Catecholic Compounds in Ctenophore Colloblast and Nerve Net Proteins Suggest a Structural Role for DOPA-Like Molecules in an Early-Diverging Animal Lineage.
Recent phylogenomic studies have proposed that ctenophores, or "comb jellies," a group of gelatinous, carnivorous, and largely planktonic marine invertebrates, are the sister group to all other animals (Dunn et al., 2008; Hejnol et al, 2009; Ryan et al, 2013; Moroz et al, 2014; Dunn and Ryan, 2015; Whelan et al, 2017). Though this "Ctenophora-sister" hypothesis is currently disputed (Pisani et al, 2015; Feuda et al, 2017; Simion et al, 2017), the resulting phylogenomic debate has sparked a resurgence of interest in the physiology and development of ctenophores (Dunn et al, 2015; Norekian and Moroz, 2016; Presnell et al, 2016) as researchers attempt to frame genomic insights from non-bilaterian animal lineages in a broader organismal biological context.
For example, the nature of a phylum-defining feature of ctenophores, the "colloblast," remains mysterious. Colloblasts are an adhesive cell type that covers the exterior surface of ctenophore tentacles and that has no obvious homolog in other animal groups. Colloblasts are used to ensnare prey in a manner akin to flypaper, and they are characterized by clusters of adhesive-containing granules. These granules rupture and release their contents on contact with prey. Little is known about the adhesive, except that it is likely proteinaceous (Franc, 1978), and it is suspected to be associated with a collagen-like, proline-rich molecule (Franc, 1985).
One known class of protein-based marine adhesives is found in both the byssal threads used to anchor mussels to their substrates and in the cement used for tube construction in some tube-dwelling polychaetes. Adhesion in these systems is mediated by the amino acid L-3,4-dihydroxyphenylalanine (L-DOPA). The 3,4-dihydroxyphenyl (catechol) substructure in L-DOPA is presumed to be produced by enzymatic hydroxylation of tyrosine (Waite, 1983; Zhao et al., 2005) via a tyrosinase (Guerette et al., 2013). When incorporated into polypeptides, L-DOPA possesses prodigious and tunable adhesive properties because of the catechol's capacity to coordinate with a wide variety of cations and biomolecules (Taylor et al., 1996;
Harrington et al., 2010; Hwang et al., 2010; Yu et al., 2011; Martinez Rodriguez et al., 2015). Further, collagen and collagen-like molecules in byssal tissue coordinate with L-DOPA-containing adhesive proteins and confer tensile strength to the byssal threads (Qin and Waite, 1995; Coyne et al., 1997; Martinez Rodriguez et al., 2015).
We hypothesized that ctenophore colloblasts contained a catechol-based adhesive, chemically similar to that found in mussel byssus, because both systems need to adhere to a variety of surfaces in a marine environment. We present data suggesting that catecholic compounds are incorporated into proteins in Pleurobrachia bachei colloblasts. Additionally, we find evidence of catechol-containing proteins associated with the nerve net beneath the locomotory comb rows of P. bachei. L-DOPA does occupy a key node in the biosynthetic pathway of the neurotransmitters dopamine, epinephrine, and norepinephrine, but ctenophores are reported not to use any of these molecules for neurotransmission (Moroz et al., 2014; Moroz and Kohn, 2016). However, our findings are in line with recent immunohistochemical observations of L-DOPA in nonneuronal cells of ctenophore polar fields (Jager et al., 2013), suggesting that the notion of a neural role for catecholic molecules in this phylum is worthy of further consideration.
Materials and Methods
Pleurobrachia bachei A. Agassiz, 1860 were collected by dipnetting on the floating docks at the University of Washington's Friday Harbor Labs in Friday Harbor, Washington (48.545234, -123.012020). All tentacle tissue was dissected away from the bodies (hereafter, "tentacle-free bodies") and was stored separately. All P. bachei bodies used in this study were free of any visible tentacle or tentacle bulb tissue. Shortly after collection, tissue samples were stored at -80 [degrees]C until use.
Arnow's reagent staining of whole ctenophores
Whole adult P. bachei (N = 5) were fixed in excess 4% paraformaldehyde (PFA) in 1 x phosphate-buffered saline (PBS) at 4 [degrees]C overnight, then washed in excess 1x PBS for 4 hours at 4 [degrees]C, then moved into successive baths of Arnow's reagent components A, B, then C (reagent A: 0.5 N HCl; reagent B: 1.45 mol [L.sup.-1] sodium nitrite and 0.41 mol [L.sup.-1] sodium molyb-date; reagent C: 1 N NaOH). Samples spent 10 minutes in each solution at 20 [degrees]C. Representative samples were dissected and photographed with identical camera settings and lighting before and after treatment.
Protein extraction, trichloracetic acid precipitation, and biochemical analysis of protein pellets
Tentacles dissected from 10 individual P. bachei were centrifuged at 4 [degrees]C for 2 minutes at 8000 x g, and any excess seawater leftover from the dissection was pipetted away. Then, 500 [micro]L of 1x PBS was added, and the tissue was sonicated on ice, using a Fisher Scientific FB505 sonicator (Hampton, NH) for 2 minutes at 30% amplitude, with a 10-second duty cycle using a micro-sonication tip. The resulting lysate was centrifuged at 4 [degrees]C for 10 minutes at 8000 x g. The supernatant was removed and reserved at -80 [degrees]C, while the pellet was rinsed in excess molecular biology-grade water. The rinse water was then removed and discarded, and this pellet was immersed in a second sonication buffer of 500 [micro]L of 0.1 mol [L.sup.-1] acetic acid/0.1% (w/v) cetrimonium bromide (CTAB) detergent, hereafter referred to as "AcOH/CTAB." The pellet was sonicated, and the lysate was centrifuged a second time, as above. The supernatant was removed and stored at -80 [degrees]C. Lysates from the 10 tentacle-free bodies were prepared in a similar manner. Because of their high water content, whole bodies were sonicated as above, but without any additional buffer. This lysate was centrifuged as above, and this supernatant, referred to as "no buffer," was removed and reserved. The pellet from this step was washed and immersed in 500 [micro]L AcOH/CTAB buffer, re-sonicated, and centrifuged as above. The AcOH/CTAB supernatant was drawn off and stored at -80 [degrees]C. Four additional standard solutions were prepared: 2 5 mg [mL.sup.-1] solutions of albumin from chicken egg white (62%-88%; Sigma-Aldrich, St. Louis, MO), 1 in 1x PBS and another in AcOH/CTAB, and 2 1 mg [mL.sup.-1] L-DOPA solutions (Tocris Bioscience, Bristol, United Kingdom), also in 1x PBS and AcOH/CTAB.
