Specific markers for cell nuclei
Introduction
Observing a live, moving Zoea under a fluorescence microscope with every nucleus shining blue after a vital stain is a fascinating sight. In the following, we will show that it is rather easy to obtain a larval whole mount with cell nuclei stained, dead or alive (Fig. 5). Common nuclear markers that can be used for the technique are, e.g., DAPI® (4′,6-Diamidino-2-phenylindole), TO-PRO-3®, and SYTOX Green®, each covering a specific section of the spectrum of light (reagents available, e.g., from Sigma, Thermo Fisher). DAPI absorbs light in the UV (358 nm) and emits in the blue part of the spectrum (461 nm), SYTOX® Green has a maximum excitation wavelength at 488 nm and emits at 523 nm, and TO-PRO-3® is active at longer wavelengths (excitation at 642 nm, emission at 661 nm). These stains are universally used in many different fields of biosciences, but until now, they are not very common in studies of decapod development.
Description of the method
For vital stains, we recommend a protocol originally designed for Chaoborus larvae (Diptera) as described in [65], and subsequently used for Zoeae [66]. Larvae are kept in small vials in sea- or freshwater (depending on the natural habitat of the animals), and then a few drops of DAPI stain solution (ca. 1 mg DAPI in 10 ml distilled water) are added. After an incubation period of between 30 min to 1.5 h in darkness, the nuclei are stained and the larvae can be studied under the microscope. Where appropriate, 7% of magnesium chloride can be used to anaesthetise the larvae.
Chemical fixation of the larvae can be done either before or after staining. Unstained chemically fixed larvae should be incubated in the fluorescence marker shortly after fixation since otherwise, the staining reagent will penetrate less well as the tissues’ biochemical composition becomes modified due to the action of the fixative. As fixatives, almost all formalin or ethanol-based mixtures like formalin-seawater or graded ethanol series can be used (see companion paper [8]). For imaging, the fixed specimens are mounted in an aqueous mounting medium suitable for fluorescence studies. Several companies such as Sigma or Thermo Fisher offer fluorescence media that already contain DAPI in an adequate concentration into which freshly fixed larvae can be embedded. After incubating larvae for ten to 30 min in these media, the nuclei of the specimen will also be stained with DAPI.
Additional comments
In cell biology and histology, fluorescent nuclear stains are commonly used as background stains for immunofluorescence labelling experiments that use specific antibodies. They help to understand the architecture of the studied organs in multichannel fluorescence analyses. Moreover, these stains can be used for nuclear and/or cellular pattern analyses (e.g. [66]). We recommend employing this technique routinely as a primary tool in developmental studies of decapod larvae to study, e.g., the number and size of nuclei in certain organs or the presence or absence of clusters of densely accumulated nuclei, viz., of embryonic cell clusters. We consider it a worthwhile but scarcely used approach, e.g., to find out whether these clusters, similar to insect “imaginal discs”, provide the source of morphogenetic waves of mitoses that bring about new components of the body during moults or during metamorphosis from Zoea to Megalopa. In addition to nuclear stains, there is a wealth of other fluorescent markers that allow to directly label organs that have hardly been used for larvae. For example, labelling tissue with phalloidin conjugated with a fluorescence marker was employed to study embryonic muscle development of crustaceans (review [1]) and may now be used for developmental studies on larvae.
An oldie, but still a goldie—classical histology
Introduction
Classical histology is used to study biological tissues to describe an organism’s microscopic anatomy and has acquired major importance since the development of the microscope in the 15th–16th century [67]. A detailed morphological and anatomical visualisation of tissues can be achieved by this technique at the micrometre scale which is essential for functional, comparative, and developmental studies of all organisms. Furthermore, using differential staining methods allows to characterise complementary histological features of the selected organs. Despite its long history and numerous competing techniques, such as micro-computed tomography or immunohistochemistry (see other sections of this paper) to analyse tissues, classical histology may still be seen as the gold standard and a frequently used approach to analyse tissues of adult crustaceans (summarised in Table 2 in [22]; see also [68,69,70,71]). Especially for small animals, this approach allows gaining insights into anatomical and morphological structures by sectioning whole specimens where other preparation or dissection is virtually impossible. To this day, serial sectioning of entire specimens is an approach regularly used to investigate morphology, anatomy, and development of decapod crustacean larvae (summarised in Table 1 of the companion paper [8]).
We summarise here the principal steps required from a living larva to a histological image. Classical histology is generally characterised by a suite of the following processes: (i) preparation, (ii) fixation procedure, (iii) dehydration and embedding, (iv) sectioning and staining, and (v) imaging session [72]. Steps (i) and (ii) were already described in the companion paper [8]. However, processing for transmission electron microscopy (TEM) using chemical fixation needs various additional steps, as will be detailed in the following, most of which are less important in solely light microscopic analyses. In general, microtomy (the technique of cutting an organism or single tissue into thin sections to allow for further observation under a light or electron microscope) determines the quality of tissue analysis because the thickness of the single sections and thus the resolution of the sample is defined by this step. Several modes of microtomy are classified in accordance with the section thickness, the microscopic technique, or the used microtome [72], for example: (1) histological sectioning (2–10 µm; Fig. 6); (2) semi-thin sectioning (0.25–2 µm; Figs. 7, 8, 9); (3) ultra-thin sectioning (50–150 nm; Figs. 10, 11).
