structure consisting of a widened gap between the membranes of adjacent cells that is filled with filamentous material, and a dense layer applied to the cytoplasmic side of the membrane limiting the gap. This dense material is contacted by numerous filaments that arise from an actin network in the apical epithelium.


While T. adhaerens lacks the tight junctions or septate junctions that typically accompany adherens junctions in Eumetazoa (Ganot et al., 2015), close membrane appositions manifesting a vague periodic structure often lie nearby, in the clefts between the lateral surfaces of cells in the epithelium (Ruthmann et al., 1986). However, the outer membrane surfaces of epithelial cell membranes are everywhere invested with a fine fuzz, and where membranes come together, such as in the appositions near adherens junctions, zipping up of the fuzz from the adjacent membranes can account for the periodic structure seen in the clefts between cells. These periodic cleft structures disappear when the adjacent membranes are pulled apart by osmotic stress induced by hypertonic seawater. Under these conditions, only the adherens junctions remain intact, and the epithelium with the adherens junction intact continues to impede the entrance of high molecular weight dextrans. Thus, the ubiquitous presence of continuous belts of adhering junctions, or zonulae adherens, provides the principal impedance barrier to diffusion of higher molecular weight dextrans across the epithelium. Places near the adherens junctions where membrane coats interlock would not be expected to exert significant filtering, because their presence is quite variable from place to place in the epithelium.

The ventral epithelium performs functions analogous to those of the digestive endothelium in Ctenophora and Eumetazoa: secretion of digestive enzymes and nutrient uptake. Secretion by lipophils results in immediate lysis of entrapped algae that can be visualized directly by examining the clouds of fluorescent phycobilins released from the lysed algae beneath the animal (Smith et al., 2015). The space under the animal's ventral surface is narrow and is determined by the length of cilia that contact the substrate. When an animal crawls over a clump of algae or debris, its ventral surface conforms to the surface of the clump. The space beneath the animal enlarges somewhat when lipophil cells secrete their digestive granules (Smith et al., 2015), and forms a pocket of variable depth enclosing released fluorescent materials. Escape of fluorescent material from around the margins of the animal is limited. Imaging of the released fluorescent material shows that it has minimal access to the interior of the animal during the short time span that it is confined under that animal.

From the impeded entry of high molecular weight fluorescent dextrans and lysed algae fragments trapped under the animal during feeding, we concluded that the zonulae adherens in the epithelium acted as a barrier filter. Large proteins and algae fragments are likely to be taken up into ventral epithelial cells by endocytosis or phagocytosis, since these cells rapidly endocytose bath-applied ferritin (Ruthmann et al.. 1986) and fluorescent dextrans. However, the larger fragments of algae and other food particles seen in concrement vacuoles in fiber cells (Grell and Benwitz, 1971; Wenderoth, 1986) must have been transferred there from other cells with direct access to the surface of the animal. Thus, the zonulae adherens in T. adhaerens provide a continuous but leaky filter that separates the ambient seawater and the intercellular spaces inside the animal.

Gland cells with secretory machinery and peptides packaged in their granules are concentrated around the circumference of the ventral epithelium, but are also present, although more sparsely, in more central regions. Peptides secreted from gland cells may have a role in regulating feeding behavior incidental to external digestion, such as initiating pauses over algae or releasing digestive enzymes (Smith et al, 2015). The present results show that peptides, in contrast to larger proteins, likely have ready access to targeted cells inside and outside the animal.

The system of zonulae adherens that encircles the epithelial cells enclosing T. adhaerens holds the epithelium together while the animal contorts during crawling and while it chums during feeding. The adherens junctions also provide a filter that allows small molecules to pass between epithelial cells. The leaky barrier in epithelia of T. adhaerens contrasts with the much tighter barrier between epithelial cells in the intestines of higher animals with enclosed digestive tracts, where belts of tight junctions exclude solutes down to the size of ions (Lee, 2015). While lysed materials may linger in these enclosed digestive tracts, the fluorescent materials released from algae lysed by digestive secretions from a T. adhaerens disappear within approximately 5 min. None of the fluorescent material crosses the epithelium during this time. The zonulae adherens that partially seal the epithelium dictate that feeding in T. adhaerens must occur primarily by direct endocytosis at the surfaces of ventral epithelial cells.