To separate whole protein from the rest of these lysates, we used tricholoracetic acid (TCA) precipitation. Ice-cold TCA (250 [micro]L, 20% w/v) was added to a like volume of lysate or standard solution; the solution was mixed by pipetting and was incubated at 4 [degrees]C for 10 minutes. These mixtures were centrifuged at 4 [degrees]C for 10 minutes at 20,000 x g. The supernatants were removed and reserved, while precipitated pellets, if present, were rinsed in excess molecular biology-grade water. Pellets were cut in half, and one half was immersed in 4% PFA overnight and was spectrally imaged by confocal microscope, as was done for whole tissue mounts, as described below. The second half of each pellet was placed on a glass slide and treated with Arnow's reagent (50 [micro]L of reagents A, B, and C applied in succession, each allowed to sit for about 1 minute before the application of the next). These Arnow-treated pellets were then photographed with standardized lighting conditions and camera settings. All TCA precipitation supernatants were likewise treated with Arnow's reagent (a 1:1:1:2 ratio of sample : reagent A : reagent B : reagent C), in order to completely neutralize the TCA. The ultraviolet-visible spectra of these Arnow-treated solutions were then measured on a NanoDrop 2000 spectrophotometer (Thermo Fisher, Waltham, MA). Droplets of Arnow-treated solutions were also photographed on slides in the same manner.
When exposed to formaldehyde, dihydroxyphenylalanine (DOPA) and related molecules form green-emitting fluorophores with known spectral characteristics (Bjorklund and Falck, 1973). To assay formaldehyde-induced fluorescence (FIF) in this system, adult P. bachei (N = 3) were fixed overnight in excess 4% PFA buffered with 1x PBS. Ctenes and tentacles were then dissected and mounted as described above. Mounted samples were imaged on a Zeiss LSM 710 laser scanning confocal microscope (Oberkochen, Germany), with a 32-channel spectral detector. Samples were excited with a laser tuned separately to 405, 458, and 488 nm (although only the 405-nm excitation resulted in a detectable emission, see Results). Emission spectra resulting from the 405-nm excitation were measured from 425-700 nm with a wavelength resolution of 3.2 nm.
Images collected in this fashion were processed using the Zeiss Zen 2.3 Lite software package. This software displays the data collected by the spectrometer detector as colorized composites, with colors of individual pixels corresponding to the perceptual color of light emitted from that region of the image. Spectral profiles for various regions of interest have been computed from the average pixel intensity within each 3.2-nm-wide spectral bin. To statistically assess the degree of similarity between spectra, we computed Pearson's correlation coefficient for each pair of spectra under consideration within the 420-650-nm wavelength range diagnostic for FIF in MATLAB (MathWorks, Natick, MA).
Immunohistochemistry and confocal microscopy
Whole adult P. bachei were fixed in excess 4% PFA in 1x PBS at 4 [degrees]C for 1.5 hours. Fixed individuals were bisected along the pharyngeal plane and moved to a solution of 1x PBS to wash out excess fixative. Fixed and washed samples were blocked overnight at 4 [degrees]C in 5% normal donkey serum in lx PBS. Blocked samples were incubated for 36 hours at 4 [degrees]C with rabbit polyclonal anti-L-DOPA antibody (ab6426, Abeam, Cambridge, MA; Research Resource Identifier [RRID]: AB_305457) 1 : 4500 in blocking solution. After this primary incubation, samples were washed 6 times with excess volumes of blocking solution, then incubated overnight in the dark at 4 [degrees]C with anti-rabbit immunoglobulin G DyLight 488-conjugated secondary antibodies (H&L, Rockland Immunochemicals, Limerick, PA), also diluted 1 : 4500 in blocking solution. After secondary antibody incubation, samples were washed 6 times with 1x PBS at 4 [degrees]C. After secondary staining, these samples were dissected to remove the tentacles and tentacle base whole, as well as sections of comb rows.
To test whether the primary anti-DOPA antibody specifically binds L-DOPA, we used a strategy of pre-incubating pure L-DOPA antigen with the primary antibody prior to introducing this mixture to the fixed tissue, as in the standard protocol described above. If the L-DOPA antigen were specifically bound to the antibody after this pretreatment, it would mask the antibody's paratope, preventing it from generating false-positive binding events to epitopes in the fixed tissue; and we would expect a negative staining result in subsequent fluorescence imaging. For these pre-incubated antigen controls, diluted primary antibodies were incubated with 1000x or 10x molar excess pure L-DOPA overnight, then used to stain samples as above. We also controlled for nonspecific binding of the secondary antibody by staining samples with secondary antibody alone. In this case, only the antirabbit immunoglobulin G DyLight 488-conjugated secondary antibody solution diluted 1 : 4500 in blocking solution was applied to the tissues; and preparations were incubated overnight, in the dark at 4 [degrees]C, then washed 6 times with 1x PBS at 4 [degrees]C.
