Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

Summary

Imaging the morphogenesis of insect embryos with light sheet-based fluorescence microscopy has become state of the art. This protocol outlines and compares three mounting techniques appropriate for Tribolium castaneum embryos, introduces two novel custom-made transgenic lines well suited for live imaging, discusses essential quality controls and indicates current experimental limitations.

Abstract

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. The observation of Tribolium embryos with light sheet-based fluorescence microscopy has multiple advantages over conventional widefield and confocal fluorescence microscopy. Due to the unique properties of a light sheet-based microscope, three dimensional images of living specimens can be recorded with high signal-to-noise ratios and significantly reduced photo-bleaching as well as photo-toxicity along multiple directions over periods that last several days. With more than four years of methodological development and a continuous increase of data, the time seems appropriate to establish standard operating procedures for the usage of light sheet technology in the Tribolium community as well as in the insect community at large. This protocol describes three mounting techniques suitable for different purposes, presents two novel custom-made transgenic Tribolium lines appropriate for long-term live imaging, suggests five fluorescent dyes to label intracellular structures of fixed embryos and provides information on data post-processing for the timely evaluation of the recorded data. Representative results concentrate on long-term live imaging, optical sectioning and the observation of the same embryo along multiple directions. The respective datasets are provided as a downloadable resource. Finally, the protocol discusses quality controls for live imaging assays, current limitations and the applicability of the outlined procedures to other insect species.

This protocol is primarily intended for developmental biologists who seek imaging solutions that outperform standard laboratory equipment. It promotes the continuous attempt to close the gap between the technically orientated laboratories/communities, which develop and refine microscopy methodologically, and the life science laboratories/communities, which require 'plug-and-play' solutions to technical challenges. Furthermore, it supports an axiomatic approach that moves the biological questions into the center of attention.

Introduction

The red flour beetle Tribolium castaneum, which belongs to the large family of darkling beetles (Tenebrionidae), has a long history within the agricultural and life sciences and is the second best studied model insect model organism after the fruit fly Drosophila melanogaster. During the last four decades, it became a powerful and popular insect model organism in developmental genetics, in evolutionary developmental biology and, during the last twenty years, in embryonic morphogenesis for a variety of reasons:

Drosophila and Tribolium both belong to the Holometabola, but diverged approximately 300 million years ago^1,2,3,4. While the embryonic development of Drosophila is commonly considered as highly derived, Tribolium shows a more ancestral mode of development that is found in a considerably larger proportion of insect species^5,6,7,8,9. Firstly, Tribolium exhibits non-involuted head development, i.e. its mouthparts and antennae emerge already during embryogenesis^{10,11,12,13,14,15}. Secondly, Tribolium follows the principles of short-germ development, i.e. abdominal segments are added sequentially from a posterior growth zone during germband elongation^16,17,18,19. Thirdly, Tribolium develops and later degrades two extra-embryonic membranes i.e. the amnion, which covers the embryo only ventrally, and the serosa, which envelops the embryo completely^20,21,22. Both membranes play a crucial morphogenetic²³ as well as protective role against microorganisms^24,25 and desiccation²⁶. Fourthly, the embryonically developing legs are fully functional during the larval life stage and serve as the primordia for the adult legs during pupal metamorphosis^{27,28,29,30,31}.

Due to their small size and modest demands, cultivation of Tribolium in the laboratory is fairly straightforward. Cultures of wild-type (WT) strains or transgenic lines typically consist of around 100-300 adults and can be kept within one-liter glass bottles (footprint 80 cm²) filled three to four centimeters high (about 50 g) with growth medium that consists of full grain wheat flour supplemented with inactive dry yeast. A water supply is not necessary. This allows even small laboratories to keep dozens of beetle cultures within small- or medium-sized commercially available insect incubators. Later developmental stages of Tribolium (larvae after approximately the fourth instar, pupae and adults) are easily separated from the growth medium by sieving. Synchronized embryos are obtained by incubating adults for short periods on egg-laying medium. For rapid development, beetle cultures are kept at 32 °C (about four weeks per generation), while stock keeping is typically performed at 22-25 °C (about ten weeks per generation).