Histological sectioning based on paraffin wax (e.g., Paraplast®) embedded tissues presents a rapid and efficient technique to observe the overall tissue organisation. A rotary microtome is needed which is a standard piece of equipment in many histology and pathology labs. Followed by classical staining techniques, such as haematoxylin–eosin (HE) or Azan, it offers a convenient way to examine samples and differentiate larval organ systems, even with the simplest bright field light microscopes (see Table 1). Unstained histological sections can also be used in a variety of studies including in situ hybridisation or immunohistochemistry. Because classical histological sections are thicker (2–10 µm) in comparison to the other techniques, this method provides an additional aspect of “depth” in the z-direction and a better insight into the coherence of tissues. Moreover, the reagents used in histological sectioning mostly are less toxic (and expensive) and therefore safer to work with. For the analysis of the internal morphology of marine crustacean larvae and juveniles, we also obtained excellent results by using histological semi-thin sectioning (section thickness of 1–1.5 µm) of whole specimens in combination with bright field microscopy. This technique needs a higher quality microtome and profits from the use of expensive diamond knives (see Table 1). Because the specimens have to be embedded in plastic resins, it is more time consuming than the former technique. Serial section series obtained with these techniques can be used to perform 3D reconstructions.
The transmission electron microscope (TEM) is one of the mightiest (and most expensive) imaging devices in life sciences with a resolution that by far surpasses that of light microscopic methods surveyed here and, therefore, allows to analyse ultrastructure in great detail. This technique is therefore fundamental to understand the cell’s ultrastructure including type, morphology, and distribution of the organelles and the cytoskeleton, and also helps to identify the cells responsible for the secretions (reviews, e.g., in [72, 73]). However, the process from the larva to the TEM image is complicated, time consuming and needs expensive chemicals and machinery (see Table 1). Hence, one should carefully consider whether the method is needed for a specific research question. For example, features such as the number of cells in an organ or analyses of the segmental organisation, or muscle fibre projections can be also unveiled with easier and faster methods than TEM. Nevertheless, whenever the development of minute structures such as sensilla, ommatidia, glands, synaptic connectivity in nervous systems and, in general, differentiation processes on the cellular level are to be studied, the TEM is the right method at hand (e.g., [74,75,76] for sensilla in general, and [77, 78] for crustacean larvae). Two main preparation techniques are used in TEM studies, the first is based on chemical fixation and represents the standard (see Table 1 in the companion paper [8]), whereas the second is based on cryofixation and is more sophisticated (surveys [72, 73, 79,80,81]). Cryotechniques circumvent some of the artefacts of chemical fixation, but are mainly useful for very small-sized specimens or objects in the 10-µm range. However, if cryofixation works, more detailed analyses, a higher resolution than with the standard protocol, and therefore even analyses of macromolecules are possible [82]. As Zoea larvae are indeed very small and include numerous minute structures, this approach should maybe be used more often, especially because high-pressure freezing extends the range of suitable specimen size into the reach of Zoea larvae or parts of them. A simple method that does not need costly equipment and can be established in any laboratory is described by Steinbrecht [83]. As the authors of this paper are not experienced in cryo-TEM, we will focus on the standard technique here.
TEM analyses using chemical fixation procedures are intricately connected with their sister technique, the semithin sectioning combined with light microscopic analysis. The latter approach originally was an aid to approach the region of interest in a specimen embedded for TEM, instead of making countless ultrathin sections. However, in times of serial 3D reconstructions, it has become a powerful tool as well (e.g., [84]). Here, the principle of reciprocity is also valid, because the quality of a 3D reconstruction based on semithin sections will improve when intermittent ultrathin sections are checked in the TEM to confirm what is ”really seen” with the light microscope. That way, it is possible to access where the real borders between organs are located or what the real functional specifications of a cell or a group of cells might be. However, various studies are using only one line of morphological evidence (e.g., SEM), to make statements about, for example, sensillar functions. Many studies would profit from adding TEM analyses to SEM, because the crucial structure–function relationships of sensilla can be better analysed by TEM since cellular and dendritic ultrastructures have to be considered. There are even sensilla that are not visible at all under the SEM [77, 78].