For all antibody experiments, after incubation with antibodies as described above, dissected pieces of tissue were mounted on glass slides with Vectamount hard-set mounting medium (Vector Labs, Burlingame, CA) and imaged on a Nikon Eclipse E800 upright microscope (Minato, Tokyo) with a BioRad Radiance 2000 laser scanning system (Hercules, CA). All tissues and treatments described here were imaged using the same laser and camera settings. For each set of conditions, N = 3 individuals were subjected to treatment.
Arnow's reagent staining
Arnow's reagent is a three-part colorimetric assay that is specific for the catecholic structure of ortho-diphenols, or o-diphenols (Arnow, 1937; Waite and Tanzer, 1981). When treated with Arnow's reagent, mono- and unsubstituted o-diphenols form a red-orange to red chromophore (Arnow, 1937; Waite and Tanzer, 1981). This assay was previously used as a histological stain in studies of DOPA-containing adhesive proteins in polychaetes (Wang and Stewart, 2012) and DOPA-hardened hydroid perisarc (Hwang et al., 2013). Treatment of whole, fixed Pleurobrachia with Arnow's reagent (N = 5 individuals) resulted in red-orange staining of the animals' tentacles (Fig. 1A, B, insets at top). The treated comb rows had a slight yellow tint but were otherwise not obviously affected by treatment with Arnow's reagent (Fig. 1B).
Of various buffer conditions used for extracting bulk protein from tentacle tissue, only AcOH/CTAB fully disrupted the tissue and extracted enough protein to form a visible pellet. This pellet, when treated with Arnow's reagent, stained a dark red color (Fig. 1B). Extraction of protein from tentacle-free Pleurobrachia bachei bodies was more straightforward, and sonication without additional buffer was sufficient to extract a visible protein pellet (Fig. 1A). Resonication of the material leftover from this initial extraction of the body tissue in AcOH/CTAB buffer resulted in further protein extraction (Fig. 1A). After TCA precipitation, protein samples from both the tentacles and the rest of the body stained a pale orange color when treated with Arnow's reagent (Fig. 1B). In all buffer conditions and all tissue types, the corresponding supernatants that resulted from TCA precipitation of total protein lacked the absorbance peaks at or around 400 and 500 nm that are characteristic of positive staining for o-diphenols by Arnow's reagent (Fig. 1B).
As a negative control, we tested protein pellets generated from solutions of low-purity chicken egg albumin with Arnow's reagent (Fig. 1 A). These samples did not stain orange, nor did they produce supernatants with absorbance peaks at 400 or 500 nm (Fig. 1B). As a positive control we tested solutions of pure L-DOPA. These solutions formed no pellet when TCA was added, but the resulting solution turned bright red and exhibited the absorbance peaks at 400 and 500 nm characteristic of o-diphenols in this assay (Fig. 1B).
Both ctene and tentacle samples that were fixed at least overnight in 4% PFA/1x PBS showed spatial patterns of autofluorescence when excited with 405-nm light. In contrast, no autofluorescence was observed in unfixed P. bachei ctenes or tentacles or in samples that had been fixed for 1.5 hours in 4% PFA/1x PBS. Laser excitations at 458 and 488 nm were also assayed, but only the 405-nm excitation produced any detectable autofluorescence. The emission spectra from both tissue regions were broad, with a full-width half-maximum value in both tissues of about 125 nm (Fig. 2B, D). The presence of a few local maxima within this broad spectrum suggests that the observed emission spectrum may be the sum of a few independent, spectrally separated emitters (Fig. 2B, D).
The FIF images of tentacles show two distinct features (Fig. 2A). We observed punctate, green-emitting, spherical granules consistent in appearance with adhesive granules in the colloblasts. In contrast, the non-colloblast regions of the tentacles emitted an unstructured, diffuse blue signal. We interpret this blue signal to be the general ctenophore tissue background fluorescence with PFA fixation. The fluorescence emission of the colloblasts was both more intense and red-shifted from that of the diffuse blue background: the colloblast granules emitted at 503, 522, and 535 nm, whereas the tissue background had local maxima at 483 and 499 nm (Fig. 2B).
In comb row tissue, the cells in the epithelium underlying the ctenes contained green-emitting granules ranging in size from 1 to 8 [micro]m in diameter, while a blue-emitting mesh similar in structure to that observed in our immunohistochemistry experiments filled most of the space between them (Fig. 2C). The ctenes themselves were also visible as blue structures similar in emission spectrum to the mesh.
Emission spectra of the mesh and ctenes exhibited maxima at 480 and 496 nm (Fig. 2D) and were, overall, similar to the tissue background of the tentacles (Fig. 2B, D). In contrast, spectra from the mesh-associated granules showed a fluorescence emission maximum at 502 nm and minor peaks at 515 and 535 nm (Fig. 2D), also similar to the emission from granules on the colloblasts' exterior (Fig. 2B, D).
We also conducted FIF assays of the protein pellets resulting from TCA precipitation (Fig. 2B, D). These protein samples exhibited emission spectra similar to those of the whole-mounted tissue samples described above. Pellets from both tentacle and body tissue exhibited emission spectra with peaks at 515 and 535 nm. The pellet from tentacle tissue also had a distinct emission peak at 675 nm, possibly due to the co-precipitation of porphyrin-containing proteins. In contrast, egg albumin pellets prepared and imaged in the same fashion showed only diffuse structure, with an emission near 515 nm, similar to the unstructured background of tentacles and ctene tissue.