Within the last decade, many standard techniques have been gradually adapted and optimized for Tribolium, as summarized in the Emerging Model Organisms books³². Of great importance are advanced genetic methods such as embryonic³³, larval^34,35 or parental^36,37 RNA interference-based gene knockdown, germline transformation with either the piggyBac^38,39 or the Minos⁴⁰ transposase system and CRISPR/Cas9-based genome engineering⁴¹. Furthermore, the Tribolium genome has been sequenced about a decade ago⁴², and is now in the third round of genome assembly release⁴³, which allows efficient and genome-wide identification and systematic analysis of genes⁴⁴ or other genetic elements^45,46. Additionally, the genomes of four other coleopteran species are available for comparative genetic approaches^47,48,49,50. In association with the sequenced genome, two large-scale genetic analyses have been performed, i.e. an insertional mutagenesis screen⁵¹ and a systematic RNA interference-based gene knockdown screen^52,53.

Fluorescence live imaging with widefield, confocal or light sheet-based microscopy (LSFM) allows to observe the embryonic morphology of Tribolium as a function of time (i.e. the morphogenesis) in a multi-dimensional context (Table 1). In widefield and confocal fluorescence microscopy, the excitation and emission light is guided through the same objective lens. In both approaches, the entire specimen is illuminated for every recorded two-dimensional plane. Hence, the specimens are subjected to very high energy levels. In LSFM, only the fluorophores in the focal plane are excited due to a decoupling of illumination and detection by using two perpendicularly arranged objective lenses (Figure 1). LSFM comes in two canonic implementations – the single plane illumination microscope (SPIM) and the digital scanned laser light sheet-based fluorescence microscope (DSLM, Figure 2) – and offers several crucial advantages over traditional approaches: (i) intrinsic optical sectioning capability, (ii) good axial resolution, (iii) strongly reduced level of photo-bleaching, (iv) very low photo-toxicity, (v) high signal-to-noise ratio, (vi) relatively high acquisition speed, (vii) imaging along multiple directions and (viii) deeper tissue penetration due to the usage of low numerical aperture illumination objective lenses^54,55,56.

LSFM has already been successfully applied in Tribolium to document nearly the entire embryonic morphogenesis⁵⁷ and to analyze the principles of extra-embryonic membrane rupture at the beginning of dorsal closure²³. To raise the attractiveness of LSFM in the Tribolium community and for insect science in general, it is of great importance to establish standard operating procedures and to improve the methods, protocols and the pool of resources to a level where the microscope becomes an ease-of-use standard tool in developmental biology laboratories, and the biological questions stay in the center of attention.

This protocol begins with the basics of Tribolium cultivation, i.e. maintenance, reproduction and embryo collection. Next, two experimental strategies are illustrated: (i) live imaging of custom-made transgenic lines and (ii) imaging of fixed embryos that were stained with fluorescent dyes (Table 2). Subsequently, three mounting techniques with slightly different purposes are explained in detail (Figure 3 and Table 3): (i) the agarose column, (ii) the agarose hemisphere and (iii) the novel cobweb holder. The protocol then explains the data acquisition procedure with LSFM. Imaging modalities and key considerations are outlined. Finally, embryo retrieval is explained and suggestions for basic data processing are provided. In the representative results, live imaging data from two novel custom-made and the Glia-blue⁵⁸ transgenic lines are shown and the respective imaging datasets are provided as a downloadable resource. Additionally, image data of fixed embryos that were stained with a variety of fluorescence dyes are presented. The discussion focuses on quality control, current limitations of the live imaging approach and the adaptation of the protocol to other species.

The protocol is written for light sheet-based fluorescence microscopes that are equipped with a sample chamber and a rotatable clamp mechanism for standardized sample holders^54,59,60, which are typically cylinder-shaped elements made of metal, plastic or glass with a diameter in the millimeter range. The protocol is also suitable for both canonic implementations, i.e. SPIM and DSLM, as well as for setups with two or more illumination and detection arms^61,62,63. The representative results show data in two spectral channels, green (illumination with a 488 nm laser, detection through a 525/50 bandpass filter) and red (illumination with a 561 nm laser, detection through a 607/70 bandpass filter), but the protocol can be expanded to three or four spectral channels.

