- Appropriate medium for the specimen of choice, e.g. E3 for zebrafish
- Low gelling temperature agarose, e.g. Sigma Aldrich A9414
- Methyl cellulose
Fluorescent beads (optional)
Fluorescent beads (or microspheres) are great samples for evaluating microscope and light sheet alignment, and they can also serve as fiducial markers for multi-view registration. Multi-color beads are labeled with four fluorescent dyes and visible in all channels. They are handy as universal test samples and to measure chromatic aberration. Single-color beads are only visible in some channels and a great add-on for multi-view, especially when placing them in an otherwise unoccupied channel. They also tend to be brighter when compared to multi-color beads, which can be an advantage during light sheet alignment. Beads come in different sizes. We found 0.5 µm diameter to be a good choice for a magnification of 10x-20x.
- Tetraspeck fluorescent beads, multi-color, 0.5 µm: Thermo Fisher T7281
- Estapor fluorescent beads, single-color, 0.5 µm
- EX 440, 458, 468 / EM 480, 510: Sigma 39555001
- EX 470, 490 / EM 525, 560: Sigma 80380016
- EX 515, 555 / EM 568, 610: Sigma 80380495
- EX 420, 515, 550 / EM 656, 720: Sigma 80380496
capillaries & plungers
Glass capillaries shape the agarose cylinder and should be considered disposables. Matching plungers are required to take up samples in liquid agarose into the glass capillaries and can typically be reused. Both are spare parts for the Brand Transferpettor pipette, and the Brand website lists details and order numbers for capillaries and plungers. Examples for matching combinations of capillaries and plungers are given here.
|Color code||Diameter||Example applications||Capillary||Plunger|
|white||tbd||Worm embryos||Sigma Z328464||Brand 701928|
|orange||0.6 / 1.2 mm||Fruit fly embryos, adult worms||Sigma Z328472||Brand 701930|
|black||1.0 / 1.5 mm||Zebrafish embryos/larvae (dechorionated)||Sigma Z328480||Brand 701932|
|green||1.5 / 2.0 mm||Zebrafish embryos in chorion||Sigma Z328502||Brand 701934|
|blue||tbd||tbd||Sigma Z328510||Brand 701936|
FEP is a transparent polymer with a refractive index similar to water, which makes it a great material for imaging live samples. Several manufacturers sell standard and custom FEP tubing by the meter.
The right tube diameter is determined by the specimen. For zebrafish 0-5 dpf, we use either 0.8 x 1.6 mm (thick walls for increased rigidity, from BOLA or Pro Liquid) or 0.8 x 1.2 mm (thin walls for imaging, from Pro Liquid). Good tube sizes for bigger samples are 1.6 x 2.4 mm (from Pro Liquid) and 2 x 3 mm (from BGB).
A 1 ml plastic syringe with a luer slip tip (Sigma Z683531 or CareTouch CTSLL1 ) and an attached injection needle is a convenient tool to take up samples into an FEP tube. Syringe needles come in different sizes with the outer diameter expressed in Gauge. This needs to match the tube’s inner diameter. Alternatively, especially for wider tubes, use a standard pipette and a matching plastic tip.
|Diameter||Wall thickness||Example application||Source||Needle|
|0.8 / 1.2 mm||200 µm||Zebrafish embryos/larvae (dechorionated)||Pro Liquid||BD #305165 Gauge 21|
AIR-TITE #89500-304 Gauge 21
Sigma Z192481 Gauge 21
|0.8 / 1.6 mm||400 µm||Zebrafish embryos/larvae (dechorionated)||BOLA, Pro Liquid||BD #305165 Gauge 21|
AIR-TITE #89500-304 Gauge 21
Sigma Z192511 Gauge 20
|1.6 / 2.4 mm||400 µm||tbd||Pro Liquid|| Grainger Gauge 16|
BD #305197 Gauge 16
|2 / 3 mm||500 µm||tbd||BGB||Grainger Gauge 14|
Sheets made of FEP can be a great starting point for custom sample chambers, bags, tubes and covers. BOLA offers FEP sheets of different thickness.
FEP tube straightening (optional)
FEP tubes typically arrive rolled-up, resulting in a more or less pronounced bend. This will result in severe sample displacements when rotating the tube in L/T/X configurations. Straightening tubes might help to mount the sample and align the image data. However, the tubes have to be straightened and cut to length before cleaning, making the process of cleaning much more elaborate and time-consuming.
