This week I’ve been diving in a little more into doing some actual research myself. Nothing breakthrough mind you, just some simple experiment to sort of understand the world around and inside of me a little better. My partner and I are looking into how the optic nerve develops inside of zebra fish and how its development may be affected by developing the fish in total darkness. We are trying to stain 1-2 day old zebra fish with Dye-I by simply poking the dye into the retina with any sort of small sharp object we can find. We’ll then separate the groups of stained fish into those that develop in a small flask with normal exposure to light (about 10-15 hours a day of UV light) and those that develop in a tin-foil-wrapped flask with no UV exposure in which feeding will be done under red light.
Every few days, I hope to take some of the fish out and look at how their brains are developing using a fluorescent microscope that allows me to see how the dye is traveling. With any luck, I’ll see a decently clear pattern of paths of nerves from the retina to the lateral geniculate. I’m not sure where it leads from there as the zebra fish has mainly the midbrain rather than our large forebrain with a thalamus and cerebral cortex. More research on my fish is definitely needed before I can do any real analyzing of the staining technique, but my real problem right now is just getting some fish stained!
Using a pin head dipped in dye, I had been trying to poke the retinas of these fish. Once the retina ruptures, almost any amount of dye should stain the retina and lead to further staining of the optic nerve as development continues (from what I’ve read, usually the rods and cones develop about 4 days at the earliest). I ran into many problems with this of course and have been fishing (oh so hilarious, I know) for new solutions ever since.
The first problem I had is that the head of the pin is about the size of the fish’s eye itself, so trying to rupture the eye with it is like trying to shove a baseball bat through my own. Of course actually rupturing the retina with this gigantic tool will only happen if my mounted fish would stop moving every time I get close to it, which is the second problem. So far, I’ve created new tools by heating TLC spotters over a flame in the organic chemistry lab and then pulling it in half so that the glass is pulled into a tiny pipette with a head just smaller than the fish’s retina. This has been successful after coupling it with a technique for knocking out my fish to keep them still. I burst a small bubble of anesthetic (0.5 MESAB) over the water that I’m using to mount my fish before injecting the fish in augar onto the slide. This does a great job in keeping the fish still so I can do my dirty work of staining its eye, but sometimes the concentration of anesthetic is too strong and the fish will die. All in all, with much more research and some patience this may turn out to be a fun little project. If you have any ideas as to where I should be looking for research and anything else I could be doing with this experiment, let me know.
~Bright Lights
I don’t suppose you have access to a microcapillary pipet puller, do you? Ultra fine point will help injection into the eye. Or see if you can get surgical needles of the highest gauge.
ultra-thin needles made out of capillary tubes are definitely the way to go here. Check around to see if there’s a needle puller in your facility. Sutter Instrument Co. makes the most popular one of these, I think.
If the ~24hpf fish is embedded in agar, you shouldn’t really need to mesab it for immobilization. A more dilute mesab solution should be sufficient for anaesthetic purposes. Check the Zebrafish Book (Westerfield) for recipes, etc.
Finally, your developmental time line is a little off. Retinal neurons start differentiating at 27hpf, beginning with the ganglion cells (the ones whose axons lead to the optic nerve). Photoreceptors ae a couple of orders removed from the optic nerve, being the sensory neurons at the top of the pathway. They start differentiating at ~46hpf. Synaptogenesis comes on the heels of this, as the neurons in questions are specified, and the whole retina is wired and functional at ~70 hpf.
I have a question, rather than any helpful ideas about staining. If you are rupturing the retina of the fish to introduce the dye, how can you then track the development of their brains with respect to light… if they can’t see?
I’m a biology n00b, so I’m probably missing something old hats know automatically.
Incidentally, I found this article to be very engaging. Your writing style is quite entertaining, and made me think a lot about what you were trying to accomplish, made me try to visualize it.
It’s DiI, not dye-I.
Just wanted to chime in and say this was interesting to me too. Sometimes the simple, practical, engineering stuff turns out to be key to the big breakthroughs, too – your approach to problem-solving should serve you well.
I wish you luck in all your fish-impaling endeavors!
Pulling glass tubes for TLC is always fun, even when your as burn prone as I am. Cheap, quick and you get to play with a Bunsen.
The suggestion of using a pipette puller to make micro pipettes is a good one, but if you don’t have access to a pipette puller, micro pipettes can be bought. They are used in IVF labs to inject sperm into eggs for a technique know as ICSI and a google search should lead you to venders. You can buy them in a range of sizes from 5 um to 30 um. They are expensive, though, and if funds are tight and you don’t have access to a puller, a pipete with a tip of around 100 um can be pulled by hand starting with capillary tubes. Take a pasteur pipette and bend the end over a bunsen burner to a 90 degree angle. Attach it to your gas source with tygon tubing and use a clamp to close the tubing most to the way. Turn on the gas, light it and use the clamp to create a very small flame. Pull your capillary tubes over the flame to create a very small bore, let it cool, and snap it. You can bevel the end by gently stroking it on jeweler’s cloth or a very fine knife shapening stone. You can check the tip under a dissecting scope and use a micrometer to measure the diameter. You could use the resulting micro pipette to rough up the retina. I’m confused about the rupturing but I don’t know much about fish eyes, only mice. Do fish have a lens and a perivitellin space? I’m thinking that if they do rupturing some of the retinal vessels then injecting your dye into the perivitelline space might be enough to allow uptake.
Yikes! That sounds gruesome.
This is no doubt a pretty dumb suggestion, but could you partially immerse a starting pistol in the fish tank and fire it and see if the concussion stuns the fish long enough for you to perform the procedure? On the downside you might give other people in the lab heart attacks, on the plus side it might impress Charles Heston.
DiI is lipophilic, and will transport in the neuronal membranes of paraformaldehyde-fixed tissues. If your question has to do with retinal connections during development, why not euthanize (with an overdose of anesthesia or whatever) zebrafish at different developmental stages, fix them in 4% paraformaldehyde, poke diI crystals into the neural retina, and store the zebrafish in a dark drawer for a few days to let the diI work its mojo? This technique works great for mouse and chick embryos, and for organ cultures; don’t see why it wouldn’t for zebrafish. The stiffness from the paraformaldehyde fixation should make diI application with your crude methods somewhat easier. I wouldn’t attempt diI-labeling in a live embryo without a picospritzer.
Am I the only reader who feels sorry for the fish?
No LC, you are not. The enthusiastic response to this makes me a little uneasy.
You may be interested in this new paper.
And perhaps this and this.