Protocol

1. Husbandry of Tribolium Cultures

NOTE: Standard conditions are defined as an incubation temperature of 25 °C and 70% relative humidity in a 12 h bright/12 h dark cycle. For more information on Tribolium husbandry, respective guidelines are available⁶⁴. This protocol requires two different flour-based media, which can be prepared in kilogram quantities and stored for several months.

1. Prior to working with flour and inactive dry yeast, store the unopened packages at -20 °C for 24 h and then let them warm up to room temperature. This procedure eliminates potential pathogens.
2. Pass full grain wheat flour and inactive dry yeast through a sieve with 710 µm mesh size, then prepare growth medium by supplementing the sifted full grain flour with 5% (w/w) sifted inactive dry yeast.
3. Pass 405 fine wheat flour and inactive dry yeast through a sieve with 250 µm mesh size, then prepare egg-laying medium by supplementing the sifted 405 fine wheat flour with 5% (w/w) sifted inactive dry yeast.
4. Pass the respective Tribolium culture through a sieve with 800 µm mesh size. Discard the leftover growth medium. Transfer the larvae, pupae and adults to a large glass dish.
5. If no progeny culture exists, transfer around 80 larvae and/or pupae to a new glass bottle to establish a new progeny culture. If no larvae and/or pupae are present, take around 40 adults instead. Add 50 g of growth medium and incubate under standard conditions.
6. Collect around 300 adults (500-700 mg) in another new glass bottle to establish the imaging culture. Add 50 g of growth medium and incubate under standard conditions. Allow the imaging culture to settle for at least 24 h in the fresh growth medium before embryo collection.
7. Replace the growth medium of the imaging culture at least every two weeks, which can be done in conjunction with embryo collection (see 3.2). After two to three months, replace the imaging culture completely by new adults from the progeny culture.
8. If a non-homozygous transgenic line is used, curate the progeny cultures frequently by removing non-transgenic animals. Ideally, generate homozygous lines if the respective transgene is not lethal or causes sterility when present on both chromosomes. Homozygous cultures typically have a higher average fluorescence level and the overall experimental conditions are narrower, since only one genotype is present.

2. Microscope Setup and Calibration

NOTE: The Tribolium embryo has an anterior-posterior length of about 600-660 µm and a transversal diameter of about 300-330 µm. Most modern CCD cameras have a sensor chip size of about 9 mm x 7 mm, i.e. a diameter of 11.4 µm, and a pixel-pixel distance of 6.45 µm. When combined with a 10X magnification objective lens, the embryo can be imaged in toto with a pixel pitch of 0.645 µm. Most modern sCMOS cameras have a larger sensor chip size of about 16 mm x 14 mm, i.e. a diameter of 21.3 µm, and a pixel-pixel distance of 6.5 µm. When combined with a 20X magnification objective lens, the embryo can be imaged in toto with a pixel pitch of 0.325 µm.

1. Attach the illumination and detection objective lenses to the microscope. Ensure that lasers and fluorescence filters match the fluorophore absorption and emission spectra very well.
2. Attach the sample chamber to the microscope. Fill the sample chamber with PBS pH 7.4 or any other appropriate imaging buffer.
3. Switch on and calibrate the microscope. Comprehensive calibration routines and troubleshooting guidelines for custom-built light sheet-based microscopes (more specifically for the DSLM implementation) have been published previously⁶⁵. For commercial devices, follow the manufacturer's instructions very carefully. Mishandling might deplete the sample chamber of the medium and cause havoc to the instrument as well as the specimen.

3. Collection of Transgenic or WT Embryos

1. Pass the imaging colony through a sieve with 800 µm mesh size. Collect adult beetles in a new glass bottle and add 10 g of egg-laying medium. Incubate the egg-laying culture for 1 h at 25 °C and 70% relative humidity in light.
2. Pass the egg-laying culture through a sieve with 800-µm mesh size to separate adults from the egg-laying medium, which contains around 30-100 embryos. Transfer adults to a new glass bottle and add either the old or 50 g of fresh growth medium.
3. If observation should start at the uniform blastoderm stage (example dataset DS0002), incubate the egg-laying medium for 15 h at 25 °C and 70% relative humidity. To start at the beginning of germband retraction (example datasets DS0001 and DS0003), incubate embryos for 23 h at 32 °C and 70% relative humidity. If other developmental stages are desired, refer to the respective literature^64,65 and adapt incubation time and/or temperature as necessary.