Description of the method: paraffin embedding, sectioning and staining (Azan)
Paraffin embedding
Prior to fixation, living larvae should be anaesthetised by a short exposure to low temperatures (see companion paper [8]). Before applying this method, animals must be immersed into the fixative and different appendages should be dissected (e.g., rostrum, dorsal spine, or limbs) for better penetration of the fixative into larval internal tissues (see paragraph on “fixation” in the companion paper [8]). We frequently used Bouin’s fluid (i.e. 40% formaldehyde, 5% glacial acetic acid, 2% saturated aqueous picric acid) to preserve the animals. Animals completely immersed in the fixative should be stored at 4 °C for at least 24 h. The larvae are then removed from the fixative and different methods can be applied for easier fluid exchange, depending on the size of the larvae. Larger specimens can be transferred into labelled vials and solutions can be exchanged using pipettes. However, for small larvae such as those of Carcinus maenas, we recommend using white embedding cassettes (Routine VI, Loose, Leica Biosystems; Fig. 6d, e) combined with the SurgiPath® Bio-Wraps (Leica Biosystems). White embedding cassettes are great tools to save time and reagents because different samples can be marked separately with a pencil on the cassette itself and placed together in one beaker (Fig. 6d). To apply this technique, cut Bio-Wraps as squares to overlay the whole area of the embedding white cassette, with one centimetre extra on each border of the cassette (Fig. 6e). Place the larvae in the embedding cassette and close it with the lid merged with Bio-Wrap in a way that there is a surplus of the Bio-Wrap on each border (Fig. 6e). This technique presents an excellent substitute during solution exchange and washing on the orbital shaker since Bio-Wraps are permeable for solutions. Moreover, larvae do not stick to the wrap which thus prevents the larvae to wash through the pores of the cassette during the washing steps with agitation. Once the larvae are placed in the cassette, start the dehydration process of the samples by submerging them in ascending ethanol concentrations: 70%, 80%, 90%, 99% for 1–2 h in each step (we recommend four different concentrations, but additional steps with other concentrations could be added) at room temperature with moderate agitation (e.g., using a laboratory orbital shaker). Note that the first ethanol step (70%) can be repeated several times until all fixative is removed. For instance, in the case of Bouin's fluid, washing in 70% ethanol should be repeated until there is no trace of yellow colour leaching out. Especially for larger samples, washing steps can be extended to ensure successful dehydration. During all the washing steps cover the beaker with, e.g., aluminium foil or Parafilm® to minimise the evaporation of the washing solutions. After the last ethanol step, submerge samples in a mixture of Tetrahydrofuran (THF) and 99% ethanol (ratio 50:50) and leave samples at room temperature overnight with medium agitation. Tetrahydrofuran is an intermediary medium that is completely miscible with paraffin and potential residues of water in all proportions. Its use ensures the complete removal of water which results in a successful embedding in paraffin. An advantage of THF is that its vapours are much less toxic than other compounds that were widely used in the past such as dioxane (diethylene dioxide). After 24 h, exchange 50:50 THF + 99% ethanol solution and wash larvae for another 6 h on a shaker. Afterwards, place the samples in clean THF and leave overnight at room temperature with medium agitation. During this step, heat paraffin granules to 60 °C using an oven or other heated chamber to ensure good fluidity of the paraffin for the next day. We use Roti®-Plast with DMSO (Carl Roth Gmbh + Co. KG) granules but any other commercially available paraplast brands for histology can be used (note that melting temperatures might vary). Incubate samples in the mixture of THF and paraffin (ratio 50:50) in a 60 °C oven and let this mixture infiltrate overnight. After infiltration of the mixture, remove the larvae from the embedding cassettes and transfer them into a fresh change of melted paraffin. Considering the viscosity of paraffin, during this step, it is recommended to remove the larvae from the cassettes and put them directly in marked glassware filled with pure paraffin (for better mixing with paraffin and easier embedding later). In order to do so, pick up the cassettes with the larvae from 50:50 THF + paraffin (wearing gloves), open the cassettes, and pick the larvae with a plastic pipette (if paraffin is still liquid) or scrape them with forceps as the paraffin solidifies. Optimise this technique based on the size of the larvae and fluidity of the 50:50 THF + paraffin solution. It is important not to touch the specimens directly and minimise the surplus solution uptake during this step. Pure paraffin series can be exchanged several times until the smell of THF is gone to be certain that the samples are in clean paraffin. Because paraffin hardens quickly at room temperature, these exchanges need to be done in the heated chamber promptly. When the samples are in pure paraffin, they are ready for embedding, and we recommend not to keep for longer than a day in this medium. Prior to embedding, metal base moulds (Carl Roth + Co. KG, 52 × 35 × 11 mm outer part, 23 × 36 mm inner part; Fig. 6d) need to be warmed up to 40 °C. Prepare an alcohol burner and heat a dissection needle. Due to the fast hardening of paraffin, these next steps should be completed quickly and preferably by two persons (one person heating the needle and smoothening the surface of the paraffin and the other pouring melted paraffin and positioning samples). The first step is to pour melted paraffin into the warmed-up embedding mould so that it covers most of its area. Using a pipette, place the sample into paraffin while positioning it centrally at the bottom of the mould. We recommend arranging two to three larvae close to each other without direct contact for easier sectioning afterwards. Larvae can be positioned as desired using hot metal tools such as needles with gentle movements as the paraffin is hardening. Smoothen the surface area with a hot dissection needle to ensure that the paraffin remains liquid. Remove the lid from the embedding cassette and press the cassette into the metal mould onto the melted paraffin with the bulging part facing the liquid paraffin so that it flows through the pores of the cassette. Add more melted paraffin to the top brackets of the embedding cassette. To ensure a good attachment of the paraffin to the cassette as it solidifies, it is important to remove any residual bubbles at the top area close to the pores by using a hot needle and pushing these bubbles to the surface of the paraffin. Residual bubbles in the solid paraffin can cause the sections to rupture or can result in breaking the sample during sectioning. Leave the prepared moulds to solidify at room temperature (for faster solidification, place moulds in the fridge). When the paraffin is completely solid, cut it between the metal and plastic border (e.g., with a razor blade) and carefully separate the metal embedding mould from the embedding cassette. If a bad attachment of the samples occurs, place the whole mould again in the 60 °C chamber until the wax is liquid again. Remove the old embedding cassette carefully without disturbing samples at the bottom of the mould and repeat the steps using a new plastic embedding cassette. A hot dissection needle can be used again to rearrange the samples as desired.
