Much of the work in the Kocot lab is made possible by the effort and generosity of colleagues who collect specimens and preserve samples for transcriptome sequencing using RNAlater. RNAlater-preserved samples need to be treated properly to preserve RNA, though. I regularly get asked for my advice on how to preserve invertebrate specimens and tissue samples using this reagent. Here's my two cents...
RNAlater is a stabilization solution for nucleic acids in tissue samples. It contains ammonium sulfate (to precipitate proteins), EDTA (a chelator of divalent cations), sodium citrate (pH buffer), and water. It has a low pH and thus it will dissolve calcium carbonate, etc.
Official manual for RNAlater:
http://tools.thermofisher.com/content/sfs/manuals/cms_056069.pdf
In my opinion, the official manual is exaggerating when it says samples can sit at room temp for up to a week and in the fridge for up to a month. In our experience with marine invertebrates, it’s more like room temperature for up to 2 days and the fridge for up to a week before risking RNA degredation.
Storage:
Store unused RNAlater at room temperature. If any precipitation forms, you can heat it to ~37° C and agitate to redissolve it.
Usage:
Tissue samples or specimens should be no larger than 1 mm in thickness to ensure penetration. Unless they are very tiny, specimens should be cut into to ensure penetration (but not homogenized!), especially if they have a waxy cuticle like an arthropod or a plant. You want a >20:1 RNAlater:tissue ratio. Avoid transferring any more seawater / culture media / etc. with the tissue sample than necessary.
After adding the tissue sample, invert the tube multiple times to help the RNAlater perfuse the sample, which will likely float. At this point you can store the sample in the freezer (ideally at -80 but -20 is OK). However, it is probably best to invert the tube multiple times right after adding the specimen/tissue sample, put it in the fridge for 15 minutes to a couple hours inverting the tube periodically, and then freeze it within 6-12 hours.
Other notes:
Sarah Gerken asked if I have a formal procedure for collecting with various preservatives that I could share with some colleagues of hers who offered to collect peracarid crustaceans for a grant we are going to write. Here is what I sent them in my attempt to formalize what we do and be thorough.
For morphology for arthropods, I defer to the experts. I’m told that 95% ethanol is OK for morphology but the legs can fall off so 70% is better for morphology. If you add a little glycerin to 95% ethanol to make it something like 1% glycerin, that can help to keep the animals a little more flexible while still preserving DNA well. For soft-bodied marine invertebrates I generally fix things in 4% formalin in sea water at least overnight, rinse thoroughly with sea water or PBS, and then transfer to 70-75% ethanol.
If you plan to do molecular work, I think it’s important to keep in mind the water content of the animal, though, and to try to have on the order of a 20:1 volume of ethanol to animal. Peracarids seem to have a lot of endogenous nucleases so I think it’s helpful to put them in the fridge or a -20 freezer right after fixation, to keep them in the fridge for long term storage, and to never let them get hot.
For transcriptomes, I put specimens or tissue samples in RNAlater or I directly freeze them at -80. For RNAlater I also use something like a 20:1 ratio of RNAlater to animal. It’s important the tissue has a high surface area to volume ratio and if it is something with a cuticle, I usually cut it into a few pieces so it can be well-perfused. In my lab we use the Takara SMART-Seq HT kit to make cDNA. It requires very little RNA. For example, for something like Idotea emarginata, one leg would be plenty of tissue. We routinely make libraries from single nematodes or other meiofaunal animals. More tissue (ideally split up among multiple tubes) is helpful, though. I would recommend trying to take multiple legs per specimen just in case we have a failure at the RNA extraction stage. If there is saltwater clinging to the specimen, I dab the tissue onto a kimwipe or paper towel before putting it into RNAlater because otherwise annoying crystals will form. To preserve tissue with RNAlater I generally put it in the tube, invert the tube several times, and pop it in the fridge for a couple hours, inverting the tubes whenever I think about it. After a couple hours to overnight (but not longer) I freeze it at -80 (-20 is OK I think, but -80 is probably better). The official RNAlater manual will tell you that RNAlater-preserved tissue can sit in the fridge for up to a month and at room temperature for up to a week but they are lairs. I think in reality it is more like in the fridge for up to a week and at room temperature for up to a day. Again it’s important that RNAlater-preserved tissue never gets hot.
Lately I have been freezing as many clean tissue samples as I can for genomics (this also works great for RNA extraction). It’s OK to just put the tissue in a cryo tube and pop it in an already cold part of the -80 but flash freezing in liquid nitrogen is better if you have it handy. Ideally the tissue would be nice and clean so one can put the lysis buffer directly into the cryo tube so the tissue thaws in it. For whole genome sequencing, more tissue is needed than for transcriptome sequencing so if it’s a big animal I tend to freeze multiple tubes containing about a pea’s worth of clean tissue each.
We usually try to collect multiple specimens of a given species and preserve separate animals or tissue samples in the appropriate fixative(s) for morphology, DNA barcoding, transcriptomics, and genomics. For large animals, it’s nice if most of the body can be saved as a morphological voucher with clean tissue samples from non-taxonomically informative parts of the body unlikely to have exogenous contamination (i.e., avoid the gut) in the other fixatives.
