Hydra magnipapillata strain 105 was used for all the experiments carried out in this study in compliance with animal welfare laws and policies (Austrian Law for animal experiments, TVG 2012, §1). Permanent mass cultures were bred and kept at 18 °C in growth chambers, and day/night light cycle at the Institute of Zoology, University of Innsbruck. Hydra cultures were fed five times per week with freshly hatched Artemia nauplii as previously described . Under these conditions, animals remained asexual and reproduced by budding. We selected animals that had at least one bud. Animals were starved for 24 h before experiments. Before fixation, animals were relaxed in 2 % urethane in culture medium for 2 min.
For bright field or differential interference contrast visualization, processed samples were examined with a Leica DM5000. Images were taken with a Leica DFC495 digital camera and a Leica LAS software.
Footprint: To collect Hydra’s footprints, polyps were placed onto glass slides and allowed attach for 30 min. After this period, polyps were gently detached with the help of a glass pipet. Glass slides bearing footprints were rinsed three times with Hydra culture medium before staining. Fresh footprints were stained using a 0.05 % solution (in culture medium) of Crystal Violet, and rinsed in culture medium.
Squeezing preparations: Living polyps were anesthetized in a 2:1 mixture of 2 % Urethane and culture medium, transferred in a drop onto a slide and slightly squeezed under a coverslip. The specimens were observed with interference contrast under the same microscopy as mentioned above.
Histology: Adult Hydra polyps were fixed in Carnoy’s fixative (ethanol, chloroform, glacial acetic acid, 6 + 3 + 1 respectively), Bouin’s fluid (saturated picric acid, 36 % formaldehyde, and glacial acid, 15 + 3 + 1 respectively), 4 % paraformaldehyde (PFA) in 0.1 M Phosphate Buffer (PBS)) and/or in Flemmings fixative (1 % chromium (VI) oxide, 2 % osmium tetroxide and glacial acetic acid, 15 + 4 + 1 respectively), dehydrated and embedded into paraplast or in Technovit 7100 resin. Paraplast sections (7 μm) and resin sections (3 μm) were produced with a Reichert Autocut 2030 (Reichert, Austria) and stained with hematoxylin and eosin (HE), periodic acid Schiff (PAS), or alcian blue (AB) pH 2.5.
Enzymehistochemistry: For peroxidase activity, Hydra polyps were fixed with 4 % PFA in 0.1 M PBS, stained with diamino benzidine (DAB + CHROMOGEN, Dako) post fixed either with 2.5 % glutaraldehyde or 1 % osmium tetroxide, dehydrated and embedded in PolyBed 812 resin. Semi thin sections (350 nm to 500 nm) were cut with a Leica ultra-microtome UCT (Leica, Austria) and stained according to Richardson et al. .
Lipid staining: PFA fixed polyps were stained with the fluorescence Nile Red method as whole mounts for lipid detection following method used by Gohad et al. . Negative controls were performed by exposing specimens to ETOH washes. Whole mounts were visualized under a Leica SP5 II confocal laser scanning microscope.
Two antibodies labelling basal disc cells were used: AE03 [35, 36], and 3G11  were kindly provided by the corresponding authors. The antibody staining method was slightly modified from the original protocols. Experiments were performed on whole mount, macerated cells and footprints. Samples were mounted in Vectashield (Vector), and visualized with a Leica DM5000, or a Leica SP5 II confocal scanning microscope. For super resolution microscopy, macerated cells samples were mounted in Mowiol and examined with a Leica TCS SP8 gSTED microscope system. Obtained super-resolution images were deconvoluted using the Huygens software from Scientific Volume Imaging implemented in the TCS SP8.
Whole mount preparation: For AE03 labelling, whole polyps were fixed in Zamboni’s fixative (2 % PFA, 0.2 % picric acid in 0.1 M PBS pH 7.2). For 3G11 labelling, whole polyps were fixed in 4 % PFA. Both fixations were done at 4 °C overnight. The following steps were applied to both antibodies: After three washes with PBS, the polyps were permeabilized with 0.5 % Triton in PBS for 30 min, and incubated with 0.5 % Triton, 1 % bovine serum albumin (BSA, w/v) in PBS with primary antibody (dilutions = AE03 1:5, and 3G11 1:1000) overnight at 4 °C. After this period, polyps were washed three times in PBS, and incubated for 2 h with fluorescein isotthiocyanate-conjugated (FITC) antimouse lgG (Dako) secondary antibody (1:200). Polyps were washed again three times in PBS and mounted.
Macerated cells: Basal discs (from approx. 150 polyps) were excised and incubated in 200 μl maceration medium (acetic acid, glycerol, and distilled water, 1:1:7) for 2 h at 30 °C. Basal discs were then mechanically disrupted by shearing them through the opening (roughly 1 mm diameter) of a pipette. The same amount of fixative, either Zamboni or PFA, were added to the medium containing cells and gently mixed. 50 μl of the sample were spread onto gelatine-coated slides and allowed dry for 20 min at RT. Steps for antibody staining were as for whole mount. Differences were a Triton concentration of 0.1 %, and an incubation time for the secondary antibody of 4 h. Slides were additionally counterstained with the DNA-specific fluorochrome, Hoechst 33342 (Life Technologies; 1 μg/ml). Samples examined with super-resolution microscopy were incubated with antimouse abberior STAR 488 (Abberior) secondary antibody diluted 1:100.
