PhD thesis - Biologisk Institut - Københavns Universitet

2KRISTOF ET AL.segmented body plan rather than an initial evolutionary steptoward segmentation in these animals. Interestingly, the lack ofcertain annelid key features, such as segmentation, coelomiccavities, nuchal organs, and chaetae, is also known for a varietyof interstitial, parasitic, and sessile annelid representatives, but toa much lesser extent for large burrowing forms, such asearthworms (reviewed in Bleidorn, 2007).Segmentation is usually considered a concerted repetition oforgans or organ systems that form subsequently from a posteriorgrowth zone along the anterior–posterior axis of an animal(Willmer, ’90). Studies on the cell proliferation patterns and theontogeny of organ systems typically associated with annelidsegments (e.g., subsequently emerging sets of paired perikaryaassociated with the ventral nerve cords, body wall ring muscles,nephridia) have proven to be ideal markers to assess whether ornot a taxon has derived from a segmented ancestor (Müller andWestheide, 2000; Hessling, 2002, 2003; Hessling and Westheide,2002; de Rosa et al., 2005; Seaver et al., 2005; Bergter et al.,2007; Brinkmann and Wanninger, 2008; Kristof et al., 2008;Wanninger, 2009). Furthermore, the growing number ofimmunocytochemical studies on neuro- and myogenesis of anumber of lophotrochozoan taxa allows for a comparison ofnervous and muscle system patterning pathways in putativelysegmented and nonsegmented clades (e.g., Hay-Schmidt, 2000;Croll and Dickinson, 2004; McDougall et al., 2006; Bergter et al.,2007; Wanninger, 2008; Wanninger et al., 2008, 2009). Althoughsome variation in annelid segment formation has been reported(Seaver et al., 2005; Brinkmann and Wanninger, 2010), thesegmentation process driven from a posterior growth zone isconsidered to be the ancestral condition for Annelida (Anderson,’66; de Rosa et al., 2005; Seaver et al., 2005; Wanninger et al.,2009). Herein, we compare the tempo–spatial distribution ofproliferating cells in Thysanocardia nigra and Themiste pyroideswith growth patterns reported for annelids. We supplement thiswork with data on myogenesis in Phascolosoma agassizii,T. nigra, and T. pyroides, and thus provide insights into theevolution of the myogenic bodyplans within the Lophotrochozoa.MATERIAL AND METHODSAnimalsAdult P. agassizii were collected in the vicinity of the FridayHarbor Laboratories (Washington) and were kept in thelaboratory until gametes were released. After fertilization, thedeveloping larvae were maintained in natural seawater atambient temperature (101C). Development was followed until15 days post-fertilization (dpf) when animals had reached the latepelagosphera stage. Adult specimens of T. nigra and T. pyroideswere obtained from Crenomytilus grayanus mussel beds. Musselaggregations were collected by scuba divers at depths of 4–8 mfrom the Vostok Bay, Sea of Japan (Russia). Adults were placed insmall tanks (15–30 specimens each) with ambient seawater(20–221C) until spawning occurred. In addition, fertilizationexperiments were performed. Adult specimens were cut open andgametes were transferred into glass jars. The eggs were fertilizedwith a few drops of a diluted sperm suspension. Embryos, larvae,and juveniles were reared in Petri dishes and glass vessels at17–191C. Elongation of the anterior–posterior axis started at3 dpf in both species. Metamorphosis occurred at 10 dpf inT. nigra and at 15 dpf in T. pyroides. Development was followedin both species until the first juvenile stages, which alreadyshowed the anlagen of the primary tentacles (i.e., at 18 dpf inT. pyroides and 15 dpf in T. nigra).EdU Labeling, F-Actin Staining, Confocal Laserscanning Microscopy,and 3D ReconstructionProliferating cells were visualized by in vivo labeling with thenucleotide analogue 5-ethynyl-2 0 -deoxyuridine (EdU) that isincorporated during the synthesis phase (S-phase) of the cell cycle.Larvae were incubated in EdU (Invitrogen, Taastrup, Denmark),diluted in filtered seawater in the following concentrations andtime intervals at 17–191C: 250 mMfor1hr,8mMfor6hr,and5mMfor 24 hr. After EdU treatment and before F-actin staining, larvaewere anesthetized by adding drops of a 3.5 or 7% MgCl 2 solution tothe seawater and were subsequently fixed in 4% paraformaldehydein 0.1 M phosphate-buffered saline (PBS) (pH 7.3) for 1.5 hr at roomtemperature or overnight at 41C. This procedure was followed bythree washes (15 min each) in 0.1 M PBS (pH 7.3) with 0.1% sodiumazide (NaN 3 ). Until further processing, samples were stored in 0.1 MPBSwith0.1%NaN 3 at 41C.After storage, larvae were rinsed in 0.1 M PBS (pH 7.3) formore than 6 hr, followed by incubation in a blocking andpermeabilization solution (saponin-based permeabilization andwash reagent with 1% BSA) overnight at 41C. Incorporated EdUwas detected with a Click-iT EdU Kit (Cat] C3005, Invitrogen,Taastrup, Denmark). The larvae were incubated with the reactioncocktail provided by the supplier (7,5 mL Alexa Flour 488 azide,30 mLCuSO 4 ,1313mL EdU reaction buffer, and 150 mL EdU bufferadditive) for 24 hr at 41C. Some specimens were additionallyincubated for 24 hr at 41C in a polyclonal rabbit anti-serotoninprimary antibody (Calbiochem, Cambridge; dilution 1:200). Thespecimens were rinsed three times for 15 min in PBS. This wasfollowed by incubation with a goat anti-rabbit Alexa 594secondary antibody (dilution 1:300; Invitrogen, Taastrup, Denmark)in 0.1 M PBS for 24 hr at 41C. Finally, the specimens werewashed three times for 15 min each in the saponin-based washreagent with 1% BSA, incubated with the nucleic acid stain4 0 , 6-diamidino-2-phenylindole (DAPI; 1:10 dilution; Invitrogen,Taastrup, Denmark), and mounted in Fluoromount G (Southern-Biotech, Birmingham, Alabama) on glass slides.For F-actin labeling, the stored larvae were washed threetimes for 15 min in 0.1 M PBS, permeabilized for 6 hr in 0.1 MPBS containing 4% Triton X-100 at 41C, and incubated in a 1:40dilution of Alexa Flour 488 phalloidin (Invitrogen, Taastrup,J. Exp. Zool. (Mol. Dev. Evol.)