To assess the degree of similarity between fluorescence emission spectra that we measured, we computed pairwise Pearson's correlation coefficients, r, for these spectra (Tables A1, A2). We restricted the input data to wavelengths less than 650 nm across all comparisons to restrict the analysis to wavelengths that are diagnostic for FIF. All pairs of spectra compared in this fashion show correlation (i.e., each pair differs from the null hypothesis of no correlation, or r = 0 with P < [10.sup.-3]). However, within each tissue, the highest values of the correlation coefficient were for pairs of tissue green-emitting granules and CTAB-extracted ctenophore proteins, and pairs of tissue blue-background, CTAB-extracted albumin samples (Tables A1, A2, bold).
In P. bachei tentacles, using a standard staining protocol with an anti-L-DOPA primary antibody followed by an appropriate fluorescent secondary antibody yielded a punctate pattern of bound antibody outlining larger spherical structures approximately 350 nm in diameter (Fig. 3A, B). The positively staining spheres are closely packed and are in turn organized into a hemispherical pattern with ~250-nm gaps between the edges of individual 350-nm spheres. This staining pattern is consistent with antibody bound to colloblast adhesive granules, as was originally identified via electron microscopy by Franc (1978). The tentillae are uniformly covered in these positively staining granules, while on the main branches of the tentacles, they are present in spiral bands running along the long axis.
The same protocol applied to comb rows yielded an extensive net-like pattern of positive-staining granules filling the area between individual comb plates and extending outward beyond the edges of the comb plates (Fig. 3F, G). The average mesh size of the pattern directly between the comb plates is 5-6 [micro]m, and it increases to 7-8 [micro]m at the distal edges of the imaged sections. Individual brightly staining granules within the mesh were about 1.4 [micro]m in diameter, and they largely appeared within the mesh outline but occasionally formed large clumps ~10 [micro]m in diameter that disrupted the mesh pattern. This mesh and its associated granules lay in a plane parallel to the body wall of the animal and formed a layer approximately 2-3 [micro]m thick. We also observed a pattern of pronounced, striated positive staining at the base of each comb plate (Fig. 3F). These striations radiated out from the comb plates and did not appear to be directly connected to the nearby mesh. The striations emanating from the comb rows occurred with a spacing of 7-8 lines every 10 [micro]m.
As other investigators have noted, fixation causes ctenophore tentacles to become somewhat "sticky" to a variety of reagents (von Byern et al., 2010; Dayraud et al., 2012). Accordingly, staining P. bachei tentacles with only fluorescently labeled secondary antibodies as a nonspecific binding control yields a staining pattern that is spatially similar to, but less efficient than, the standard protocol described above, producing a stain that is ~30% dimmer, as estimated by average grayscale value (Fig. 3C). In comb row preparations, this nonspecific staining control protocol yielded no staining of the mesh, granules, or ctene striations observed in the traditional dual-antibody stain described above (Fig. 3H).
Because these treatments yielded staining patterns that were similar spatially but different in intensity, we included an additional antigen specificity control. If the primary antibody can specifically bind its antigen in this pre-incubation, the antibody's paratope will be masked, preventing it from subsequently binding to epitopes in the tissue. Any staining observed in this treatment must be due to either nonspecific binding of the primary antibody to the tissue or to interactions between the tissue and the pure antigen itself. In P. bachei tentacles, this treatment yielded a markedly different staining pattern from secondary antibodies alone. The punctate staining pattern of the standard protocol was abolished, leaving a fainter, unstructured stain throughout the tissue (Fig. 3D). In P. bachei comb rows, the same control experiment revealed neither the mesh structure, nor the granules between and to the sides of the comb plates, nor the bright striations emanating from the comb plates. In all comb row control experiments, only a faint background signal from the comb plates remained (Fig. 3I). This suggests that the primary anti-L-DOPA antibody used does bind specifically to pure L-DOPA, but that in the tentacles, L-DOPA from the pre-incubation step interacted with the tissue in a nonspecific fashion to produce the resulting faint, unstructured pattern.
We found evidence of non-canonical, catecholic amino acids in colloblast granules of the ctenophore Pleurobrachia bachei. Surprisingly, we also found similar catecholic granules associated with the comb row subepithelial nerve net. Our finding that these analytes are detectable in TCA-precipitated protein pellets from P. bachei tissue lysates, but not in the associated supernatants, suggests that these catecholic amino acids are not free in solution but rather are associated with and potentially incorporated into proteins.
Previous investigations of catecholic metabolites in ctenophores utilized capillary electrophoresis in combination with mass spectrometry or fluorescence detection to probe tissue extracts against a specific set of standard molecules (Moroz et al., 2014). These studies concluded that there was no L-DOPA (or dopamine) present in ctenophores, at least as a small signaling molecule in solution (Moroz et al., 2014; Moroz and Kohn, 2015). The results of the present study do not contradict this prior work. Capillary electrophoresis can accurately detect very small quantities of metabolites in a biological sample relative to specific standards, but it may fail to detect or distinguish close chemical variants of these standards without coupling to a second analytical technique such as mass spectrometry (Cruz et al., 1996; Lopez-Montes et al., 2013). Similarly, the multiple local maxima in our FIF data suggest that the catecholic residues in P. bachei tentacles and comb rows may be closely structurally related to L-DOPA, rather than L-DOPA per se, and thus were unlikely to have been part of a set of chemical standards. Furthermore, our data suggest that catecholic amino acids are present in P. bachei in a polymerized, proteinaceous form, while Moroz et al. (2014) specifically assayed for small, solution-state catecholamines and their precursors.