Histological sectioning
Prior to sectioning using a rotary microtome (e.g., a Leica Microtome RM2145), a heating plate needs to be warmed up to 40 °C. Mark the cleaned microscope slides and warm them up on the heating plate. To decrease the water surface tension for an even distribution of water on the slide, place a droplet of glycine on the slide and spread it equally over the whole slide (e.g., using a paintbrush). Before sectioning, trim the solidified paraffin block with a razor blade in a trapezoid shape to reduce the surplus paraffin medium around your specimen. The blocks may be cooled for better trimming [85]. Make sure that the steel blade on the microtome is new and without any damage or physical scars to avoid section ruptures during sectioning. Position the block onto the microtome holder as desired and start trimming the block until the sample becomes more visible. Preservation in Bouin’s fluid stains the samples with yellow colour, and since the yellow Bouin’s fluid penetrates deep into the tissue (especially if some appendages are cut), the yellow stain will not disappear completely. This gives the advantage during sectioning on the microtome that the samples can be easily identified in the paraffin block. Traces of the fixative will be dissolved in the later steps after sectioning, during the washing steps on the slides. However, extremely tiny or transparent samples such as those fixed in PFA (which do not leave a colour trace of the fixative) can be stained prior to the embedding in paraffin. The most widely used solutions for this are toluidine blue, cresyl violet, or a low graded iodine solution (0.5–1% in ethanol after the final dehydration step; see previous chapter), where a drop of one of these solutions for a few seconds on the sample enhances the contrast between the sample and paraffin block. Once the sample is getting closer to the blade, switch to fine sectioning for the desired section thickness (good results can be achieved in a range of 2–10 µm). Pay special attention when the first tissue appears. Additionally, since the larvae of some species are extremely small, you can ensure the proper starting point of the sampled tissue by checking the sections under a microscope. When the first sampled tissues appear within the block, put water drops on the previously prepared heated slides and place sections onto the water surface. Since paraffin sections stick to each other, usually 3–5 sections (depending on the dimensions of the trimmed block) can be sectioned in a row, allowing for a better orientation of consecutive sections. A paintbrush, previously soaked in water, can be used to detach the sections and place them on the slides covered with water. As the heated water evaporates, the paraffin sections stretch and eventually stick to the slide. Note that transferring the sections from the blade to the slides is a tedious process and different techniques can be applied to achieve this goal es exemplified in reference [85]. Before the histological staining, the slides must be dried and can be stored at room temperature in a dark and dry place covered with aluminium foil for several months, or placed in the same way at 4 °C for even longer storage.
Staining and mounting
Once the sections are dried on the slides, any desired histological stain can be used. Numerous dyes (acidic or basic) are available and can be used for staining, depending on the tissues or cells of interest. For an overview of the sample and revealing the general tissue anatomy, complex mixtures such as Periodic acid-Schiff reaction (PAS), Masson’s trichrome, van Grieson, Azan, Mallory, etc. (reviewed in [67, 72, 85, 86]) can be used for larvae as exemplified in [23]. In principle, all stains used in analyses of adult crustacean histology such as, e.g., the popular trichromatic Masson–Goldner stain [87,88,89,90] can also be applied to sections of larval tissues (reagents are available, e.g., from MORPHISTO®). These methods complement each other and allow to analyse different cellular features of a given organ. A highly recommended compilation of staining techniques specific for crustaceans is available [85]. A differential analysis of an organ’s histology using different stains can be complemented by immunohistochemistry (see below) to specifically localise cellular components such as elements of the cytoskeleton or transport proteins. In the following, we will elaborate on the Azan staining technique after Geidies [72, 86] as an example to describe steps that typically characterise a histological stain, and which we have successfully used to reveal different tissues on the paraffin sections (Fig. 6a, b). This formalin-free trichrome stain offers a great overview of the tissue, where extended washing of the samples with water does not affect the staining itself [91]. Azan staining differentiates connective and reticular tissues (blue), cell nuclei and erythrocytes (red), and muscle tissues (red–orange).
To save time and volume of exchange solutions, the slides can be placed into slide holders and stained together in one tray. The staining process starts by washing the slides with the attached sections in xylene for 10 min to remove the paraffin from the sections and to clear (i.e. make transparent) the tissues (any other clearing agent, such as the less toxic Roti-Histol®, can be used to remove the paraffin from the sections). Transfer slides into 99% iso-propanol and submerge them for 5 min to remove the xylene. Incubate slides in a descending alcohol series (we recommend a minimum of three steps: 96%, 80%, 60%) for 5 min in each concentration to rehydrate the samples. For thicker sections or bigger samples, more intermediate steps could be added, such as 90% and 70% (both for 5 min). Submerge the slides in distilled water for 5 min and afterwards incubate in a nuclear fast red solution for 30 min for selective staining. Dip the samples into distilled water and transfer them to 5% phosphotungstic acid to incubate for 5 min. Phosphotungstic acid in the dye mixture provides sufficient mordanting and also ensures the right acidity (pH: ~ 2–3) necessary for good results with the aniline blue-orange G mixture. Immerse shortly in distilled water and place slides into aniline blue-orange G solution for 10 min to stain collagen, reticular fibres and muscles. Clean the slides by shortly dipping them into distilled water, then incubate in 60% ethanol, next in 80% ethanol, and finally leave them in 96% ethanol for 5 min each step. Submerge slides in 99% isopropanol for five min and then incubate slides in xylene for at least 5 min to remove potential paraffin residues. Remove the slides from the xylene, dry them out on paper tissue and by using a piece of tissue carefully remove any residual xylene drops on the slide without touching the sample. After the xylene has entirely evaporated (after a few min), mount the slides immediately with 150 µl of Roti-Histokitt II (xylene-free polymeric mounting medium) per slide. Carefully cover samples with coverslips avoiding any bubbles and let the slides dry for at least 24 h at room temperature. Prepared dry slides can then be stored at room temperature in a dark and dry place and can last for years or even decades.
