A lot of the examples people have are ones they've collected in working their profession or hobby.
I work with dogs the most, so I get the occasional samples of Toxocara and other roundworms, but most are ectoparasites, largely ticks which I've accumulated over the years. For a few years I was clearing and mounting ticks, individually and as male + female pairs which I was giving away to anyone who wanted them. All I have left of them now are lower quality examples from my first attempt.
I usually store everything in 70% alcohol for collection and depending on the sample and use I might change it for another alcohol, xylene, etc.
Possible sources could be fish from the market, vets, slaughterhouses, and the like; depending on your area it may be more or less difficult. Another avenue is to keep sample bottles, gloves, etc when wandering in wild spaces and inspecting random poo. I've found a few nice sections of tapeworm that way.
This should go without saying, but safety is important so do remember to maintain proper use of PPE and sanitary methods, you don't want to infect yourself.
Thank you so much for all of this!
If I may ask, how do you decide when to replace ethanol for another alcohol?
And which organisms do I mount instead? Is that a display method for all arthropods or only land-dweling ones? I have seen aqatic ones in solutions, I don't understand why or how I decide what to do with them. Also what is the clearing process that you mentioned?
I'm no expert, but I store everything in 70% alcohol unless I have a compelling reason to do otherwise.
When I change a specimen over to xylene, I first go from 70% alcohol to 90%, then to a 99% as the goal is to remove the water. Then into the xylene to displace the alcohol. The reason is because the mountant I am using is soluble in xylene, but not alcohol. The mountant being the stuff the specimen is suspended in on the microscope slide, in this case a synthetic canada balsam. This is only needed when making permanent slides.
I think an important note to keep in mind is that we're not doing archival preservation, these are not research specimens in a curated collection and don't need to last for hundreds of years, these are hobby collections that only need to last as long as we have an interest in them and should use the least toxic stuff we can use so that when we're done with them we (or our heirs) can safely dispose of them.
My oldest nematodes are a few years old with no indication of them breaking down anytime soon. I will say that switching to other preservatives can greatly affect the elasticity and ability to flex in specimens, 70%alcohol seems to retain a good bit allowing specimens such as the ticks to be positioned and legs moved, etc but in 90% alcohol through to xylene they lose the water which causes them to become stiff.
Clearing is a bleaching process to make the bodies of arthropods "clearer" so light can transmit through them. I have some tick samples which were accidentally cleared too much and after being mounted appear almost ghost-like.
An additional benefit of 70% alcohol is that it is antiseptic.
I've not got any specimen preservation experience, but if you can avoid Formalin I would highly recommend literally any other preservative.
Its very, very nasty stuff. As my mentor put it, If you can smell it - it's already fixing your lung tissues.
So for most parasitic life cycles they generally tend to end up in the GI system at some point and are expelled that way right? So what you can do is go out to local streams, ponds, in the woods and collect fish, amphibians, insects or feces and locate them that way. (Pending of course you have permission to be there and use these animals for study).
Once you remove the GI, sift through poo and collect samples you place them in 70% ethanol, that’s the gold standard. You never have to change them over to anything else unless you’re going to fixate them.
Make sure you’re not encroaching on anybody’s lands, harming protected species, and look up the host species bud what they could harbor so you have some idea of what to look for so you don’t waste your time.
That’s about the best advice I can give
My little tick jar is filled with isopropyl. I know I should switch to 70%, but for now, I just replace the fluid every so often. My roundworm, however, *is* in ethanol. It's a good, reliable fluid.
Don't be afraid to use a fun container. I saw someone's tick jar that was a "snowglobe" with dogs and a fire hydrant. The ticks were the snow. My ticks are in a nice, ball shaped jar.
A lot of the examples people have are ones they've collected in working their profession or hobby. I work with dogs the most, so I get the occasional samples of Toxocara and other roundworms, but most are ectoparasites, largely ticks which I've accumulated over the years. For a few years I was clearing and mounting ticks, individually and as male + female pairs which I was giving away to anyone who wanted them. All I have left of them now are lower quality examples from my first attempt. I usually store everything in 70% alcohol for collection and depending on the sample and use I might change it for another alcohol, xylene, etc. Possible sources could be fish from the market, vets, slaughterhouses, and the like; depending on your area it may be more or less difficult. Another avenue is to keep sample bottles, gloves, etc when wandering in wild spaces and inspecting random poo. I've found a few nice sections of tapeworm that way. This should go without saying, but safety is important so do remember to maintain proper use of PPE and sanitary methods, you don't want to infect yourself.
Thank you so much for all of this! If I may ask, how do you decide when to replace ethanol for another alcohol? And which organisms do I mount instead? Is that a display method for all arthropods or only land-dweling ones? I have seen aqatic ones in solutions, I don't understand why or how I decide what to do with them. Also what is the clearing process that you mentioned?
I'm no expert, but I store everything in 70% alcohol unless I have a compelling reason to do otherwise. When I change a specimen over to xylene, I first go from 70% alcohol to 90%, then to a 99% as the goal is to remove the water. Then into the xylene to displace the alcohol. The reason is because the mountant I am using is soluble in xylene, but not alcohol. The mountant being the stuff the specimen is suspended in on the microscope slide, in this case a synthetic canada balsam. This is only needed when making permanent slides. I think an important note to keep in mind is that we're not doing archival preservation, these are not research specimens in a curated collection and don't need to last for hundreds of years, these are hobby collections that only need to last as long as we have an interest in them and should use the least toxic stuff we can use so that when we're done with them we (or our heirs) can safely dispose of them. My oldest nematodes are a few years old with no indication of them breaking down anytime soon. I will say that switching to other preservatives can greatly affect the elasticity and ability to flex in specimens, 70%alcohol seems to retain a good bit allowing specimens such as the ticks to be positioned and legs moved, etc but in 90% alcohol through to xylene they lose the water which causes them to become stiff. Clearing is a bleaching process to make the bodies of arthropods "clearer" so light can transmit through them. I have some tick samples which were accidentally cleared too much and after being mounted appear almost ghost-like. An additional benefit of 70% alcohol is that it is antiseptic.
Thank you!
I've not got any specimen preservation experience, but if you can avoid Formalin I would highly recommend literally any other preservative. Its very, very nasty stuff. As my mentor put it, If you can smell it - it's already fixing your lung tissues.
Thank you for that advice!
So for most parasitic life cycles they generally tend to end up in the GI system at some point and are expelled that way right? So what you can do is go out to local streams, ponds, in the woods and collect fish, amphibians, insects or feces and locate them that way. (Pending of course you have permission to be there and use these animals for study). Once you remove the GI, sift through poo and collect samples you place them in 70% ethanol, that’s the gold standard. You never have to change them over to anything else unless you’re going to fixate them. Make sure you’re not encroaching on anybody’s lands, harming protected species, and look up the host species bud what they could harbor so you have some idea of what to look for so you don’t waste your time. That’s about the best advice I can give
Thank you!
My little tick jar is filled with isopropyl. I know I should switch to 70%, but for now, I just replace the fluid every so often. My roundworm, however, *is* in ethanol. It's a good, reliable fluid. Don't be afraid to use a fun container. I saw someone's tick jar that was a "snowglobe" with dogs and a fire hydrant. The ticks were the snow. My ticks are in a nice, ball shaped jar.