As I wrote about before, my semester lab project for neurobiology has to do with regeneration. The idea is to damage the spinal cord and observe wonderful regeneration. This proposal was based on some articles I read about regeneration of zebrafish hearts, fins, tails, etc. Unfortunately I haven’t had much luck so far.
Last week, armed with an exacto knife, I performed my first round of spinal cord butchering on fifteen zebrafish that were only a few days old. The zebrafish are captured with a glass pipet and then immobilized using auger that’s just warm enough to be in liquid phase. They are then mounted on a slide under the stereoscope. I quickly discovered many faults in my methods, one of which being that I captured the fish in too many drops of water so when the auger was added in a test tube, the fish weren’t completely immobilized. I would then carefully approach a fish with the exacto knife on the slide under the scope and it would turn into a bucking bronco. Eventually I perfected the art of capturing the few day old zebrafish with the pipet and putting them in test tubes in only one drop of water, thus partially solving the immobilization issue.
The second problem I encountered is that there is nothing exact about an exacto knife under a stereoscope. Accomplishing this spinal cord severing is much like peeling an orange with a baseball bat in that it’s extremely difficult without making a mess. Even with the fish immobilized the tail doesn’t really stay put when pressure is applied with the seemingly crowbar sized exacto knife. The key to this dilemma, although I have yet to master it, is probably making the layer of auger on the slide as thin as humanly possible so that there isn’t as much room for movement.
All the zebrafish from my first attempt died. Two of them were alive for a day or so but barely. I did another round of zebrafish butchering with fifteen more fish yesterday (yes I enjoy spending my Sunday afternoon in the neurolab) and from what I could tell, all but two of them died immediately. I’ll have to see if the elite two are still swimming around today but if not, I’ll try yet another round of fifteen and see how it goes. If anyone has any ideas for instruments or methods that could improve my success, feel free to insert a comment.
Don’t push down with the knife. Use it to slice. Set the blade where you want to make the incision and pull it sideways in a sawing motion. Your cuts will be cleaner and there will be less damage to the fish from smashing.
How about using a little tricaine to knock them out?
http://en.wikipedia.org/wiki/Tricaine_methanesulfonate
It would make it much less stressful for both you and the fish. It’s available from Sigma, but someone at your university likely has some.
By auger you mean agar? For a cutting instrument, some really fine cuts can be made with glass blades harvested from aged window glass recovered from old buildings – where the glass has deformed from gravity over many decades. It can be broken to create thin, curved blades that are very sharp. The blades can be fixed with glue, etc. to a handle. this is just one suggestion for getting a sharp blade costing more in effort than in money.
A properly anesthetized fish will not move. Try MS-222 (tricaine), recipe in the Zfish book (see Zfin.org), or a similar anesthetic.
I suggest pouring some agar into your petri dishes and letting those solidify. Then, capture your anesthetized fish, and lay it onto the agar. Working quickly, draw off the water, and anchor the fish by placing a drop of your barely-liquid agar over the posterior half of the fish. Agar will stick to agar, and the fish will be anchored. Leave the head/gills uncovered so that a drop of MS-222 water can be placed there.
With the fish so anchored, and working under a dissecting ‘scope, a sharpened tungsten needle or an insect pin can be used to perform the spinal surgery. An exacto knife is a poor tool for this, as you are learning. Both needles or pins can be fused into pulled-glass pipettes with as little as an alcohol flame. And they make great zfish surgery tools!
After surgery, flood the dish with clean fish water, and carefully cut away the agar. The fish will float free, and can be moved to a warm dish with clean fish water for recovery. In my experience, a properly anesthetized fish will recover in less than 2-3 minutes (though your particular surgery could have a significant effect, of course).
I am happy to answer further technique questions if you have them. You may email me at my username @ gmail DOT com. Good luck!
Not to avoid your main question, but I did see a rather cool little speech about 2 weeks ago by a woman from Harvard who had induced regeneration in a tadpole past the normal tail regeneration stage, by using a technique she had used with playhelminthes before time, by introducing gap junction proteins (From fungi if I remember) into the organism that could not be shut-down by the normal innate processes of development. Basically put, as she said, “Mary Shelly had it right all along…” and it is an electric field that helps establish regeneration, and not some inducing protein as most believe…
Similar to the ‘old window glass’ suggestion, you might try using flaked obsidian? I’ve heard that it’s sharper than surgical steel with a narrower edge. I can’t vouch for that but it IS wicked sharp and readily available.