Richard Leapman, Xiaobing Chen, and Maria Aronova from the Laboratory of Cellular Imaging and Macromolecular Biophysics, National Institutes of Biomedical Imaging and Bioengineering, National Institutes of Health, Bethesda, MD, provided the FEI Technai electron microscope and produced the darkfield tomograms. We thank Christine Winters and Rita Azzam for extensive technical assistance with electron microscopy, and Christine Winters and Tatiana Mayorova for careful review of the manuscript.

Literature Cited

Adams, E. D. M., G. G. Goss, and S. P. Leys. 2010. Freshwater sponges have functional, sealing epithelia with high transepithelial resistance and negative transepithelial potential. PLoS One 5: el5040.

Arendt, D., M. A. Tosches, and H. Marlow. 2016. From nerve net to nerve ring, nerve cord and brain--evolution of the nervous system. Nat. Rev. Neurosci. 17: 61-72.

Brightman, M. W., and T. S. Reese. 1969. Junctions between intimately apposed cell membranes in the vertebrate brain. J. Cell Biol. 40: 648-677.

Chen, X., C. A. Winters, and T. S. Reese. 2008. Life inside a thin section: tomography. J. Neurosci. 28: 9321-9327.

Dickinson, D. J., W. J. Nelson, and W. I. Weis. 2011. A polarized epithelium organized by beta- and alpha-catenin predates cadherin and metazoan origins. Science 331: 1336-1339.

Farquhar, M. G., and G. E. Palade. 1963. Junctional complexes in various epithelia. J. Cell Biol. 17: 375-412.

Field, M., and R. A. Frizzell, eds. 1991. Handbook of Physiology: Gastrointestinal System, Vol. 4. American Physiological Society, Bethesda, MD.

Ganot, P., D. Zoccola, E. Tambutte, C. R. Voolstra, M. Aranda, D. Allemand, and S. Tambutte. 2015. Structural molecular components of septate junctions in cnidarians point to the origin of epithelial junctions in Eukaryotes. Mol. Biol. Evol. 32: 44-62.

Grell, K. G., and G. Benwitz. 1971. Die Ultrastruktur von Trichoplax adhaerens F. E. Schulze. Cytobiologie 4: 216-240.

Grell, K. G., and G. Benwitz. 1974. Spezifische Verbindungsstrukuren der Faserzellen von Trichoplax adhaerens F.E. Schulze. Z Naturforsch. Sect. C Biosci. 29: 790-798.

Grell, K. G., and A. Ruthmann. 1991. Placozoa. Pp. 13-27 in Microscopic Anatomy of Invertebrates, Vol. 2, Placozoa, Porifera, Cnidaria, and Ctenophora, F. W. Harrison and E. A. Ruppert, eds. Wiley-Liss, New York.

Halanych, K. M. 2015. The ctenophore lineage is older than sponges? That cannot be right! Or can it? J. Exp. Biol. 218: 592-597.

Hulpiau, P., and F. van Roy. 2010. New insights into the evolution of metazoan cadherins. Mol. Biol. Evol. 28: 647-657.

Hyafil, F., C. Babinet, and F. Jacob. 1981. Cell-cell interactions in early embryogenesis: a molecular approach to the role of calcium. Cell 26: 447-454.

Jackson, A. M., and L. W. Buss. 2009. Shiny spheres of placozoans (Trichoplax) function in anti-predator defense. Invertebr. Biol. 128: 205-212.

Jekely, G. 2011. Origin and early evolution of neural circuits for the control of ciliary locomotion. Proc. Biol. Sci. 278: 914-922.

Jekely, G. 2013. Global view of the evolution and diversity of metazoan neuropeptide signaling. Proc. Natl. Acad. Sci. USA 110: 8702-8707.

Jekely, G., J. Paps, and C. Nielsen. 2015. The phylogenetic position of ctenophores and the origin(s) of nervous systems. EvoDevo 6: 1.