Description of the method: plastic embedding, semithin sectioning, and staining with Toluidine Blue and according to Holländer and Vaaland
Embedding in plastic resins
We obtained good results (Fig. 7) with larvae fixed in FAE fixative or Bouin’s solution (see section on fixation in the companion paper [8]). Because of the high toxicity of the fixatives and embedding media, all embedding steps need to be carried out under a fume hood. After washing the specimens in three changes (20 min each) of phosphate-buffered saline (PBS; 0.1 M, pH 7.2, 1.8% sucrose), the larvae are post-fixed for 1 h by immersion in 2% osmium tetroxide at room temperature. The specimens are then washed in three changes (20 min each) of PBS. Afterwards, the samples are transferred to 30% acetone and dehydrated through an ascending series of acetone (in concentration steps of 10%) to 100%. The dehydrated samples are transferred using micro dissecting needles in Eppendorf® plastic tubes filled with a 1:1 mixture of acetone and Araldite (Araldite® epoxy resin kit, Agar Scientific). Initially, the plastic tubes are left closed for 3 h and opened for 20 h afterwards. The completely soaked samples are then transferred into small glass dishes (embryo dishes) filled with 100% Araldite using micro dissecting needles. Also, the embedding blocks are filled with 100% Araldite. When preparing the embedding medium, the surplus of the liquid and unpolymerised 100% Araldite can be drawn into plastic syringes and stored frozen (− 20 °C) until further use. To ensure the best quality of Araldite, the syringes need to be defrosted at least 1 h before use and should be defrosted not more than three times. To avoid and remove air inclusion within the specimen and in the corners of the embedding blocks, heat the glass dishes with samples and embedding blocks to 60 °C for 15 min and subsequently expose them to a vacuum (200 mbar) for 20 min. In a final step, the samples are transferred into the pre-filled embedding blocks. Samples should be positioned at the tip of the embedding blocks using micro dissecting needles (Fig. 8a). The embedding blocks, including the samples, are then incubated for 48 h at 60 °C for final polymerisation. The viscosity of the embedding medium decreases right before polymerisation, so we recommend checking the position of the samples after 15 min and eventually correct the position using micro dissecting needles. The hardened embedding blocks should cool down for at least 30 min before further use.
Sectioning
Before sectioning, thoroughly clean the microscope slides so that the sections will adhere without any folds. To that end, the adhesion microscope slides should be immersed in 96% ethanol (or a 9:1 mixture with 25% ammonia [84]) in cuvettes for 3 days, briefly rinsed with distilled water, and immersed in distilled water for at least 1 day until further use. Transfer the microscope slide onto a heating plate at 60 °C until all water has evaporated (about 30 min). Alternatively, the glass slides may also be used in a wet state directly for section uptake. The following descriptions are based on experiences with a rotary microtome (e.g., HM355 by Thermo Fisher Scientific). The hardened embedding block should be trimmed with a razor blade into a trapezoid, creating a truncated 45° pyramid with the specimen close to front face (Fig. 8b, c). For linking sections into serial bands, the upper side of the trimmed block needs to be sealed with a 1:1 mixture of superglue and xylene (for discussion of different glues see [84]). After preparation of the embedding blocks, the sectioning knife (e.g., Histo Jumbo diamond knives by DIATOME) should be carefully cleaned with ethanol (96%) and the collecting basin needs to be filled with distilled water. In a first sectioning step, the samples are trimmed roughly (“quick & thick”, e.g., sectioning thickness: 15 μm; sectioning speed: 10 units in our device-specific setting), and trimming needs to be stopped before the embedded sample is reached by the knife. For semi-thin sectioning, the process needs to be slowed down (sectioning speed: 5 units), and sectioning thickness should be reduced (sectioning thickness: 1–1.5 μm). The single sections should stick together in a band due to the one-sided sealing with superglue [84]. The length of the section bands depends on the operator’s experience and sample size, but should not be longer than 15 attached sections as the sections unfold (and thus expand a little) when drying on the heating plate. The section band needs to be lifted carefully from the knife and left floating within the water bath. Afterwards, the microscope slide needs to get immersed carefully into the water bath. The microscope slides should be moved carefully underneath the section bands and carefully lifted out of the water. When immersing the microscope slides underneath the section chain, it is useful to reduce the surface tension of the water bath by placing a pipette crosswise in front of the microscope slide (Fig. 6d).
