Key words

1 Introduction

The phylum Placozoa [1] comprises flat (approx. 20–30 μm in height) discoid animals with a body size commonly less than 4 mm in diameter [2,3,4,5] (Fig. 1). One recently described species, Polyplacotoma mediterranea, can reach a size of up to 10 mm by adopting a highly ramified and highly flexible body shape [6]. In contrast, specimens of the other described species, Trichoplax adhaerens [2] and Hoilungia hongkongensis [7], as well as all other undescribed species never grow larger than 3–4 mm in diameter. The sandwich-like body of placozoans lacks any kind of symmetry but possesses a clear top-bottom polarity (Fig. 2a) [4, 8, 9]. The upper epithelium is facing the water column while the lower epithelium adheres to the substrate [2, 3, 10]. Upside-down flipped animals rotate and bring their lower epithelium back into contact with the surface quickly. Flipped animals perform this rotation by beating with the cilia of the upper epithelium. During this phase, the lower epithelium glides along itself until it regains contact with the ground [2, 3].

Fig. 1
figure 1

Different vital stages of placozoans. (a) Light microscopy image of the morphologically most simple metazoan animal, the placozoan Trichoplax adhaerens (“Grell clone”). (b) Degenerative stage of Trichoplax adhaerens (“Grell clone”). Under unfavorable conditions the upper epithelium of the animal lifts up and forms a hollow bubble (red arrow). In most cases the specimen will die shortly thereafter. (c) Degenerative stage of Trichoplax sp. H2 (“Vieste clone”). These thread-shaped stages can be frequently found in old placozoan cultures and are likely caused to some extend by unfavorable water chemistry

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Fig. 2
figure 2

Ultrastructure and feeding behavior of placozoans. (a) Schematic cross section of Trichoplax adhaerens . The typical placozoan bauplan consists of an upper epithelium, a lower epithelium and a fiber cell layer between both epithelia. The shown schematic bauplan and cell types are a synthesis of recent studies on the placozoan ultrastructure [11,12,13]. Please note that the actual number of placozoan somatic cell types is likely even higher and that each respective major cell type might summarize multiple sub-cell types. (b) Illustration of the typical placozoan feeding behavior. The animal glides over a food particle (green) to form an external digestion cavity. Digestive enzymes (yellow) are secreted into this extracorporeal feeding cavity and nutrition uptake is performed by the lower epithelium by means of phago- and likely pinocytosis. Panels a and b are modified after [40]

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The three-layered placozoan bauplan consists of at least nine differentiated somatic cell types: upper and lower epithelial cells, fiber cells, sphere cells, three types of gland cells, lipophil cells, and crystal cells [2, 3, 11,12,13] (Fig. 2a). Noteworthy, multiple other somatic (sub)cell types are awaiting their description [13, 14]. In addition to the so far identified differentiated somatic cell types, pluri- or omnipotent stem cells are found near the contact zone between lower and upper epithelium [15, 16]. The fiber cell layer, which is sandwiched between the upper and lower epithelium, plays a major role, for instance, in animal body contraction [3, 17]. The inter-connected fiber cells are the contractile elements and therefore also play a major role in animal locomotion. Lipophil cells secrete enzymes for extracellular digestion (Fig. 2b) and are exclusively found in the lower epithelium [11]. The lower epithelium also harbors three different types of gland cells (type 1, 2 and 3), which synthesize neuropeptides or mucus (in case of type 2 gland cells) [12]. Given that gland cells of type 1 and 3 possess a cilium, they have been suggested to be secretory sensory cells [12]. It is worth mentioning that type 3 gland cells can also be found in the upper epithelium [12]. The recently described sphere cells in the upper epithelium [13] include the shiny spheres, which might play a role in predator defense [18]. Finally, the so-called crystal cells are located at the margin of the animal body [11] and serve functions in gravity perception [19]. Besides morphological studies, a single-cell RNAseq study has indicated the existence of even more somatic cell types in placozoans [14].