Neither L-DOPA nor dopamine affects ciliary beat in P. bachei (Moroz et al., 2014). Moreover, P. bachei, like all other ctenophores studied, does not encode the ionotropic receptors required for catecholamine signaling (Moroz et al., 2014; Moroz, 2015). However, Moroz and colleagues also predicted dozens of novel putative peptidic prohormones in P. bachei and other ctenophore species; and they suggested that these peptides could be involved in signaling through the numerous epithelial sodium channels or orphan G protein-coupled receptors also identified by their studies (Moroz et al., 2014; Moroz, 2015; Moroz and Kohn, 2015). The addition of a variety of post-translational modifications to prohormones during intracellular processing is common, but the physiological significance of most of these additions is poorly understood (Perone and Castro, 1997). Our data are compatible with a scenario in which a population of the putative prohormones or proneuropeptides identified by Moroz and colleagues contains post-translationally modified, catecholic amino acids that would likely have gone undetected by assays designed to detect catechols as solution-state small molecules. Further molecular biological work to isolate these peptides and study their activity in vivo is necessary to assess this question.
Our observation that both the tentacles and the ctenes of P. bachei only fluoresce after an overnight treatment with formaldehyde demonstrates that this fluorescence is the result of the slow reaction of formaldehyde with molecules in the tissue. FIF has previously been used to detect and to distinguish dopaminergic and serotonergic neurons. In general, the FIF emission peak moves to a longer wavelength with additional substitution on an aromatic ring: 5-hydroxytryptamine-and 5-hydroxytryptophan yield a green-yellow fluorescence; L-DOPA has an emission maximum around 480 nm but an overall greenish appearance due to the long tail of the spectrum; while 5-hydroxydopamine (5-OH-DA) has an emission peak of about 500 nm that shifts to 530-540 nm at high concentrations, as seen in some vertebrate neurons (Tranzer and Thoenen, 1967; Baumgarten et al., 1972).
Though the granules in the nerve net and in the colloblasts are presumably functionally distinct, their formaldehyde-induced fluorophores have similar emission spectra, with a major peak near 510 nm and multiple minor peaks at longer emission wavelengths. This is consistent with the granules containing a mixture of multiply substituted phenolic rings, possibly at high concentrations. Furthermore, the observed positive Arnow staining of TCA-precipitated protein pellets from both tentacle and tentacle-free body tissue lysates suggests that these catecholic compounds are present in proteins in these tissues.
Our evidence that catecholic amino acids are localized to the adhesive granules of Pleurobrachia colloblasts suggests a role for these compounds in colloblast adhesion and prey capture. This is potentially similar to mussel and polychaete adhesive systems. Mussel byssus (Waite and Tanzer, 1980; Waite, 1983; Benedict and Waite, 1986; Deacon et al., 1998;Zhaoand Waite, 2006; Guerette et al., 2013) and tube-dwelling polychaete cement (Zhao et al., 2005; Becker et al., 2012; Wang and Stewart, 2012) are both secreted marine biological adhesives whose material properties are derived from catecholic amino acids in polypeptides. However, whether this chemical similarity constitutes a synapomorphy between any of these groups is difficult to assess without more detailed biochemical and bioinformatic information about catecholic proteins and their biosynthesis in these groups.
In mussels, the inclusion of DOPA in byssal thread adhesive proteins is thought to be mediated by a tyrosinase (Guerette et al., 2013). Tyrosinases are a class of enzymes distinct from tyrosine hydroxylases in mechanism, substrate specificity, and expression patterns. Pleurobrachia bachei and other ctenophores reportedly do not express tyrosine hydroxylases (Moroz et al., 2014; Moroz and Kohn, 2015, 2016), which is notable because in species that use catecholamines such as dopamine as neurotransmitters, tyrosine hydroxylase catalyzes the initial step of these molecules' synthesis. However, P. bachei does express a transcript annotated as a tyrosinase (Moroz et al., 2014, 2018). Tyrosinases (sometimes referred to as polyphenol oxidases, monophenol monoxygenases, catechol oxidases, cre-solases, or catecholases) are well known for their role in producing melanin in vertebrates. They are, however, found in all domains of life, and, as their numerous monikers suggest, they mediate reactions with various substrates to diverse functional ends (Sanchez-Ferrer et al., 1995; Claus and Decker, 2006; Mayer, 2006; Decker et al., 2007; Zaidi et al., 2014). Though the most well-known tyrosinase reactions are the conversion of tyrosine to L-DOPA and the oxidation of L-DOPA to L-dopaquinone (Raper, 1926; Sanchez-Ferrer et al, 1995; Ito and IFPCS, 2003), other reactions, such as the hydroxylation of L-DOPA residues in adhesive mussel foot protein peptides to 5-hydroxydo[rho]a (also known as "L-TOPA"), have also been observed (Burzio and Waite, 2002). Additional reactions and substrates have been observed in plant and fungal systems (Mayer, 2006). Because of the flexibility of phenol chemistry and the variable binding pocket geometry of tyrosinases, sequence data alone are insufficient to predict the activity of a given tyrosinase domain-containing protein. Thus, the presence of a tyrosinase domain in the P. bachei genome is congruent with the multiply substituted catechols that we observe in the animal; but in the absence of experimental information about its enzymatic activity, it is not possible to say with certainty whether this enzyme is producing these residues.
We were surprised to see evidence of catecholic amino acids in P. bachei neurons; however, this finding has precedent in other invertebrates. Several species of anthozoans have been found to contain significant quantities of DOPA and DOPA-like derivatives, such as 5-cysteinyldopa and 5-hydroxydopa, in their nerve nets (Carlberg et al., 1982; Carlberg, 1983, 1990; Hudman and McFarlane, 1995). In cnidarians, Carlberg suggested both that DOPA may be neuroactive in its own right and that DOPA derivatives such as 5-hydroxydopa play a role in modulating that activity (Carlberg, 1990). There is a potential thread connecting this observation and catecholic adhesive proteins, because anthozoan nerve nets have also been found to contain hydroxyarginine (Makisumi, 1961), another non-canonical amino acid that is present in mussel byssus adhesive (Papov et al., 1995). Chemical similarities in the nerve nets of ctenophores and cnidarians and the adhesive structures of mussels, polychaetes, and ctenophore colloblasts are intriguing, but it is unclear whether they are evidence of a deeply rooted origin for catecholic amino acids in animal proteins or whether they are a result of the labile chemistry of phenolic compounds.