Staining and mounting
The staining after Holländer and Vaaland [92] (1% phenylenediamine in methanol-isopropanol) strongly stains nervous tissues. Moreover, it has been shown that phenylenediamine also stains nucleoli and cytoplasm of other cell types as well, and therefore allows visualisation of several types of tissues within a crustacean larva (Fig. 7). In this method, it is likely that the formation of highly stained oxidation products by phenylenediamine in combination with the reduction of tissue-bound osmium plays an important role [92]. As osmium tetroxide (from the post-fixation process) only stains unsaturated lipids, while phenyldiamine does not stain lipids, it could be shown that the combination of both reagents results in enhanced staining of unsaturated lipids and some staining of saturated lipids [93]. The following steps should be carried out under the fume hood due to the high toxicity of the reagents. The microscope slides with the semi-thin sections should be incubated in the staining solution for ca. 14 h. Afterwards, the microscope slides need to be taken out of the staining solution and washed three times for 20 min each with iso-propanol. The microscope slides need to be rinsed for another 3 × 5 min with distilled water and dried afterwards on a hot plate. As an alternative to the stain presented above, good results were also achieved using Toluidine blue (1% Toluidine blue, 0.25% Pyronin G in a solution of 1% sodium tetraborate, [94]). Note however, that in all available stains for plastic embedded tissues are suitable for larvae also (reviewed in [67, 72, 85]) and we describe only those stains with which we have hands-on experience. Both staining solutions described here should be prepared at least 24 h before use to ensure that all crystals are dissolved. Toluidine blue, also known as tolonium chloride, is a dye with a high affinity for acidic tissue components and organelles rich in DNA and RNA such as, e.g., the cell nucleus [95]. This stain is a fast technique since the sections need to be covered with the staining solution only for a few seconds (ca. 30 s at room temperature). For both staining techniques, it is advisable to check the results in between as these stainings are nearly irreversible and overstaining makes it difficult to analyse sections due to dark sepia or blue tones. After staining, the microscope slides should be rinsed several times with distilled water until no more dye bleeds into the washing medium, and finally dried on a heating plate (60 °C) for approximately 30 min. The final step before microscopy is mounting the coverslips on the microscope slides to protect the sections. We recommend using Roti®-Histokitt II to cover-slip the specimens. Three small droplets should be put on each microscope slide, covered slowly with coverslips (avoiding bubbles), and left to dry for 24 h at room temperature.
Description of the method: ultrathin sectioning and transmission electron microscopy
Plastic embedding
The dissection and fixation of the specimens for TEM while cooling the samples on ice are described in the companion paper [8] and briefly summarised in the introduction to Sect. 2 of this contribution. The next step, in traditional protocols, is dehydrating the specimens in a graded series of 30%, 50%, 70%, 90%, and 96% ethanol followed by 100% isopropanol, 100% propylene oxide, and then a mix of propylene oxide and resin [73, 96]. In many labs, however, this procedure was replaced by a graded acetone series, because acetone is a good solvent for epoxy resin. In addition, 100% acetone needed for the last dehydration steps can be kept free of water quite easily on granular calcium chloride. A feasible dehydration protocol would thus be 30%, 50%, 70%, 90%, 100% acetone for 10 min each, and then 3 times 20 min dried pure acetone. At the 70% acetone step, the specimen vial should be taken from the ice bath and exposed to room temperature, avoiding surrounding water, because otherwise there is a risk of accidental rehydration of the specimens. Afterwards, the specimens are incubated in a 1:1 mix of acetone and resin for 30 min to 2 h maximum, and then in the pure resin. This step can last overnight, but after a few hours, the lids of the vials should be removed to allow the acetone to evaporate. After that, the specimens are placed in fresh unused resin into embedding forms and oriented according to the desired sectioning angles. Some typical accessories for carrying out these and the following steps are shown in Fig. 11.
Osmium tetroxide, and cacodylate buffer, as well as aldehydes and the resin components, are very poisonous and have to be handled according to strict safety rules. Hence, the use of gloves and eye cover when preparing the solutions is essential. Always leave the solutions under a fume hood, and if possible, use a special “preparation hood” where also the dissecting microscope is placed.
Preparing the epoxy resin needs some preliminary considerations, since resins of different hardness can be obtained depending on the proportion of the resin’s components. In most epoxy resins (Epon, Araldite®, Glycidether 100, etc.), there is a soft component and a harder one. Hence, it is possible to prepare resins whose hardness matches the needs of the specimens to study. After the two resin components are mixed together, the accelerator/activator is stirred in according to the manufacturer’s instructions. For soft specimens, medium to soft resin is suggested, for hard ones (e.g., those containing hard arthropod cuticle), a harder resin is recommended.
Once the specimens are placed in resin into the embedding form, they are heated in an incubator or heating cabinet at 40 °C. Initially, the resin becomes more fluid due to the rise in temperature which can improve its infiltration. After about an hour, before the resin becomes waxy, there is still opportunity to improve the specimens’ orientation within the embedding form, if necessary. After that, the specimens are put back into the oven and the temperature is increased to 60 °C overnight so that the resin polymerises completely. After removing the specimens from the incubator, they should be left in the embedding form until they have cooled down to room temperature. Fresh from the incubator, they are still quite soft and flexible but at room temperature, they will become hard. An alternative to this basic technique is vacuum embedding [97], a technique which is often successful in specimens that prove difficult to be embedded using the standard protocol.
Sectioning
Trimming, sectioning, and staining are in principle quite similar to semithin sectioning as described above (Fig. 8). Nevertheless, some specific tricks and techniques to obtain good ultrathin sections will be described in the following (see also Fig. 11). An ultramicrotome (e.g., Leica Microsystems) instead of a simple rotation microtome is needed and a diamond knife for ultrathin sections (e.g., DIATOME®) is strongly suggested instead of glass knives. With a diamond ultrathin knife, good sections can be made as a standard.