Alternately, maybe a very fine needle would work better? I’m not as conversant with fish biology as you obviously are, but would a puncture work for what you’re doing as oppposed to an incision?
Off-topic, but it hasn’t deformed. Glass doesn’t do that. Old glass windows simply never were even; the technique to get them even is very recent. Sometimes they are thicker at the top than at the bottom…
Off-topic, but it hasn’t deformed. Glass doesn’t do that. Old glass windows simply never were even; the technique to get them even is very recent. Sometimes they are thicker at the top than at the bottom…
The idea is to cut the spinal cord in two and see if it heals, so…
The idea is to cut the spinal cord in two and see if it heals, so…
Ctenotrish beat me to it. I was also going to suggest heating some glass (perhaps from a small test tube or pipette) and stretching it till it’s fine as hair. If you want, you can also compress it at the edge to make it flat. You may not get the cleanest cut, but it will allow you to puncture through, at the very least sever the spinal cord.
Another suggestion would be to use tweezers to break off the sharp edge at the point of the exacto-knife and then tape the tweezers along the middle to continuously grip the flaked metal edge.
You’re paying some good money to attend a university, and when you need technical help with a hands-on aspect of your project, you…ask the blogotube hive mind? Why not ask whatever professor is going to end up evaluating your project?
For larger cuts you should use some Lumsden bio-scissors. They were my best friends come dissection time when I was in school.
For finer cutting in soft tissue, I would recommend using very fine tungsten needles.
And as Pete asked: why not get your prof to help you with this?
Who are these idiots who think professors know what to do in the lab? Ask a technician instead!
The auger/agar error reminds me of one of my PhD supervisor’s classics. He was pouring a gel to do some RFLPs or RAPDs, and turned round to the technician and asked “For the agarose, do I use agar agar or bacto-agar?”.
Bob
Ah, this is a great showcase of how the web can be uniquely valuable. It is important to get to working methods fast, there are such things as misplaced patience. :-
I don’t see it mentioned, but FWIW, reference samples, including some that are diverted at key points in a process, is a tool to validate what your experiment did (and what your methods did (this time)).
Ah, this is a great showcase of how the web can be uniquely valuable. It is important to get to working methods fast, there are such things as misplaced patience. :-
I don’t see it mentioned, but FWIW, reference samples, including some that are diverted at key points in a process, is a tool to validate what your experiment did (and what your methods did (this time)).
@David:
I followed that, just wasn’t sure if a needle puncture would be sufficiently large to sever the cord. (I’ve only got a very vague idea how big these fish are)
My wife uses the Zebrafish as a model. So I called her and this is what she says:
Use adult zebrafish. Use “Mesab” to anesthetize the fish. Go to zfin.org and the protocols are there.
She also says to use a laser, forceps or scissors, razor blade, depending on the level of precision you need.
And, she, once again, says you should use ADULTS for regeneration as that’s regeneration, not a development process. She also says she believes there is work being done in this area/model and you might find some literature on PubMed, so search it.
Re: #10
Judging by the answers so far, Blue came to exactly the right place.
Of course, one of the answers was less than useless…
— CV
you would probably need IRB permission for this though.
A low tech approach I’ve found that works well is a old straight razor blade (I guess you could dissect a safety razor cartridge) cut on the bias and fixed with forceps. These blades tend to give clean cuts on all but the most heavily fixed tissue and make acceptable cheap alternatives to vibratome/microtome blades if you don’t have access to them (although you can pick up a pack of 20 microtome blades for less than $10 these days from most biotech supply firms).
Perhaps we could help you more if you give us a little more information on exactly what you’re doing – stages, assays etc ect.
it hasn’t deformed. Glass doesn’t do that.
This surprised me, but http://en.wikipedia.org/wiki/Glass#Behaviour_of_antique_glass supports it.
Sometimes they are thicker at the top than at the bottom…
Why not the other way around? Ah … “When actually installed in a window frame, the glass would be placed thicker side down for the sake of stability and visual sparkle.[10] Occasionally such glass has been found thinner side down or on either side of the windows edge, as would be caused by carelessness at the time of installation.”