The presence of catecholic amino acids in P. bachei colloblast proteins suggests a possible catechol-mediated adhesion mechanism for these organisms. Meanwhile, though the function of catecholic molecules in the ctenophore nerve net is unknown, the presence of o-diphenols there suggests that the history of neurotransmitter evolution is potentially more complicated than initially thought. Evidence of catecholic amino acids in both tissues highlights the functional plasticity and evolutionary complexity of biological catechols, as well as the potential for the resurgent study of ctenophores' unique physiology to reveal surprising insights with connections to other animal systems.
Drs. Sonke Johnson, Mark Goulian, Paul Janmey, and Kim Sharp provided comments that improved the manuscript. The authors declare that they have no competing interests. Andrea Stout of the Cell and Developmental Biology microscopy core in the University of Pennsylvania Perelman School of Medicine provided expert assistance with confocal imaging. AMS gratefully acknowledges funding from the Packard Foundation, the Structural Biology Training Grant (National Institutes of Health 2T32GM008275-26), and National Science Foundation Division of Materials Research 1351935.
Arnow, L. E. 1937. Colorimetric determination of the components of 3,4dihydroxyphenylalanine-tyrosine mixtures. J. Biol. Chem. 118: 531-537.
Baumgarten, H. G., A. Bjorklund, A. F. Holstein, and A. Nobin. 1972. Organization and ultrastructural identification of the catecholamine nerve terminals in the neural lobe and pars intermedia of the rat pituitary. Z. Zellforsch. Mikrosk. Anal. 126: 483-517.
Becker, P. T., A. Lambert, A. Lejeune, D. Lanterbecq, and P. Flammang. 2012. Identification, characterization, and expression levels of putative adhesive proteins from the tube-dwelling polychaete Sabellaria alveolata. Biol. Bull. 223: 217-225.
Benedict, C. V., and H. J. Waite. 1986. Location and analysis of byssal structural proteins of Mytilus edulis. J. Morphol. 189: 171-181.
Bjorklund, A., and B. Falck. 1973. Cytofluorometry of biogenic monoamines in the Falck-Hillarp method, structural identification by spectral analysis. Pp. 171-181 in Fluorescence Techniques in Cell Biology. A. A. Thaler and M. Semetz. eds. Springer. Berlin.
Burzio, L. A., and J. H. Waite. 2002. The other Topa: formation of 3.4.5trihydroxyphenylalanine in peptides. Anal. Biochem. 306: 108-114.
Carlberg, M. 1983. Evidence of dopa in the nerves of sea anemones. J. Neural Transm. 57: 75-84.
Carlberg, M. 1990. 3,4-dihydroxyphenylethylamine, L-3,4-dihydroxy-phenylalanine and 3,4,5-trihydroxyphenylalanine: oxidation and binding to membranes. A comparative study of a neurotransmitter, a precursor and a neurotransmitter candidate in primitive nervous systems. J. Neural Transm. 81: 111-119.
Carlberg, M., L. Laxmyr, E. Rosengren, and R. Elofsson. 1982. 5-OH-Dopa and 5-S-cysteinyldopa: new biogenic amino acids in invertebrates. Comp. Biochem. Physiol. C Comp. Pharmacol. 73: 23-25.
Claus, H., and H. Decker. 2006. Bacterial tyrosinases. Syst. Appl. Microbiol. 29: 3-14.
Coyne, K. J., X.-X. Qin, and J. H. Waite. 1997. Extensible collagen in mussel byssus: a natural block copolymer. Science 277: 1830-1832.
Cruz, L., R. W. Garden, H. J. Kaiser, and J. V. Sweedler. 1996. Studies of the degradation products of nisin, a peptide antibiotic, using capillary electrophoresis with off-line mass spectrometry. J. Chromatogr. A 735: 375-385.
Dayraud, C, A. Alie, M. Jager, P. Chang, H. Guyader, M. Manuel, and E. Queinnec. 2012. Independent specialisation of myosin II paralogues in muscle vs. non-muscle functions during early animal evolution: a ctenophore perspective. BMC Evol. Biol. 12: 1-21.
Deacon, M., S. Davis, J. Waite, and S. Harding. 1998. Structure and mucoadhesion of mussel glue protein in dilute solution. Biochemistry 37: 14108-14112.
Decker, H., T. Schweikardt, D. Nillius, U. Salzbrunn, E. Jaenicke, and F. Tuczek. 2007. Similar enzyme activation and catalysis in hemocyanins and tyrosinases. Gene 398: 183-191.
Dunn, C. W., and J. F. Ryan. 2015. The evolution of animal genomes. Curr. Opin. Genet. Dev. 35: 25-32.
Dunn, C. W., A. Hejnol, D. Q. Matus, K. Pang, W. E. Browne, S. A. Smith, E. Seaver, G. W. Rouse, M. Obst, G. D. Edgecombe et al. 2008. Broad phylogenomic sampling improves resolution of the animal tree of life. Nature 452: 745-749.
Dunn, C. W., S. P. Leys, and S. H. D. Haddock. 2015. The hidden biology of sponges and ctenophores. Trends Ecol. Evol. 30: 282-291.
Feuda, R., M. Dohrmann, W. Pett, H. Philippe, O. Rota-Stabelli, N. LartiUot, G. Worheide, and D. Pisani. 2017. Improved modeling of compositional heterogeneity supports sponges as sister to all other animals. Curr. Biol. 27: 3864-3870.