A shared feature of all placozoans is the exceptionally high degree of body plasticity due to the absence of any kind of skeleton or other solid body parts [2, 4, 5]. Placozoans constantly change shape by contracting and relaxing their flat body which causes locomotion. Another mode of locomotion is mediated by ciliary beats of the lower epithelial cells, which is not accompanied, however, by shape changes [2]. It has recently been suggested that the amino acid glycine induces fiber cell contractions and also the activation of ciliated locomotion in Trichoplax adhaerens [20]. The locomotion activity of placozoans mirrors the vitality of the animals and is correlated to food availability and food uptake [21,22,23,24]. Locomotion pauses for different reasons, for example, during the phase of extracellular digestion of food particles [21,22,23,24]. This extracellular digestion (Fig. 2b) is carried out by release of secreted enzymes of so far unknown composition (for further details, see [23]). Although the cells involved and the underlying physiological mechanisms are yet unknown, placozoans are able to perceive light [25].

Placozoans show exceptional regeneration capacities. For instance, partial mechanical disruptions will usually heal very rapidly within minutes [26] (see Note 1). This high regeneration capacity relates to the simple body architecture, which lacks any kind of organs or other complex morphological structures, even a basal membrane and a complex extracellular matrix are missing [17]. The high regenerative capacity of the animals allows placozoans to reproduce very efficiently by binary fission, which is the dominant mode of reproduction under laboratory conditions [2, 3, 5] (see Note 2). For this type of vegetative reproduction, animals constrict in the center region of the body to form two daughter individuals of approximately the same size. The plane of each fission is orthogonal to the previous division plane [27] and a certain ratio of inner vs. marginal cells is required to trigger fission [26]. Daughter individuals stay connected for several hours by a cellular thread, which mechanically breaks when the daughter individuals move in different directions at the very end of the fission process [2, 3]. The resulting wounds heal rapidly and the original wound borders cannot be traced back under the light microscope 30 min after regeneration. The critical step of the wound healing process of both the outer margin as well as the central region of the animal body is the contraction of the adjacent epithelial cells which brings the cells within the cell layers in close contact [26].

Comprehensive studies on whole body regeneration in Trichoplax adhaerens have been conducted by Ruthmann and Terwelp (1979) [28], and by Schwartz (1984) [26]. Both studies applied procedures outlined by Miller in 1971 [29], but without citing this original work. Miller has been a methodical pioneer for modern placozoan regeneration experiments, although some of his results contradict findings by Ruthmann and Terwelp [28] and Schwartz [26]. Ruthmann and Terwelp [28] tested different chemicals (using for example colchicine or trypsin–EDTA-supplemented calcium−/magnesium-free ASW, see Note 3) to dissociate the placozoan body. Succeeding dissociation of animals into small cell clusters or even single cells, their reaggregation was studied after bringing the cells into close proximity via gentle centrifugation.

Studies from Kuhl and Kuhl (1963, 1966) [30, 31] reported first observations on the healing process of animals after cutting off a small piece from the animal body. The second key study on placozoan regeneration by Schwartz (1984) [26] focused on mechanical manipulation of the animal body. Schwartz studied the regeneration capacities of placozoan specimens after selective removal of marginal and/or center cells in a quantitative fashion and identified an approximately 20–25 μm thick circumferential marginal zone of presumably particular relevance for regeneration processes. This morphologically cryptic (i.e., macroscopically indistinguishable) zone consists of specific cell types, which cannot be found in the center of the animal. The differential cell type distribution from the edge to the center of the animal implies that cell type populations from both areas are needed to allow full recovery of animals after mechanical disruption. Schwartz also conducted a series of transplantation experiments (see Note 4). For example, parts of the marginal zone from a donor animal were transplanted into the central body region of an acceptor animal (Fig. 3). An important result from these experiments was the observation that cells from the animal margin do not de- or redifferentiate when transplanted into the center of the animal. Successfully integrated marginal body fragments led to a hole in the central part of the body, resembling a doughnut-like appearance, with marginal cells lining the central hole. Eventually, such holes were shifted (by an unknown mechanism of active cell movement) toward the outer margin of the animal where the transplanted and the original marginal zones fused [26]. Due to the microscopic size of placozoans, only body parts comprising all three cell layers have so far been successfully transplanted into an acceptor animal. Body parts consisting just of the upper or lower epithelium alone, respectively, have not yet been successfully isolated and transplanted. Schwartz [26] also showed that even small animal fragments keep their top-bottom polarity after excision, that is, that these fragments tightly reattach to the substrate by means of the lower epithelium.