Considering how much I’ve cursed trying to cut crystals and getting them to do my bidding on the slide, I don’t envy you your work with fish.
As they say: never work with children or animals – you seem to be doing both in one.
I assumed IRB permission was already granted.
And the razor blade is my wife’s favorite. Cheap. Low-tech. Cuts well and clean.
Agree with most of what’s said here. You can’t use adults without IRB, IACUC, training, etc.
What exactly are you hoping to regenerate? If you just want to lesion the spine and see if the axons of spinal neurons regrow, how will you tell? If you are cutting off the tail and with it some spinal cord… it’ll heal, but it won’t grow back.
If it’s the former, it is regeneration, as long as the same cells whose axons you sever are sending out new ones… hard to be sure, though, since there is still neurogenesis and new axons. And its true that the embryo is a much more permissive environment for axon growth than the adult (larvae have very little myelin, for example).
And yes, a razor blade on larval fish (the spinal cord is around 0.1mm across, if that) is like trying to cut bread with a brick.
Electolytically sharpened tungsten needles are the way to go. Look online, start with the thinnest wire you can find and sharpen with a low voltage power supply and 1M NaOH. I think a 9V battery works too.
You might try raising them in sterile Ringers instead of normal fish water after surgery, and adding some antibiotic.
Oh, btw, PZ posted on fish anesthetic on his old site: http://pharyngula.org/index/weblog/comments/how_to_euthanize_a_fish/
Using a lower dose, and/or for a shorter time, is what gets you the difference between recovery from surgery, and euthanized. Note especially all the different names for the same ‘cocktail’ . . . I had forgotten that MS-222 is also commonly know as mesab until I saw one of the comments above.
I’m sorry, I came across this just now and can’t refrain from making a comment. This is so horrible, it makes me really sad to see how low you value a life. Well, I guess it’s in the name of science and so on, but there is so much science to be done which doesn’t involve tortureing and killing!
Good luck with your project, though. I really hope you’ll get something usefull out of it.
The tail can move again?
The tail can move again?
My only experience with small animals was trying to dissect a preserved Amphioxus (lancelet). I soon abandoned the scalpel for a dissecting pin.
>The tail can move again?
that doesn’t tell you if there was axonal regeneration or new neurons. as i said before, there is still a lot of neurogenesis at this stage.
Ask the lab manager! Do not interrupt the PI unless the lab is on fire. If you only want to sever the spine, use a pulled Pasteur pipet, it will do the job, or the Lumsden bio-scissors will work too.
And to all the houty touty’s out there who said don’t ask blogdom, you are wrong!!! (3 !!! must mean it’s important).
This is why Al Gore invented the internet! (one more time; ask the lab manager/tech and not the PI).
Use a tungsten filament.
It’s odd that half of you are giving very good and helpful suggestions and the other half are chiding PZ for asking a question where there’s no way he’ll get very good and helpful suggestions.
Why is it that you don’t need IRB approval for “spinal cord butchering on fifteen zebrafish that were only a few days old.” and yet you would for full grown fish. That seems weird and kind of wrong. Are there somehow less ethical issues if the fish are young?
I’d make sure you don’t need approval.
PZ for asking a question
Uh, you think PZ wrote this article?
Ah, yes. I dissect a certain piece of E11.5 mouse brains, and I’ve likened getting the brain out at that stage (even using very fine forceps) to separating sheets of wet tissue paper with two pairs of pliers.
I agree with the suggestion of a tungsten needle for the zebrafish, although I think a pair of fine forceps could probably do the job pretty well.
Other people have nominated precise tools, but I’d like to put in another nomination for glass. Really fine glass bits are evidently sufficient to dissect certain insects, according to an old friend.
Granted, said old friend was working with ad hoc tools decades ago, so make of it what you will.
“there is nothing exact about an exacto knife”
I concur. You’d assume zebrafish should be tricky, but even drywall can sometimes be a pain in the ass.
It’s more of a in-the-close-ballpark-o knife. But try putting that on a small package label.
I hope the two of you get on well…
>Why is it that you don’t need IRB approval for “spinal >cord butchering on fifteen zebrafish that were only a few >days old.” and yet you would for full grown fish.
Where I am, zebrafish under 5 days old are fair game for anything, over that they fall under IACUC review. It’s arbitrary, I don’t how you decide when a fish embryo can experience pain or whatever. Its nervous system is on par with that of an insect through most of larval development.