4. Embryo Dechorionation

NOTE: The chorion is the outer layer of the eggshell and consists of proteins and polysaccharides. It is strongly light absorbent but not essential for proper development and can thus be chemically removed.

1. Pass the egg-laying medium through a sieve with 300 µm mesh size. Collect the embryos in a cell strainer with 100 µm mesh size and discard the leftover egg-laying medium.
2. Prepare a 6-well plate as follows: fill the A1, A2, A3 and B3 wells with 10 mL PBS pH 7.4 and the B1 and B2 wells with 9 mL PBS. Then carefully add 1 mL sodium hypochlorite to B1 and B2. Observe the progress of steps 4.3 to 4.8 under the stereo microscope. CAUTION sodium hypochlorite is corrosive.
3. Insert the cell strainer into the A1 well (PBS pH 7.4) and wash the embryos for one minute under gentle agitation. Larger flour particles should detach from the embryos.
4. Transfer the cell strainer to the B1 well (sodium hypochlorite in PBS pH 7.4) and shake the plate vigorously for 30 s. The remaining flour particles should detach completely from the embryos.
5. Transfer the cell strainer to the A2 well (PBS pH 7.4) and wash the embryos for 1 min under gentle agitation.
6. For dechorionation, transfer the cell strainer to the B2 well (sodium hypochlorite in PBS pH 7.4). Shake the plate vigorously until the chorion ruptures and detaches from the embryos. The chorion looks like a thin, semi-transparent membrane with lacerated edges, while dechorionated embryos have a plain silhouette.
7. Transfer the cell strainer to the A3 well (PBS pH 7.4) and wash the embryos for 1 min under gentle agitation. If any chorion fragments are still attached to the embryos after the washing, repeat step 4.6.
8. Store the embryos in the B3 well (PBS pH 7.4). Storage for several hours does not harm the embryos, but keep in mind that development continues.
9. For non-homozygous transgenic lines, remove any non-fluorescent embryos if possible. For fixation and staining of WT or transgenic embryos, continue with step 5. For live imaging of transgenic embryos, continue with step 6.

5. Vitelline Membrane Permeabilization, Fixation, and Staining

NOTE: The vitelline membrane is the inner layer of the eggshell. It is impervious for fluorescent dyes and thus has to be chemically permeabilized before staining.

1. Fill a scintillation vial with fixation/permeabilization solution (1.5 mL 4% (w/v) paraformaldehyde in PBS pH 6.9 and 1.5 mL n-heptane). Transfer embryos with a paintbrush to the vials. Cover vials with aluminum foil to protect fluorophores from light and incubate for 30 min on a rocking platform at 50 rotations per minute. CAUTION paraformaldehyde is toxic and corrosive, n-heptane is flammable and toxic.
2. Remove fixation solution and treat embryos three times for 15 min in 3 mL 0.1% (v/v) Triton X-100 in PBS pH 7.4 and subsequently once for 15 min in 3 mL 1% (v/v) Triton X-100 in PBS pH 7.4 on a rocking platform at 50 rotations per minute. CAUTION Triton X-100 is corrosive.
3. Remove permeabilization solution and add 0.5 mL staining solution to the vial. Exemplary staining solutions are provided in Table 2. Stain embryos overnight at 4 °C on a rocking platform at 50 rotations per minute. For the dyes proposed in Table 2, washing is not necessary.

6. Preparation of Mounting Agarose

1. Add 1 g low-melt agarose to 100 mL PBS pH 7.4 and heat the mixture for 2-3 min in a microwave oven at 800 W. If small agarose particles are still present, repeat the heating process in 30 sec intervals until the particles dissolve completely. The mounting agarose does not need to be prepared freshly every time, the solidified agarose can be re-liquefied by heating as described above. This process can be repeated 5-6 times before too much water has evaporated.
2. Allow the agarose to cool down to approximately 40-45 °C, then fill two 1.5 mL reaction tubes with the heated agarose and keep those tubes at 35 °C in a thermoblock.