The idea is to straighten the tube by heating it to 180 °C and slowly cooling it down. The process happens either in stainless steel tubing or in glass tubing. Both have to have the right inner diameter to fit FEP tubes in. We now use two steel tubings of the length of around 50 cm. Inner diameter is either 2.4 or 1.6mm.
- Start procedure the afternoon before intended FEP tube cleaning
- Cut FEP tubes slightly longer than steel tubing
- If possible, fill smaller FEP tube in larger FEP tubes (example: put 1.2 x 0.8 FEP tube in 2.4 x 1.2 FEP tube, and both together in steel tube)
- Place FEP tubes in steel tubing, with a little end of FEP tube sticking out
- Heat these tubes to 180 °C for 2 hours, e.g. in large autoclaves from cleaning service
- After 2 hours, take out tubes and let cool down at room temperature for at least 5 hours
FEP tube cleaning
FEP tube are typically dusty/dirty upon arrival and require cleaning. Cleaning long pieces of tube before cutting them to length will make that process easier. To avoid contamination, use gloves and/or forceps to handle clean tubes.
Flush FEP tubes twice with 1 M NaOH (Merck) using syringe, 0.45 μm PVDF filter (see below) and matching syringe needle. Place the flushed tubes in Falcon tubes with 0.5 M NaOH and ultrasonicate them for 10 minutes. Transfer the tubes into a small basin and flush them with double-distilled H2O and with 70% ethanol. Place the tubes in fresh Falcon tubes with 70% ethanol and ultrasonicate for 10 minutes. Finally, flush the tubes with double-distilled water, cut them to length and store them in Falcon tubes with double-distilled water.
All solutions should be degassed and filtered using a syringe filter (Millex-HV PVDF 0.45 μm, #SLHV033RS) before use.
Mounting for L/T/X configuration
There are numerous techniques of mounting samples for the L, T and X configurations. Here, we describe three distinct strategies: First, mounting in a free-hanging agarose cylinder (left). Second, mounting in an FEP tube, either in solid agarose (center) or in low-concentration agarose (right).
Mounting in free-hanging agarose cylinder
- Melt 1.5% agar, keep it at 37ºC.
- Treat samples with tricaine.
- Put the plunger in the glass capillary such that the white end barely comes out of the capillary.
- Put the sample in agar, with as little water as possible.
- Take up about 3cm (1”) of agar in the capillary by pulling the plunger.
- Take up the fish, head down. Keep the fish close to the end of the tube.
- Let agar set 1-2 minutes and transfer into a 12ml falcon with tricaine E3. Be careful not to move the plunger or the fish might be released.
- Once you are at the microscope and ready to image, push the plunger down such that the fish is just outside the capillary.
Mounting in FEP tube
- Prepare a petri dish coated with a thin layer (1-2mm) of 1.5-5% agar.
- Melt 0.1% agar, keep it at 37ºC.
- Treat samples with tricaine.
- Cut the clean and straightened FEP tube to the desired length with a razor blade.
- Mount the FEP tube on the blunt end needle.
- Place the sample in agar with as little extra medium as possible.
- Fill the FEP tube with agar first, then take up the fish, head down. Keep the fish close to the end of the tube.
- Carefully stick the FEP tube into the agar coated dish at a 90º angle. Avoid bubbles between plug and agar. Rotate the tube and take it out.
- Check by eye or stereoscope that there is an agar plug at the end of your FEP tube. If there is no plug, repeat the previous step.
- Cut the FEP tube at the edge of the blunt end needle.
- Transfer FEP tube into an Eppendorf tube with tricaine E3 until ready to image.
You can also take up the fish in 1.2-1.5% agar and not use the plug. The FEP tube will provide additional stability. However, keeping the developing fish in solid agar will impair its development. We recommend using 0.1% agar and a plug for imaging for more than one hour.
- Kaufmann, Mickoleit, Weber & Huisken, Development 2012: Multilayer mounting enables long-term imaging of zebrafish development in a light sheet microscope
- Weber, Mickoleit & Huisken, Jove 2013: Multilayer Mounting for Long-term Light Sheet Microscopy of Zebrafish
- Weber, Mickoleit & Huisken, Methods in Cell Biology 2014: Light sheet microscopy