Footprint: Secreted material was collected on glass slides as described for light microscopy purposes. Immunofluorescence staining with AE03 and 3G11 was carried out as described above.
We tried to elucidate the subcellular binding of AE03 and 3G11, but several different immunogold approaches failed: i) post-embedded immunogold on both, cryo and chemically fixed material , ii) post-embedded immunogold on thawed cryosections according Tokuyasu , and iii) pre-embedded with horseradish labelled streptavidin conjugated antibodies.
Configuration of actin filaments in the basal disc were detected by phalloidin staining. Experiments were performed using amputated basal discs. Animals were let to attach to a glass slide for approximately 1 h and were then amputated. Basal discs were fixed with 4 % PFA for 1 h at room temperature, then washed three times for 10 min in PBS- 0.5 % Triton, and then incubated in Alexa 488 phalloidin (Invitrogen) in a concentration of 1:400 for 1 h at RT in the dark. Afterwards, they were washed three times for 10 min with PBS. Samples were mounted in Vectashield (Vector), and visualized with a Leica SP5 II confocal scanning microscope.
Scanning electron microscopy
Hydra polyps were fixed in 4 % PFA for 24 h. They were dehydrated in graded ETOH, dried by the critical point method (with CO2 as transition fluid), mounted on aluminium stubs, coated with gold in a sputter coater, and observed with a JEOL JSM-6100 scanning electron microscope.
Transmission electron microscopy
Conventional chemical fixation and cryofixation were performed basically as described by Holstein et al. . Chemical fixation: Hydra polyps were allowed to attach on a dialysis membrane, relaxed in 2 % urethane for 3 min and immediately fixed with a combined 2.5 % glutaraldehyde and 1 % osmium tetroxide fixative, dehydrated in a graded acetone series and embedded in Polybed 812 resin.
High pressure freezing (HPF) and freeze substitution
Basal discs were dissected and frozen with a HPM-010 (HPF apparatus from BAL-TEC, Baltzers, Liechtenstein), freeze substituted with acetone containing osmium tetroxide and uranyl acetate, and embedded into PolyBed 812 as previously described . Thick sections (350 nm) and ultrathin sections (70 nm) were cut with a Leica ultra-microtome UCT (Leica, Austria), mounted on copper grids and stained with uranyl acetate and lead citrate and examined with a Zeiss Libra 120 energy filter transmission electron microscope using zero loss electrons. Images were taken with a TRS 2048 high speed camera (Tröndle, Germany) and visualized through Olympus SiS iTEM 5.0 software.
Cytochemical detection of PAS-positive 1–2 vicinal diols was carried out according to Thiery . Sections from cryofixed samples were mounted on gold grids, exposed to periodic acid, thiocarbohydrazide, and silver proteinate. Negative control included omission of periodic acid treatment.
Peroxidase activity was detected in Hydra polyps fixed with 4 % PFA in 0.1 M PBS. They were stained with diamino benzidine (DAB + CHROMOGEN, Dako), and post fixation with 2.5 % glutaraldehyde/1 % osmium tetroxide, dehydration and embedding into Polybed 812. Images were taken without any further section post-staining. Controls included the inhibition of peroxidase activity by incubating samples in 3 % hydrogen peroxide for 20 min.
Electron energy loss spectroscopy (EELS) and electron spectroscopic imaging (ESI) were performed on ultrathin sections in order to detect element distribution in structures rich in nitrogen (N) and phosphorus (P) within the basal disc cells. The EELS charts and ESI images were collected using the software iTEM 5.0©. Distribution of N and P was measured according to a three-window power law difference ESI model with an energy slit width of 15 eV. Here, two background images and one image at the ionization edge of the appropriate element, K edge 397 eV for N and L2,3 edge at 129 eV for P were taken. Maximum element distribution was finally mix mapped with an inverted high contrast image taken at 250 eV.
The window-one was placed at 382 eV, the window-two at 350 eV, and window-three at 410 eV. The difference of the three windows coincide with the onset of the N –K ionization edge at 397 eV. The background image was set with the subtraction model obtained from the EELS analyses default, which ensures that only the energy loss from the ion under examination is mapped. Likewise, for constructing the distribution map of P, the window-one was placed at 121 eV, window-two at 110, and window-three at 153 eV. The energy-loss contribution of three windows coincide with the onset of P –L2,3 ionization edge of 129 eV.
Atomic force microscopy
Footprints from Hydra were collected on glass slides. The samples were let dry at room temperature and analyzed with Atomic Force Microscopy (AFM- Peak Force Tapping™, PFT). Data collection was achieved by applying controlled, low forces on the tip of the cantilever during imaging, which allows a direct comparison between the morphology and the adhesive properties at the nanometer (nm) scale. With the PFT method, an adhesion profile can be obtained by evaluating one force-distance curve for each pixel of the obtained image. Thereof, the adhesion force is defined as the maximum force needed to pull off the cantilever tip. The probe (silicon tip on silicon nitride cantilever—SNL, Bruker, Santa Barbara, CA, USA, k¼0.12 N/m) was calibrated on a stiff surface prior to the experiment for the measurements of the mechanical properties, in order to quantify the tip sample force.