7. Embryo Mounting and Insertion into the Sample Chamber

1. Option 1: Agarose column (Figure 3A-C, first column; Table 3, second column)
   1. Fill a 1.0 mL syringe with distilled water and attach it to one of the glass capillary openings by using a small rubber hose. Fill the capillary halfway with distilled water.
   2. Pick an embryo with a paintbrush and place it on the inner side of the other glass capillary opening. Proceed quickly to the next step before the embryo dries out.
   3. Stick the capillary into the liquid agarose and slowly draw a few µL of liquid agarose alongside the embryo into the capillary. Then quickly increase the drawing force so that the embryo is dragged along with the agarose. In the end, a few µL of agarose should be above and below the embryo. Set the capillary aside for a few minutes and let the agarose solidify.
   4. Shorten the capillary with a diamond glass cutter to a length of about 5 mm. The remaining fragment should be completely filled with agarose and the embryo should be within the second quartile.
   5. Insert the steel cylinder with the pin into the sample chamber, and then stick the capillary fragment on top of the pin. The agarose column should protrude a few millimeters from the capillary fragment with the part that contains the embryo.
2. Option 2: Agarose hemisphere (Figure 3A-C, second column; Table 3, third column)
   1. Draw 200 µL of liquid agarose into a 1.0 mL syringe, then use it to fill the steel pipe from the top with agarose until a hemisphere forms at the bottom opening. The hemisphere diameter should be equal to the pipe diameter. Wait a few minutes until the agarose has solidified.
   2. Pick an embryo with a paintbrush and rearrange it on the tip of the brush so that the anterior end becomes accessible. Dip the agarose hemisphere into liquid agarose to cover it with a thin film.
   3. Place the embryo with its anterior end upright on the pole of the agarose hemisphere. Turn the steel pipe around and correct the position of the embryo with the brush. If necessary, stabilize the embryo by adding 2 µL agarose to the embryo/hemisphere border. Ideally, less than half of the embryo surface should be covered in agarose and the flanks should be as steep as possible.
   4. Insert the steel pipe with the hemisphere and embryo slowly into the sample chamber.
3. Option 3: Cobweb holder (Figure 3A-C, third column; Table 3, fourth column)
   1. Pick an embryo with a paintbrush and set it aside.
   2. Cover the slotted hole of the cobweb holder with 5-8 µL liquid agarose, then remove most of the agarose until only a thin agarose film remains.
   3. Place the embryo onto the agarose film and adjust its position. Carefully move the embryo to the x-axis center of the slotted hole, and align its elongated anterior-posterior axis with the elongated axis of the slotted hole.
   4. Insert the cobweb holder with the mounted embryo slowly into the sample chamber. Position the cobweb holder so that it does not interfere with the excitation and emission light, i.e. 45° relative to the x- and z-axis.

8. Light Sheet-based Fluorescence Microscopy

1. Decide along how many directions (and along which orientations) the embryo should be imaged. Four directions (along the orientations 0°, 90°, 180° and 270°) typically provide a good trade-off between coverage and energy load.
2. Set up the number of fluorescence channels. The major parameters for each channel are the laser line, the fluorescence filter, the laser power and the exposure time. For long-term live imaging, it is important that the integrated energy load does not exceed the tolerance of the embryo. An exposure time of 25-50 ms ensures fast imaging, while a laser power between 100 and 300 µW should not affect the viability of the embryo. For fixed samples, exposure time and laser power can be at least doubled.
3. For live imaging assays, configure the time lapse. The complete embryogenesis of Tribolium takes about seven days at room temperature (23 ± 1 °C) and a little less than three days at 32-35 °C⁶⁴. Define the temporal interval according to the morphogenetic events that should be observed. For short temporal intervals, consider lowering exposure time and laser power (see 8.2).
4. Position the embryo in transmission light mode along the x- and y-axes into the center of the field of view without exposing it to the laser. Rotate the embryo by 360° in 90° steps to examine the embryo for any damage that could have occurred during the mounting process.
5. Rotate the embryo appropriately around the y-axis to align the embryonic axes with the microscope axes, for example the lateral axis with the x-axis, i.e. the dorsoventral axis with the z-axis. When using the cobweb holder technique, alignment might not be possible.
6. In the fluorescence mode, define the z-stack for each direction. Add 25-50 µm as spatial buffer in front of and behind the embryo. Including the buffer, the typical z-stack covers around 350-400 µm for a Tribolium embryo. Also define the z-spacing, which should be four times the lateral spacing, i.e. the spacing along the x- and y-axes⁶⁶.
7. If two or more embryos are imaged in parallel, restart at 8.4 with the next embryo.
8. For long-term imaging, ensure that the sample chamber is filled with sufficient PBS pH 7.4 or other imaging buffer. Also cover the sample chamber opening.
9. Start the imaging process. For long-term live imaging, monitor the correct acquisition along all directions in all channels for all embryos. Occasionally check during the next days that the embryos are alive and in the appropriate position.