Franc, J.-M. 1978. Organization and function of ctenophore colloblasts: an ultrastructural study. Biol. Bull. 155: 527-541.
Franc, J.-M. 1985. La mesoglee des Ctenaires: approches ultrastruc-turale. biochimique et metabolique. Ph.D. dissertation, Universite Claude Bernard Lyon 1, Lyon, France.
Guerette, P. A., S. Hoon, Y. Seow, M. Raida, A. Masic, F. T. Wong, V. H. B. Ho, K. W. Kong, M. C. Demirel, A. Pena-Francesch et al. 2013. Accelerating the design of biomimetic materials by integrating RNA-seq with proteomics and materials science. Nat. Bioteclmol. 31: 908-915.
Harrington, M. J., A. Masic, H.-A. Niels, J. Waite, and P. Fratzl. 2010. Iron-clad fibers: a metal-based biological strategy for hard flexible coatings. Science 328: 216-220.
Hejnol, A., M. Obst, A. Stamatakis, M. Ott, G. Rouse, G. Edgecombe, P. Martinez, J. Baguna, X. Bailly, U. Jondelius et al. 2009. Assessing the root of bilaterian animals with scalable phylogenomic methods. Proc. R. Soc. Biol. Sci. B 276: 4261-4270.
Hudman, D., and I. D. McFarlane. 1995. The role of L-DOPA in the nervous system of sea anemones: a putative inhibitory transmitter in tentacles. J. Exp. Biol. 198: 1045-1050.
Hwang, D. S., H. Zeng, A. Masic, M. J. Harrington, J. N. Israelachvili, and J. H. Waite. 2010. Protein- and metal-dependent interactions of a prominent protein in mussel adhesive plaques. J. Biol. Chem. 285: 25850-25858.
Hwang, D. S., A. Masic, E. Prajatelistia, M. Iordachescu, and J. H. Waite. 2013. Marine hydroid perisarc: a chitin- and melanin-reinforced composite with DOPA-iron(III) complexes. Acta Biomater. 9: 8110-8117.
Ito, S., and IFPCS (International Federation of Pigment Cell Societies). 2003. The IFPCS presidential lecture: a chemist's view of melanogenesis. Pigment Cell Res. 16: 230-236.
Jager, M., C. Dayraud, A. Mialot, E. Queinnec, H. le Guyader, and M. Manuel. 2013. Evidence for involvement of Wnt signalling in body polarities, cell proliferation, and the neuro-sensory system in an adult ctenophore. PLoS One 8: e84363.
Lopez-Montes, A. M., A.-L. Dupont, B. Desmazieres, and B. Lavedrine. 2013. Identification of synthetic dyes in early colour photographs using capillary electrophoresis and electrospray ionisation-mass spectrometry. Talanta 114: 217-226.
Makisumi, S. 1961. Guanidino compounds from a sea-anemone. Anthopleura japonica Verrill. J. Biockem. 49: 284-291.
Martinez Rodriguez, N. R., S. Das, Y. Kaufman, W. Wei, J. N. Israelachvili, and J. H. Waite. 2015. Mussel adhesive protein provides cohesive matrix for collagen type-lA. Biomaterials 51: 51-57.
Mayer, A. M. 2006. Polyphenol oxidases in plants and fungi: going places? A review. Phytochemistry 67: 2318-2331.
Moroz, L. L. 2015. Convergent evolution of neural systems in ctenophores. J. Exp. Biol. 218: 598-611.
Moroz, L. L., and A. B. Kohn. 2015. Unbiased view of synaptic and neuronal gene complement in ctenophores: Are there pan-neuronal and pan-synaptic genes across Metazoa? Integr. Comp. Biol. 55: 1028-1049.
Moroz, L. L., and A. B. Kohn. 2016. Independent origins of neurons and synapses: insights from ctenophores. Philos. Trans. R. Soc. B Biol. Sci. 371: 20150041.
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.
Moroz, L. L., P. L. Williams, and A. Kohn. 2018. NeuroBase: a comparative neurogenomics database. [Online]. University of Florida. Available: https://neurobase.rc.ufl.edu [2018, November 21].
Norekian, T. P., and L. L. Moroz. 2016. Development of neuromuscular organization in the ctenophore Pleurobrachia bachei. J. Comp. Neurol. 524: 136-151.
Papov, V. V., T. V. Diamond, K. Biemann, and J. H. Waite. 1995. Hydroxyarginine-containing polyphenolic proteins in the adhesive plaques of the marine mussel Mytilus edulis. J. Biol. Chem. 270: 20183-20192.
Perone, M. J., and M. G. Castro. 1997. Prohormone and proneuropeptide synthesis and secretion. Histol. Histopathol. 12: 1179-1188.
Pisani, D., W. Pett, M. Dohrmann, R. Feuda, O. Rota-Stabelli, H. Philippe, N. Lartillot, and G. Worheide. 2015. Genomic data do not support comb jellies as the sister group to all other animals. Proc. Natl. Acad. Sci. U.S.A. 112: 15402-15407.
Presnell, J. S., L. E. Vandepas, K. J. Warren, B. J. Swalla, C. T. Amemiya, and W. E. Browne. 2016. The presence of a functionally tripartite through-gut in Ctenophora has implications for metazoan character trait evolution. Curr. Biol. 26: 2814-2820.
Qin, X., and J. Waite. 1995. Exotic collagen gradients in the byssus of the mussel Mytilus edulis. J. Exp. Biol. 198: 633-644.
Raper, H. S. 1926. The tyrosinase-tyrosine reaction. Biochem. J. 20: 735-742.
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, S. A. Smith et al. 2013. The genome of the ctenophore Mnemiopsis leidyi and its implications for cell type evolution. Science 342: 1242592.