Lee, S. H. 2015. Intestinal permeability regulation by tight junction: implication on inflammatory bowel diseases. Intest. Res. 13: 11-18.

Leys, S. P., and A. Riesgo. 2011. Epithelia, an evolutionary novelty of metazoans. J. Exp. Zool. B Mol. Dev. Evol. 318: 438-447.

Marlow, H., and D. Arendt. 2014. Evolution: ctenophore genomes and the origin of neurons. Curr. Biol. 24: R757-R761.

Miller, P. W., 1). N. Clarke, W. I. Weis, C. J. Lowe, and W. J. Nelson. 2013. The evolutionary origin of epithelial cell-cell adhesion mechanisms. Curr. Top. Membr. 72: 267-311.

Moroz, L. L., K. M. Kocot, M. R. Citarella, S. Dosung, T. P. Norekian, I. S. Povolotskaya, A. P. Grigorenko, C. Dailey, E. Berezikov, K. M. Buckley et al. 2014. The ctenophore genome and the evolutionary origins of neural systems. Nature 510: 109-114.

Ruthmann, A., G. Behrendt, and R. Wahl. 1986. The ventral epithelium of Trichoplax adhaerens (Placozoa): cytoskeletal structures, cell contacts and endocytosis. Zoomorphology 106: 115-122.

Ryan, J. F., K. Pang, C. E. Schnitzler, A.-D. Nguyen, R. T. Moreland, D. K. Simmons, B. J. Koch, W. R. Francis, P. Havlak, NISC Comparative Sequencing Program et al. 2013. The genome of the ctenophore Mnemiopsis leidyi and its implications for cell type evolution. Science 342: 1242592.

Schulze, F. E. 1883. Trichoplax adhaerens, nov. gen., nov. spec. Zool. Anz. 6: 92-97.

Schulze, F. E. 1891. Uber Trichoplax adhaerens. Abh. K. Preuss. Akad. Wiss. Berl. 1-23.

Smith, C. L., F. Varoqueaux, M. Kittelmann, R. N. Azzam, B. Cooper, C. A. Winters, M. Eitel, D. Fasshauer, and T. S. Reese. 2014. Novel cell types, neurosecretory cells, and body plan of the early-diverging metazoan Trichoplax adhaerens. Curr. Biol. 24: 1565-1572.

Smith, C. L, N. Pivovarova, and T. S. Reese. 2015. Coordinated feeding behavior in Trichoplax, an animal without synapses. PLoS One 10: e0136098.

Sperling, E. A., and J. Vinther. 2010. A placozoan affinity for Dickinsonia and the evolution of late proterozoic metazoan feeding modes. Evol. Dev. 12: 201-209.

Srivastava, M., E. Begovic, J. Chapman, N. H. Putnam, U. Hellsten, T. Kawashima, A. Kuo, T. Mitros, A. Salamov, M. L. Carpenter et al. 2008. The Trichoplax genome and the nature of placozoans. Nature 454: 955-960.

Takeichi, M. 1977. Functional correlation between cell adhesive properties and some cell surface proteins. J. Cell Biol. 75: 464-474.

Thiemann, M., and A. Ruthmann. 1990. Spherical forms of Trichoplax adhaerens (Placozoa). Zoomorphology 110: 37-45.

Tucker, R. P., and J. C. Adams. 2014. Adhesion networks of Cnidarians: a postgenomic view. Int. Rev. Cell Mol. Biol. 308: 323-377.

Ueda, T., S. Koya, and Y. K. Maruyama. 1999. Dynamic patterns in the locomotion and feeding behaviors by the placozoan Trichoplax adhaerens. BioSystems 54: 65-70.

Wenderoth, H. 1986. Transepithelial cytophagy by Trichoplax adhaerens F. E. Schulze (Placozoa) feeding on yeast. Z. Naturforsch. Sect. C Biosci. 41: 343-347.


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Received 3 June 2016; accepted 12 September 2016.

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Author:Smith, Carolyn L.; Reese, Thomas S.
Publication:The Biological Bulletin
Article Type:Report
Date:Dec 1, 2016
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