Fig. 3
figure 3

Tissue transplantation in Trichoplax adhaerens . (a) From the “donor” animal, which has been stained with methylene blue, a piece consisting of marginal as well as center cells (lower left) has been cut off with an acupuncture needle. (b) The “acceptor” animal (unstained) has been prepared for the transplantation by means of cutting a hole into the center of its body. (c) The “donor” fragment is placed into the body hole of the “acceptor” animal. (d) Complete intergrowth of the “donor” fragment and the “acceptor” animal (18 h posttransplantation). Please note that the marginal cells of the “donor” fragment keep their state and form a new small margin in the central body region of the “acceptor” animal. (Picture taken from [33])

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The following protocols are based on previously published animal handling and manipulation techniques and may serve as an updated basic guideline to perform whole-body regeneration experiments in placozoans.

2 Materials

  1. 1.

    Stereomicroscope (magnification from 6.5× to 50×).

  2. 2.

    Cold light source with gooseneck light guide.

  3. 3.

    Glass petri dishes (4 cm and 15 cm diameter).

  4. 4.

    Handheld salinometer.

  5. 5.

    Food source: algae culture of Pyrenomonas helgolandii (or Rhodomonas salina) or rice grains [25].

  6. 6.

    Artificial sea water (ASW): 35 g/L commercial Ocean Sea Salt in ddH2O.

    Filter at 4 μm, adjust to pH 7.8 and leave at least 24 h to settle (see Note 5). Autoclave if needed for downstream application. Store at room temperature.

  7. 7.

    Filter paper (retention range > 4 μm).

  8. 8.

    Methylene blue stock solution (MBSS): 0.5 g methylene blue powder in 100 mL ddH2O.

  9. 9.

    Lipophilic membrane dye stock solution: 2 g dye (e.g., DiI, Invitrogen) in 200 μL 70% (v/v) EtOH in ddH2O.

  10. 10.

    Single depression microscope slides.

  11. 11.

    Acupuncture needle (0.18 mm diameter or thinner, length ideally >30 mm).

  12. 12.

    Calcium- and magnesium-free artificial seawater (CMF-ASW) + egtazic acid (EGTA): 26.24 g NaCl, 0.671 g KCl, 4.687 g Na2SO4, 2.15 mM NaHCO3, 10 mM Tris–HCl, 2.5 mM EGTA in 1 L ddH2O (see Note 6).

  13. 13.

    High-salt ASW: 1 g NaCl in 100 mL ASW.

  14. 14.

    Fixative A: ice-cold 4% (w/v) paraformaldehyde (PFA), 0.2% (v/v) glutaraldehyde in high-salt ASW.

  15. 15.

    Fixative B: ice-cold 4% (w/v) PFA, in high-salt ASW.

3 Methods

Placozoan specimens are extremely fragile which is highly relevant when working with live animals. The standard procedure for transferring placozoans from one place (e.g., culture dish) to another (e.g., microscopy slide) is to first detach the animal by means of a focused water jet gently pipetted from the side of the animal (see Note 7). Afterward, floating specimens can be pipetted into the designated place. Sometimes placozoans adhere very strongly to their substrate (the name giving feature for Trichoplax adhaerens , the “sticky hairy plate” [2, 3]), and cannot be detached without body disruptions. Another, even more disruptive, threat to the fragile animal body is the exposure to air—attached placozoans that fall dry for 1 s only get irreversibly disrupted. Therefore, the regular exchange of artificial seawater (ASW) in the culture dishes must be performed as rapidly and as cautiously as possible and an uninterrupted liquid film has to constantly cover the placozoans. Direct contact of placozoans to large underwater air bubbles, for example, as generated by algae in the culture dishes, is not harmful to the animals. Obviously, it is the rapid change of surface tension, and not the air itself, that is disruptive.

3.1 Animal Cultivation

Clonal animal material can be accessed through the corresponding author Bernd Schierwater who cultures a variety of placozoan model species in his lab. Alternatively, nonmodel placozoans can be randomly acquired from the walls of tropical sea water aquaria in pet shops, zoos, or even private aquaria. In these cases, a clonal lineage from one single starting animal has to be established and this clone has to be genetically characterized in advance of usage [32].

  1. 1.

    Fill a glass petri dish (see Notes 8 and 9) 2/3 with ASW.

  2. 2.

    Add 1 mL of food source (see Notes 10 and 11).

  3. 3.

    Cover the petri dish to minimize evaporation.

  4. 4.

    Detach placozoans from the wall of the shipment vessel (e.g., a 50 mL tube) by gently pipetting a focused water jet on their side (see Note 7).