In the following, we refer here to some aspects which in our experience are crucial for a successful ultrathin sectioning and subsequent TEM analysis. As suggested in the previous section about semithin sections, the trimming (with a razor blade) should create a truncated 45° pyramid with the specimen close to the front face having the shape of a trapezoid. In small arthropods, such as larvae, it is usually better not to trim up to the specimen but leave a little resin edge around. It is important that the two parallel edges of the trapezoid, which will become the upper and the lower edge of the sections, are perfectly parallel to get straight ribbons of ultrathin sections instead of spiral-shaped ribbons. It is important to practice adjusting the block in reference to the diamond knife. It needs much experience to operate the micrometre screws for adjusting the block and the knife in combination with the upper and lower illumination devices of the microtome. This allows to precisely adjust all three angles using the “shadow line” between block and knife to allow for effective sectioning. With a well-trimmed block and a good knife, sectioning of very good ultrathin sections is not as difficult as beginners often think. For high resolution, the sections should be really thin, of grey or silver interference colour (40–70 nm). For relatively low resolution, sections of golden interference (70–90 nm) are sufficient. In ultrathin sectioning, silver sections are, therefore, more of a gold standard than gold sections. As in semithin sectioning, another essential issue is “fishing” the sections from the water bath attached to the knife and transferring them to the grids in perfect orientation. This can be done from above or below—resulting in mirror-symmetrical pictures in the TEM—with the help of formvar coated copper “slit” grids and a clean eyelash glued to a tooth stick, a basic tool of the ultramicrotomist. It can be used to split ribbons of sections into handy pieces and to orientate the sections along the grid edge. Another important trick is to stretch the sections with a little bit of chloroform vapour emitted by a piece of moist filter paper held close to the grid but not touching it. One more parameter is essential for the entire procedure, the sectioning angle, usually indicated for each diamond knife in the manufacturer’s instructions. However, on some occasions, this angle should be slightly adjusted. If the section thickness is not homogenous within a ribbon of sections, increase the angle; if there are fissures or cracks within the sections, decrease the angle.
Additional comments
Despite its long history and numerous competing technical approaches, classical histology is still a commonly used method to investigate organisms’ cells and tissues. While histological (paraffin) sectioning is a fast and fairly simple approach to analyse organs of interest, higher-resolution methods, such as semi-thin and eventually ultra-thin sectioning provide additional insights into cells and tissues of organisms only a few millimetres in size such as crustacean larvae (Figs. 6, 7, 10). Another advantage of classical histology is the long-term stability of the finished samples. When stored correctly (dark, cool, dry), the sections (as well as the embedded blocks) can last for decades. Nevertheless, classical histology is technically challenging, and a highly invasive method as it leads to alteration and destruction of the specimen. For instance, paraffin embedding is not suitable for extremely hard samples, such as strongly calcifying organisms, where due to softer paraffin, samples usually break during the sectioning process. Furthermore, this procedure is prone to artefacts such as section loss, distortion, and staining artefacts. In addition, classical histology is time and labour consuming, as well as costly because expensive microtomes and microscopes are needed (see Table 1). Compared to light microscopy-based histology, a TEM analysis requires more work and time, but the higher resolution power from the TEM pays off when results about minute structures and cellular ultrastructures are needed.
Overall, to date (and not only limited to samples of invertebrates), the combination of histological sectioning, staining, and microscopic assessment of 2D sections, is still the gold standard for structural and compositional analysis of biological tissues [98]. Serial sections are excellent starting material for 3D reconstructions of selected organ systems such as the nervous system (Fig. 9). In a broader perspective, it has been shown that the combination of multiple analytical methods with classical histology might be the most effective approach to gain a holistic insight into the anatomy of invertebrate organisms (e.g., [22, 23, 98, 99]) because single methods sometimes cannot fulfill all requirements. Nevertheless, the selection and combination of methods depends on the objective and certain species characteristics and therefore, should be chosen with care.
X-ray microscopy (µCT)
Introduction
The application of X-ray microscopy (µCT) to soft tissues has now become a standard in imaging techniques (Figs. 12, 13). In the field of anatomy, µCT has been demonstrated to be a useful analytical tool applied in a rapidly growing variety of metazoan taxa, such as cnidarians [100], plathelminths [101], nematomorphs [99], nematodes [102], polychaetes [103, 104], molluscs [98, 105], echinoderms [106, 107] as well as arthropods [108,109,110,111,112,113]. Modern, lab-based µCT-scanners deliver high resolution allowing the investigation of tiny specimens with body sizes of free-living crustacean larvae ranging from 75–195 µm in Tantulocarida [114], the smallest arthropods in the world, up to 5 cm in length, e.g., in Stomatopoda [115]. Since it is a non-destructive method allowing to investigate intact specimens as a whole (reviews [109, 116]) as well as in the anatomical context (Figs.12, 13), X-ray microscopy opens up numerous opportunities for correlative microscopic workflows such as the combination with serial block-face imaging [117], scanning- or transmission electron microscopy [118], including histological sectioning for light microscopy [98, 118], or the combination with magnetic resonance tomography [119] of one and the same specimen.
Description of the method
In general, imaging of biological samples based on µCT results from different absorption coefficients of tissues and of how X-ray radiation is absorbed by these samples. Thus, X-ray absorption is dependent on the atomic number based on the tissues' biochemical composition [120]. The best results in terms of resolution, reduced scanning time, and contrast can be achieved by scanning samples that were previously fixed, contrast-enhanced and critical point dried. For a better resolution of integumentary structures, dried specimens can be additionally coated analogous to the sample preparation (sputter coating) for scanning electron microscopy [121].