Sanchez-Ferrer, A., J. N. Rodriguez-Lopez, F. Garci'a-Canovas, F. Garcia-Carmona. 1995. Tyrosinase: a comprehensive review of its mechanism. Biochim. Biophys. Acta 1247: 1-11.
Simion. P., H. Philippe, D. Baurain, M. Jager, D. J. Richter, A. D. Franco, B. Roure, N. Satoh, E. Queinnec, A. Ereskovsky et al 2017. A large and consistent phylogenomic dataset supports sponges as the sister group to all other animals. Curr. Biol. 27: 958-967.
Taylor, S. W., D. B. Chase, M. H. Emptage, M. J. Nelson, and J. H. Waite. 1996. Ferric ion complexes of a DOPA-containing adhesive protein from Mytilus edulis. Inorg. Chem. 35: 7572-7577.
Tranzer, J. P., and H. Thoenen. 1967. Electronmicroscopic localization of 5-hydroxydopamine (3,4,5-trihydroxy-phenyl-ethylamine), a new "false" sympathetic transmitter. Experientia 23: 743-745.
von Byern, J., C. Mills, and P. Flammang. 2010. Bonding tactics in ctenophores: morphology and function of the colloblast system. Pp. 29-40 in Biological Adhesive Systems: From Nature to Technical and Medical Application, J. Byern and I. Grunwald. eds. Springer, Vienna.
Waite, J. H. 1983. Evidence for a repeating 3,4-dihydroxyphenylalanine-and hydroxyproline-containing decapeptide in the adhesive protein of the mussel, Mytilus edulis L. J. Biol. Chem. 258: 2911-2915.
Waite, J. H., and M. L. Tanzer. 1981. Specific colorimetric detection of o-diphenols and 3.4-dihydroxyphenylalanine-containing peptides. Anal. Biochem. 111: 131-136.
Waite, J. H., and M. L. Tanzer. 1980. The bioadhesive of Mytilus byssus: a protein containing L-DOPA. Biochem. Biophys. Res. Commun. 96: 1554-1561.
Wang, C. S., and R. J. Stewart. 2012. Localization of the bioadhesive precursors of the sandcastle worm, Phragmatopoma califomica (Fewkes). J. Exp. Biol. 215: 351-361.
Whelan, N. V., K. M. Kocot, T. P. Moroz, K. Mukherjee, P. Williams, G. Paulay, L. L. Moroz, and K. M. Halanych. 2017. Ctenophore relationships and their placement as the sister group to all other animals. Nat. Ecol. Evol. 1: 1737-1746.
Yu, J., W. Wei, E. Danner, R. K. Ashley, J. N. Israelachvili, and H. J. Waite. 2011. Mussel protein adhesion depends on interprotein thiol-mediated redox modulation. Nat. Chem. Biol. 7: 588-590.
Zaidi, K. U., A. S. Ali, S. A. Ali, and I. Naaz. 2014. Microbial tyrosinases: promising enzymes for pharmaceutical, food bioprocessing. and environmental industry. Biochem. Res. Int. 2014: 854687.
Zhao, H., and H. J. Waite. 2006. Linking adhesive and structural proteins in the attachment plaque of Mytilus californianus. J. Biol. Chem. 281: 26150-26158.
Zhao, H., C. Sun, R. J. Stewart, and H. J. Waite. 2005. Cement proteins of the tube-building polychaete Phragmatopoma californica. J. Biol. Chem. 280: 42938-12944.
Table A1 Pairwise Pearson's correlation coefficients (r) for colloblast microspectrofluorimetry spectra Tentacle Tentacle Albumin background pellet pellet Colloblast granules 0.664 0.939 0.475 Tentacle background 0.843 0.967 Tentacle pellet 0.702 Albumin pellet
All pairs of spectra share some similarity, but the most similar pairs of spectra (highest values of r) from within the tentacle samples are colloblast granules and tentacle AcOH/CTAB extracted pellet, and tentacle background and albumin AcOH/CTAB extracted pellet (in bold).
Table A2 Pairwise Pearson's correlation coefficients (r) for ctene microspectrofluorimetry spectra Ctene Body Albumin background pellet pellet Ctene granule 0.927 0.981 0.929 Ctene background 0.852 0.998 Body pellet 0.853 Albumin pellet
All pairs of spectra share some similarity, but the most similar pairs of spectra (highest values of r) from within the ctene samples are the AcOH/CTAB extracted pellets of ctene granules and bodies AcOH/CTAB extracted pellet and ctene background and albumin AcOH/CTAB extracted pellet (in bold).
JAMES P. TOWNSEND (1,*) AND ALISON M. SWEENEY (2,[dagger])
(1) Biochemistry and Molecular Biophysics Graduate Group, Perelman School of Medicine, University of Pennsylvania, Philadelphia, Pennsylvania 19104-4799; and (2) Department of Physics and Astronomy, University of Pennsylvania, 209 South 33rd Street, Philadelphia, Pennsylvania 19104-6202
(*) ORCID: 0000-0002-4782-6083.
([dagger]) To whom correspondence should be addressed. Email: firstname.lastname@example.org. ORCID: 0000-0002-6009-8551.
Received 9 November 2017: Accepted 7 September 2018; Published online 10 December 2018.
Abbreviations: AcOH, acetic acid; CTAB, cetrimonium bromide; DOPA, dihydroxyphenylalanine; FIF, formaldehyde-induced fluorescence; L-DOPA, L-3,4-dihydroxyphenylalanine; o-diphenols, ortho-diphenols; PBS, phosphate-buffered saline; PFA. paraformaldehyde; TCA. tricholoracetic acid.
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|Author:||Townsend, James P.; Sweeney, Alison M.|
|Publication:||The Biological Bulletin|
|Date:||Feb 1, 2019|
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