  5. 5.

    Transfer floating placozoans into the petri dish by pipetting.

  6. 6.

    Store the culture at room temperature (ideally at 23 °C, tolerable range from 20 °C to 25 °C) under a 12:12 h light–dark cycle (see Note 12).

  7. 7.

    Every two weeks replace half of the ASW with fresh ASW to prevent the accumulation of waste products.

  8. 8.

    Add 1 mL of food source after replacement of ASW.

  9. 9.

    Repeat steps 7 and 8 until the animal density reaches 300 specimens per dish (see Notes 13 and 14).

  10. 10.

    Follow step 4 to detach 50 animals.

  11. 11.

    Pour 50% of the volume (including detached animals) to a new petri dish.

  12. 12.

    Add fresh ASW to the original and the new culture dishes.

  13. 13.

    Add 1 mL of food source to both cultures.

  14. 14.

    Repeat steps 6–13 to maintain the placozoan culture.

3.2 DiI Staining

Dyes like DiI can be used to trace placozoan cells or transplants in live specimens [27, 33]. The following protocol refers to the procedures as already described in [33] as well as own unpublished data.

  1. 1.

    Pipet 30 μL of the membrane dye stock solution onto a depression microscope slide.

  2. 2.

    Wait for 1 h for the solution to dry completely at room temperature.

  3. 3.

    Detach placozoans from their culture dish by gently pipetting a focused water jet on their side.

  4. 4.

    Transfer the detached placozoans to the dried slide.

  5. 5.

    Do not cover the slide since removal of the cover glass might disrupt the animals.

  6. 6.

    Incubate at room temperature in the dark for 7 h in a wet chamber.

  7. 7.

    Transfer stained animals into a petri dish filled with ASW to wash them.

  8. 8.

    Proceed with mechanical manipulation experiments (see Subheading 3.4).

3.3 Methylene Blue Staining

Different vital stainings, such as Janus Green, Neutral Red, Cresyl Blue, Crystal Violet, or Methylene Blue, have been successfully tested for placozoans (e.g., [26, 27, 34]). The protocol below refers to the protocol from Schwartz 1984 [26].

  1. 1.

    Transfer placozoans into a 4 cm petri dish filled with 8 mL ASW.

  2. 2.

    Add 200 μL MBSS to the dish for the liquid to become dark blue.

  3. 3.

    Stain animals for 45–60 min.

  4. 4.

    Transfer stained specimens by pipetting into a new petri dish containing fresh ASW to wash the animals.

  5. 5.

    Proceed with mechanical manipulation experiments (see Subheading 3.4).

3.4 Mechanical Manipulation

The following protocol refers to the procedures described in Schwartz 1984 [26].

  1. 1.

    Transfer live specimens into a glass petri dish prefilled 2/3 with fresh ASW to remove algae.

  2. 2.

    Wait for 5 min for the animals to attach and become completely flattened.

  3. 3.

    Cut a small piece from one (stained) animal by using an acupuncture needle (Fig. 3a).

  4. 4.

    Create a hole in the center of another animal by using an acupuncture needle (Fig. 3b).

  5. 5.

    Immediately transfer the small animal fragment by pipetting into the middle of the hole of the other animal (Fig. 3c).

  6. 6.

    Follow the wound healing/regeneration process (Fig. 3d) by visual inspection and taking pictures/time-lapse videos (see Note 15).

3.5 Animal Fixation

This protocol follows [35].

  1. 1.

    Follow steps 4 and 5 in Subheading 3.1 to transfer animals to be fixed into separate petri dishes.

  2. 2.

    Wait until all animals have settled on the bottom of the petri dishes.

  3. 3.

    Remove as much ASW as possible before adding fixative.

  4. 4.

    Carefully add ice-cold fixative A until the petri dishes are filled 2/3.

  5. 5.

    Incubate at room temperature for 90 s.

  6. 6.

    Carefully remove fixative A.

  7. 7.

    Carefully add ice-cold fixative B.

  8. 8.

    Incubate at 4 °C for 1 h.

  9. 9.

    Carefully remove fixative B.

  10. 10.

    Wash animals three times in ice-cold DEPC-treated H2O.

  11. 11.

    Immediately proceed with respective downstream protocols (e.g., in situ hybridization).