If you’re worried about how fish are treated, zebrafish labs are the last place you should be worried about. Go to a fish market sometime, or a pet shop.
Use a laser. But don’t go attaching lasers to the heads of any fish or sharks. Once you start down that path it never ends well.
Fine Science Tools (no relationship) sell thin, breakable razor blades, find them here:
http://tinyurl.com/2modbh
Don’t shell out for the breakers/holders, a pair of No3 forceps will do. Glue them to the ends of orange sticks (wooden bbq skewers will do as well) with Araldite and you will have fine, very sharp micro knives. Finer than micro scissors and with less crushing of tissue and larger and more intuitive than electrolitically sharpened tungsten needles.
I have used them on both mouse and chick embryos to good effect.
Trained baby sharks with friggin laser-beams on their foreheads. It’s the only way to go.
Why is it that you don’t need IRB approval for “spinal cord butchering on fifteen zebrafish that were only a few days old.”
I don’t know about UMM, but at most universities the Institutional Review Board oversees biomedical, behavioral, and educational research that involves Homo sapiens. The IACUC (Institutional Animal Care and Use Committee) oversees research involving all other animals, including embryos and larvae of vertebrate species.
Sharpened tungsten wire “scalpels” and #5 watchmakers’ forceps are my tools of choice for microdissections. Tungsten wire pieces can be purchased pre-sharpened from Fine Science Tools, or you can sharpen your own by running current through a potassium hydroxide solution. An old train set transformer works well for this-have the instrumentation service at the university rig it for you, or find a teenager who is more electromagnetically competent than yourself (which would be almost all of them, in my case). You can purchase needle holders (aka pin clamps) from FST, or design your own holders from polymer clay.
In a pinch, as in no time to wait for special tools, I’d try glass. I’m a glass artist, and the best glass for producing really thin but reasonably durable edges seems to be any thick coloured art glass. Any stained glass worker you know will have tons of shards lying around, and all you need to do to get useful bits is to cut and break a piece crudely.
Just for clarification:
– Students do have agar and mesab available in the lab. I know from long experience that just learning to manipulate a piece of tissue that is 3mm long and a few hundred microns thick is a challenge.
– I don’t allow destructive experimentation on the adult fish in the lab. This is an experiment that assesses regrowth after perturbation in larvae and is an exercise in microdissection and neural staining.
– I’ve been out of town for a few days, but this week we’ll review the procedure to see what is going wrong.
Those poor fish. Gotta feel sorry for em. I’ve had, have Zebras who lived for six-seven years.
It’s a bit of an investment but if you forsee doing more microsurgery type stuff, you can look into buying a gastromaster. I think they’re about $3K and you can do some incredibly precise cutting with them. They are sold by the manufacturer (somewhere in Illinois, I think) and if you google ‘gastromaster’, you’ll find it.
I would try heating up a thin glass rod, pulling it to the requisite diameter and then breaking it with a twisting motion. Alternatively, get some minutennaedlen (pretty close on the spelling) which are tiny insect pins and sharpen them with 2000 grit paper under the microscope. I would suggest practicing on a few dead fish first. When you are trying to do something you don’t really know how to do it is often helpful to break the task down into steps and master each step sequentally.
Have you thought about using a sapphire blade/lance? They’re much more affordable ($30-$115, depending on the specifics) than the diamond lances and are about the same sharpness — they just wear down more quickly than diamond lances (though they’re a *lot* better than stainless steel lances for wear).
I remember seeing them here: http://www.wpiinc.com/products/microdissecting/knives/sapphire/index.html
From their web page:
Diamond knife quality at a fraction of the price
The sapphire knife is a precision cutting instrument for use in ophthalmic, neurologic and plastic surgery research procedures. It provides a superb cut far exceeding that of a finely honed steel knife. Under magnification of the optical microscope, even the best steel blade appears saw-toothed with metal flakes loosely attached to the cutting edge. Such a blade will produce a rough cut and damage to tissue. In contrast, the blades of WPI’s sapphire knife can be polished to a far greater smoothness (less than 0.05 µm RMS). An incision made by a sapphire knife results in significantly less damage than a conventional steel knife and no metal particles are left behind in the tissue, so wounds heal faster with less scaring.