9. Embryo Retrieval and Quality Control

1. Once imaging has completed, remove the sample holder with the embryos from the sample chamber. Only embryos from live imaging assays have to be retrieved. Fixed and stained embryos can be discarded.
2. If a live imaging assay was performed, transfer the embryo to an object slide. For the agarose column technique, extract the embryos from the surrounding agarose with a scalpel, micro-knife or razor blade. For the agarose hemisphere technique, transfer the hemisphere with the flat side to the object slide. Embryo removal is not necessary. For the cobweb holder technique, gently dismount the embryos from the agarose film with a paintbrush. Incubate the object slide in a small glass dish in saturated humidity atmosphere under standard conditions until the larvae hatch.
3. After hatching, transfer the larvae to individual wells of a 24-well plate and add 500 mg of growth medium to each well. Incubate under standard conditions. Compare the total development time and morphology of the imaged larvae to non-imaged WT and transgenic larvae (see discussion). In assays that characterize WT development, no obvious morphological aberrations should ideally be noticeable.
4. Once the larvae develop to healthy adults, provide them with appropriate WT partners of the same background strain. After two weeks, check for progeny production. Also consider to check the fertility of the progeny by mating them inter pares. Only when the imaged embryos develop into fertile adults that produce fertile progeny, the imaging procedure can be considered non-invasive.

10. Image Data Processing

NOTE: For image data processing, the open-source image processing software ImageJ⁶⁷ or its derivate FIJI⁶⁸ are recommended. Both versions as well as comprehensive documentation are found at www.imagej.net. The standard file format for LSFM data is the tagged image file format (TIFF). The intrinsic container option allows the storage of image stacks such as z-stacks or time series of z maximum projections within a single file that can be opened in ImageJ or FIJI via drag-and-drop.

1. If necessary, rotate the z-stacks to align the anterior-posterior axis of the embryo with the y-axis (Image → Transform → Rotate). Please note that the rotation parameters have to be set individually for each direction, but not for the time lapse and the fluorescence channels.
2. Crop the z-stacks along the x-, y- and z-axes so that only minimal background (20 - 40 pixels along the x- and y-axes, 5 - 10 planes along the z-axis) remains (for the x- and y-axes, use the Rectangular Selection Tool, then Image → Crop; for the z-axis, use Image → Stacks → Tools → Slice Keeper). Please note that the cropping parameters have to be set individually for each direction, but not for the time points and the fluorescence channels.
3. Calculate the z maximum projection for each z-stack (Image → Stacks → Z Project, choose Max Intensity). Intensity projection calculations are data simplification procedures that remove one spatial dimension.
4. For long-term live imaging data, consider using an ImageJ or FIJI script that performs processing steps 10.1 to 10.3 for all directions, all time points and all fluorescence channels⁶⁵. Ideally, concatenate all z maximum projections per channel and direction and save them as a time series in one TIFF container. Alternatively, the BigDataViewer⁶⁹ plug-in for FIJI can be used to navigate through Terabyte-large datasets.
5. Depending on the transgenic line and imaging modalities, dynamic intensity adjustments of the time stacks might be desirable due to photo-bleaching and/or fluorophore expression fluctuations. Either the ImageJ- or FIJI-intrinsic Bleach Correction function (Image → Adjust → Bleach Correction, choose Histogram Matching) or dedicated software⁶⁵ are suggested.
6. If the embryo was recorded and/or along multiple directions and in multiple fluorescence channels, horizontal combination of the directions (Image → Stacks → Tools → Combine) and channel merge (Image → Color Merge → Channels) are recommended.
7. Registration and fusion of z-stacks acquired along multiple directions⁷⁰, eventually in combination with deconvolution⁷¹, allows the calculation of superior three-dimensional images of uniform quality and isotropic resolution. Both plug-ins are open source and pre-installed in FIJI, but they require the presence of landmarks (fluorescent microspheres, e.g. beads) around the specimen.
8. Open-source software frameworks for large-scale quantitative data extraction and analysis are also available, for example for segmentation and tracking of cell nuclei⁷² or cell shape recognition⁷³.

