Hello Kunal,

we used to image some zebrafish embryos on a inverted confocal microscope: in order to do that we just used a chambered coverslip, which was a standard microscopy coverslip, with a silicone layer, 1 mm thick on its surface, dividing it in multiple separated chambers (it looked like the ones in the picture here: http://www.sigmaaldrich.com/labware/labware-products.html?TablePage=9578334).

With one of these coverslips, we filled a chaber of low melting point agarose gel including the zebrafish, removed all the excess agarose on top, and sealed it with a standard coverslip to prevent the agarose from drying out. Basically we obtained a sandwich of two coverslips, 1mm apart from each other, with the silicone layer around the zebrafish to prevent "crushing" it.

It worked quite nicely, and embryos survived in there for several hours.

I hope this helps!

The chambered coverslip is a good option. Many of the silicone ones are peel-and-stick, which makes them easy to use. They hold liquids, but their size doesn't hold the larvae in a specific orientation.

We also have good luck using an agarose gel (1-1.5%) with V-shaped channels cut into it. This allows you to fill the channel with your anesthetic (MS-222 or cold water), keep the larva moist, and orient it to your needs. The point of the V (oriented downward) keeps a tight hold on the larvae to keep it from moving/rolling. Because you can fill the channels with fluid, you can add/remove individuals using a transfer pipette, preventing damage. We also use a piece of fine fishing line to make small adjustments to the larvae while it is in the channel; this will also help prevent damage.

Descriptions of how to make these gels are in the zebrafish microinjection literature and the 3 resources I've added below. Many of these methods use molds to create the channels. You can create your own molds - see Sabaliauskas 2006 or try something similar to an electrophoresis comb.  Alternatively, if you are careful, you can make the channels using a scalpel or single-edged razor blade -- I suggest doing this cutting under a microscope. Once made, the gel can be reused -- we keep ours submersed in buffer solution at 4C between uses. 

http://zfin.org/zf_info/zfbook/chapt5/5.1.html

Book The Zebrafish Book. A Guide for The Laboratory Use of Zebraf...

Thank you Paolo and Melissa! Paolo, do you use a specific one? could you share a product or catalog number?

Melissa, with regards to the channels, are they cut throughout the thickness of the gel to allow for imaging on an inverted microscope?

We make the gel in a basic plastic Petri dish. Fill dish ~1/2 - 3/4 full of molten agarose and let it solidify at 4C on a flat, even surface. When solid, remove the plate and make your cuts. The channels only need to be 1-3 mm deep and ~3 mm wide (top of the V). The channel should be such that the larva stays put and immersed where you can see it. If the channel is too deep and/or the top of the V too narrow, then you risk the larvae getting stuck where you can't move it or get it back out, and it could be difficult to image. 

Since my last reply, I thought of another technique that we have used with some success. When the gel has set up but is not yet solidified, place 5-6 capillary tubes in lines on top of the gel. They should sink in but not be covered. When the gel has solidified, use forceps to remove them while being careful not to tear the gel. This will create round bottom channels. They aren't quite as good as the V channels with keeping fish orientation, but they are much faster and easier to make. I even know some people that have fashioned their own molds/stamps using them which have an additional depth to hold more liquid for longer imaging sessions (see attached pictures).

https://www.thermofisher.com/us/en/home/references/molecular-probes-the-handbook/tables/coverwell-chamber-gaskets.html

Hello Kunal,

unfortunately, it was work i was doing in my former group, and i never ordered the coverslips, we just had them around for another project. something from the page on sigma aldritch i linked should be good, just make sure the size of the well is big enough to accomodate the zebrafish.

Also, the capillary made channels Melissa suggested should be easy to make, and could work nicely on an inverted microscope, if you make the agarose layer thin enough.

As a last solution, our first tests were made with square section borosilicate capillaries from CM scientific (like these: http://www.cmscientific.com/proddetail.php?prod=8100-length), you can get quite wide ones with thin walls, and since the sides are flat they work really well with microscopes, you just fix the whole capillary over your objective, without coverslip or anything.

We attached a syringe to one end of the capillary with a flexible rubber tube, and sucked in the zebrafish in low melting agarose. We moved out of that solution because the zebrafish did not last very long in them (1-2 hours), but you can give them a try.

Hi

I usually do live imaging using confocal microscope. Hope this method suit your situation.

In our case, we use glass bottom dish for live imaging. Basically it is small dish with cover slip as the bottom. So, what you need to do is apply some melted low melting agarose on it and mount your sample in it. Once the gel is solidify (usually within 20 min), put feeding water on top of it (1-2mL) and you can use it for imaging. In our case, the water will not dry out for at least 5-6 hours.

I have been using the following protocol adopted from the mentioned article.

I anaesthetize larvae using 200mg/L tricaine and make a 0.08% w/v low melt agarose gel in the same concentration of tricaine. I usually add approximately 130 uL of low melt agarose in each well (48 well plate) and let it cool down before adding the larvae. The anaesthetized larvae can then easily be positioned and they remain stable in the gel for 4-5 hours.

The advantage is that you can plate multiple embryos (critical for my study). I don't know if you're imaging single larvae.

Article Developing a Novel Embryo-Larval Zebrafish Xenograft Assay t...

Hi, I think one of the adaptations when using LMP agarose is to mount the anestetized larvae in the agarose (0.5-1%) and after solidified cover with E3 medium with tricaine...maybe it will avoid it to dry. Best wishes.