Here, we briefly summarise methodological steps of the sample preparation for a workflow enabling a consecutive immunohistochemical and/or histological investigation (for further details of other possible tissue preparations see our extensive manual on µCT in decapod larvae [122]). First of all, the larvae need to be properly fixed. Note that the choice of fixative has an effect on tissue preservation as well as the choice of agents for contrast-enhancement. Bouin's solution or 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS) or seawater have turned out to be useful fixatives that allow immunohistochemical labelling after µCT-scanning (see companion paper [8]). After a proper fixation for at least two hours at room temperature, the tissues need to be post-fixed in Dent's fixative (20% DMSO in methanol) and further dehydrated in two graded steps up to 100% methanol. At this stage, larvae can be stored at − 20 °C. Note that fixation with methanol disrupts or denatures binding sites of some epitopes in tissues to some extent (e.g., of F-actin resulting in a limited detectability by, e.g., phalloidin conjugated with fluorescent dyes). Ethanol or isopropanol represent an alternative to methanol. However, depending on the scientific question, any of the subsequently used staining agents need to be carefully checked in advance for their compatibility with methanol-fixed tissues. After dehydration and before scanning, the tissues should be incubated in 2% iodine in methanol for contrast-enhancement for at least 24 h at room temperature. After two to three washing steps using pure methanol, the larvae can be directly scanned ideally in sealable plastic chambers filled with fresh methanol (for further details on mounting of samples, see [122]). After successful scanning, the sample needs to be rehydrated in a decreasing series of methanol in TRIS-buffer and finally be washed in several changes of PBS-TX (0.3% Triton X-100 in 0.1 M PBS) to remove any reminiscent iodine from the tissue. After washing, the tissues are ready for any further processing for histological sectioning followed by immunohistochemical or histochemical treatments.
Additional comments
The main advantage of X-ray microscopy is that it is a virtually non-destructive method allowing to investigate intact specimens as a whole. Furthermore, this technique provides a high resolution even to the cellular level in well-fixed and contrasted specimens. Finally, X-ray microscopy opens up numerous opportunities for correlative microscopic workflows of one and the same specimen. For example, we established workflows allowing for combining µCT with consecutive immunohistochemical (see next chapter) or histochemical labelling of the same specimen, and also combining µCT of Cobalt-filled axonal projections with subsequent sectioning [123]. This way, general information on the spatial relationships of the whole internal anatomy can be assembled before using other destructive techniques such as histological sectioning. In other words, the collection of µCT-based datasets can be considered as an anatomical backup and is therefore also used for taxonomy in zoological museums or collections for digital preservation of type material being referred to as cybertypes [110, 124, 125]. For the study of type material, the undesirable brown staining of specimens due to the use of agents for contrast enhancement such as 2% iodine in ethanol is fortunately reversible. These agents can be completely removed after enduring (several days) of washing in several changes of fresh volumes of pure ethanol (Fig. 14) before transferring the samples back to 70% ethanol as often used in collections. Since µCT-imaging is a less-destructive method, it is one of the few techniques enabling scientists to have a look into the inside of, for instance, rare fossil specimens such as amber inclusions of, e.g., insect larvae (e.g., [126, 127]) or Cambrian arthropods [128]. A re-investigation by applying µCT of already described fossil crustacean larvae [129,130,131] would thus be feasible. Here, nature already embedded the samples millions of years ago.
Another advantage is that µCT produces isometric data which are favourably useful for morphometric approaches in contrast to anisometric imaging such as, e.g., microscopy of serial sections or confocal laser scanning microscopy where z-axially refractive mismatches can occur [111, 132]. Some possibilities to visualise these data sets are exemplified in Fig. 12. One of the most commonly used analytical processing is to generate 3D reconstructions of selected organs or tissues (Figs. 12, 13); and see Table 2). In principle, 3D reconstructions can also be generated from microscopic images based on section series (Fig. 9). However, image stacks based on µCT provide isometric datasets meaning that the pixel-size equals in all three dimensions (depending on the scanning-settings and magnification). Furthermore, artefacts only occur due to chemical fixation, drying of samples, or scanning. In contrast, image series generated from sections often suffer from more artefacts and need to be aligned before they can be used for 3D reconstruction. In addition, these datasets are anisometric which means that the pixel-size especially of the z-axis is ultimately dependent on the physical section thickness, or if generated using a confocal laser scanning microscope, the pixel sizes have to be manually corrected due to possible refractive mismatches caused by the immersion media.
Because µCT-imaging is, to put it simply, based on differences in tissue densities, it is less specific than, e.g., techniques using histological staining or antisera raised against a variety of antigens. The major disadvantage is of course that the acquisition, as well as the service and maintenance of a lab-based µCT-device are quite expensive (see Table 1). What is more, sample preparation and µCT, like most other techniques, also have specific disadvantages. For delivering good contrast, especially soft tissues need to be fixed and incubated in contrast-enhancing agents in advance which often cause tissue modifications such as shrinking (see [111, 122]). However, by using methanol as outlined above, we obtained excellent results for later immunohistochemical labelling against anti-synapsin (SYNORF1) in combination with histochemical detection of the cell proliferation marker EdU previously in vivo injected (Krieger, unpublished). The possibility of combining these techniques makes the protocol especially interesting for studying developmental stages such as larvae. If specimens are not intended to be used for additional staining experiments or immunohistochemistry, µCT of critical point dried samples delivers better contrasts as resulting from wet-scanning in, e.g., high-grade alcohols [22, 122]. Furthermore, by using dried samples, a specimen container becomes redundant so that the working distance can be decreased resulting in higher resolutions and shorter exposure and thus scanning times. However, dried material can be still later used for scanning electron microscopy (SEM).