Results

This protocol describes an experimental framework for fluorescence imaging of living or fixed and stained Tribolium embryos with LSFM. Due to the low levels of photo-bleaching and photo-toxicity, a direct consequence of its optical sectioning capability, LSFM is particularly well suited for long-term live imaging.

The novel AGOC{ATub'H2B-mEmerald} #1 transgenic line expresses a histone2B-mEmerald fusion protein under co...

Discussion

Quality control

In live imaging assays, the preparation and recording procedure must be non-invasive, i.e. neither the mechanical and chemical handling (collection, dechorionation, mounting onto the sample holder) nor the integrated energy load during the observation should affect the viability of the specimen. For studies that characterize WT development, it is recommended to only use data from experiments in which the embryo survives the recording process, is retrie...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
full grain wheat flour	Demeter e.V.	1.13E+08	US: whole wheat flour, UK: whole meal flour
405 fine wheat flour	Demeter e.V.	1.13E+08	US: pastry flour, UK: soft flour
inactive dry yeast	Flystuff / Genesee Scientific	62-106
Phosphate-buffered Saine (PBS), pH 7.4	Thermo Fisher Scientific	10010-023
sodium hypochlorite, ~12% active Cl	Sigma Aldrich	425044-250ML	Caution: sodium hypochlorite is corrosive
low-melting agarose	Carl Roth	6351.2
6-well plate	Orange Scientific	4430500
24-well plate	Orange Scientific	4430300
glass capillaries, internal Ø 0.46 mm	Brand GmbH + Co KG	7087 09
SYTOX Green	Thermo Fisher Scientific	57020	Staining solution preparation is explained in Table 2
YOYO-1 Iodide	Thermo Fisher Scientific	Y3601	Staining solution preparation is explained in Table 2
BOBO-3 Iodide	Thermo Fisher Scientific	B3586	Staining solution preparation is explained in Table 2
Alexa Fluor 488 Phalloidin	Thermo Fisher Scientific	A12379	Staining solution preparation is explained in Table 2
Alexa Fluor 546 Phalloidin	Thermo Fisher Scientific	A22283	Staining solution preparation is explained in Table 2
sieve, 800 µm mesh size	VWR International	200.025.222-051
sieve, 710 µm mesh size	VWR International	200.025.222-050	for growth medium preparation (step 1.1)
sieve, 300 µm mesh size	VWR International	200.025.222-040
sieve, 250 µm mesh size	VWR International	200.025.222-038	for egg laying medium preparation (step 1.2)
glass dish, Ø 100 mm × 20 mm	Sigma Aldrich	CLS70165102
cell strainer, 100 µm mesh size	BD Biosciences	352360
paint brush, head Ø 2 mm	VWR International	149-2121
syringe, 1.0 mL	B. Braun Medical AG	9166017V
scintillation vials	Sigma Aldrich	M1152-1000EA
paraformaldehyde (PFA)	Sigma Aldrich	158127	Caution: paraformaldehyde is toxic and corrosive
n-heptane ≥99%	Carl Roth	8654.1	Caution: n-heptane is flammable and toxic
Triton X-100	Sigma Aldrich	X100-100ML	Caution: Trition X-100 is corrosive