3.6 Animal Body Dissociation and Reaggregation

In general, the success of regeneration, with the eventual rebuilding of a vital whole animal, is highly dependent on the random aggregation of all cell types (potentially including pluri- or omnipotent stem cells). The protocol below refers to the procedures as described in Ruthmann and Terwelp [28] as well as Sebe-Pedros et al. [14].

  1. 1.

    Transfer 50 live animals into a petri dish containing fresh ASW to remove algae.

  2. 2.

    Transfer animals by pipetting into a new petri dish prefilled 2/3 with CMF-ASW + EGTA (see Note 3).

  3. 3.

    Wait for 2–5 min until the cells start to disperse, beginning from the margin of the animal.

  4. 4.

    Carefully pipet animals 5–10 times up and down to accelerate the complete dissociation process of the entire animal body.

  5. 5.

    Pipet cell suspension into 1.5 mL tube and add fresh ASW.

  6. 6.

    Centrifuge the dissociated cells for 2 min at 150 rcf in a table centrifuge.

  7. 7.

    Remove and discard the excess ASW from the tube.

  8. 8.

    Leave the concentrated cell suspension for reaggregation overnight.

  9. 9.

    Monitor the ongoing reaggregation process under the stereomicroscope.

4 Notes

  1. 1.

    The recently reported “spontaneously occurring body disruptions” in Trichoplax adhaerens [36] are likely the result of inappropriate culture conditions.

  2. 2.

    Neither age nor any kind of developmental stages can be deduced from placozoan morphology. Likewise, division (i.e., vegetative reproduction) cannot be precisely predicted as it does not seem to be size-related.

  3. 3.

    CMF-ASW can be supplemented with 0.1% (v/v) trypsin to improve animal body dissociation [28].

  4. 4.

    The rate of success of mechanical manipulation/transplantation experiments on placozoans is strongly related on the vitality of acceptor animals; moving acceptor animals are frequently repelling the donor fragment.

  5. 5.

    Alternatively, we have successfully used the following composition for ASW: 26.29 g NaCl, 0.74 g KCl, 0.99 g CaCl2, 6.09 g MgCl2•6H2O, 3.94 g MgSO4•7H2O in 1 L ddH2O [37]. ASW should be prepared at least 24 h in advance of usage. Ideally, ASW should settle for 3–5 days.

  6. 6.

    We took the composition of CMF-ASW from a previously published recipe [38].

  7. 7.

    The process of picking and transferring large numbers of placozoans can be simplified by stepwise adding 1 M MgCl2 or alternatively 5% (v/v) total volume of ethanol [39] into the culture petri dish. After gently shaking, animals will detach and can be pipetted in large batches into a new petri dish. Remaining MgCl2−/ethanol-enriched seawater has to be replaced afterward by regular ASW. Avoid adding an excess of MgCl2, and application times longer than 2 min, since both will increase disintegration of animals.

  8. 8.

    Rinse glassware with ASW before usage to remove remnants of detergents.

  9. 9.

    For most experiments sterile working conditions are not necessary, that is, glassware does not need to be autoclaved unless downstream analyses require sterile work.

  10. 10.

    In addition to standard algae as food sources, placozoans can also be cultured on glass slides which are covered with a natural marine biofilm. Biofilm-covered slides can be obtained by placing clean (new) slides into a seawater aquarium or (if possible) a natural marine environment for at least seven days. Before usage, the biofilm-covered slides should be washed in ddH2O for several hours to avoid placozoan contamination from the field/aquarium.

  11. 11.

    Depending on the food source, coloring of placozoans may look transparent/light white, pink, light yellow, light green, light brown, or blue. Healthy animals fed with P. helgolandii usually acquire a pinkish coloring due to the storage of breakdown products of algae digestion.

  12. 12.

    Frequently measure salinity in the culture dishes and adjust with ddH2O to keep salinity constant at 35‰ ± 2‰. Avoid dropping of ddH2O directly onto the animals.

  13. 13.

    The sporadically occurring sudden decline of a particular placozoan culture due to unfavorable conditions can be minimized by the weekly split of cultures and by keeping multiple cultures at different places.

  14. 14.

    Indicators of unfavorable culture conditions are animals with a large central bubble between the upper and lower epithelium or elongated placozoans (thread-shaped) (Fig. 1b and c, respectively).

  15. 15.

    The exposure of animals to a directional light source may cause an increase of animal movement activity, which has to be taken into account during the regeneration process.