If you come across aplacophorans and there are multiple specimens of the same species, please put some in formalin, some in ethanol, some in RNAlater, and directly freeze some. If you just get one of something and it is big, please put the middle 1/3 of the body in RNAlater and the head and tail in ethanol. If you get just one of something that is small but big enough to cut in half, please put the head in ethanol and the tail in RNAlater (the sclerites always point to the posterior). If you get something tiny that you would just smush if you tried to cut it, please just throw it in ethanol.
RNAlater is a stabilization solution for nucleic acids in tissue samples. It contains ammonium sulfate (to precipitate proteins), EDTA (a chelator of divalent cations), sodium citrate (pH buffer), and water. It has a low pH and thus it will dissolve calcium carbonate, etc.
Official manual for RNAlater:
http://tools.thermofisher.com/content/sfs/manuals/cms_056069.pdf
In my opinion, the official manual is exaggerating when it says samples can sit at room temp for up to a week and in the fridge for up to a month. In our experience with marine invertebrates, it’s more like room temperature for up to 2 days and the fridge for up to a week before risking RNA degredation.
Storage:
Store unused RNAlater at room temperature. If any precipitation forms, you can heat it to ~37° C and agitate to redissolve it.
Usage:
Tissue samples or specimens should be no larger than 1 mm in thickness to ensure penetration. Unless they are very tiny, specimens should be cut into to ensure penetration (but not homogenized!), especially if they have a waxy cuticle like an arthropod or a plant. You want a >20:1 RNAlater:tissue ratio. Avoid transferring any more seawater / culture media / etc. with the tissue sample than necessary.
After adding the tissue sample, invert the tube multiple times to help the RNAlater perfuse the sample, which will likely float. At this point you can store the sample in the freezer (ideally at -80 but -20 is OK). However, it is probably best to invert the tube multiple times right after adding the specimen/tissue sample, put it in the fridge for 15 minutes to a couple hours inverting the tube periodically, and then freeze it within 6-12 hours.
Other notes:
Sarah Gerken asked if I have a formal procedure for collecting with various preservatives that I could share with some colleagues of hers who offered to collect peracarid crustaceans for a grant we are going to write. Here is what I sent them in my attempt to formalize what we do and be thorough.
For morphology for arthropods, I defer to the experts. I’m told that 95% ethanol is OK for morphology but the legs can fall off so 70% is better for morphology. If you add a little glycerin to 95% ethanol to make it something like 1% glycerin, that can help to keep the animals a little more flexible while still preserving DNA well. For soft-bodied marine invertebrates I generally fix things in 4% formalin in sea water at least overnight, rinse thoroughly with sea water or PBS, and then transfer to 70-75% ethanol.
If you plan to do molecular work, I think it’s important to keep in mind the water content of the animal, though, and to try to have on the order of a 20:1 volume of ethanol to animal. Peracarids seem to have a lot of endogenous nucleases so I think it’s helpful to put them in the fridge or a -20 freezer right after fixation, to keep them in the fridge for long term storage, and to never let them get hot.
For transcriptomes, I put specimens or tissue samples in RNAlater or I directly freeze them at -80. For RNAlater I also use something like a 20:1 ratio of RNAlater to animal. It’s important the tissue has a high surface area to volume ratio and if it is something with a cuticle, I usually cut it into a few pieces so it can be well-perfused. In my lab we use the Takara SMART-Seq HT kit to make cDNA. It requires very little RNA. For example, for something like Idotea emarginata, one leg would be plenty of tissue. We routinely make libraries from single nematodes or other meiofaunal animals. More tissue (ideally split up among multiple tubes) is helpful, though. I would recommend trying to take multiple legs per specimen just in case we have a failure at the RNA extraction stage. If there is saltwater clinging to the specimen, I dab the tissue onto a kimwipe or paper towel before putting it into RNAlater because otherwise annoying crystals will form. To preserve tissue with RNAlater I generally put it in the tube, invert the tube several times, and pop it in the fridge for a couple hours, inverting the tubes whenever I think about it. After a couple hours to overnight (but not longer) I freeze it at -80 (-20 is OK I think, but -80 is probably better). The official RNAlater manual will tell you that RNAlater-preserved tissue can sit in the fridge for up to a month and at room temperature for up to a week but they are lairs. I think in reality it is more like in the fridge for up to a week and at room temperature for up to a day. Again it’s important that RNAlater-preserved tissue never gets hot.
Lately I have been freezing as many clean tissue samples as I can for genomics (this also works great for RNA extraction). It’s OK to just put the tissue in a cryo tube and pop it in an already cold part of the -80 but flash freezing in liquid nitrogen is better if you have it handy. Ideally the tissue would be nice and clean so one can put the lysis buffer directly into the cryo tube so the tissue thaws in it. For whole genome sequencing, more tissue is needed than for transcriptome sequencing so if it’s a big animal I tend to freeze multiple tubes containing about a pea’s worth of clean tissue each.
We usually try to collect multiple specimens of a given species and preserve separate animals or tissue samples in the appropriate fixative(s) for morphology, DNA barcoding, transcriptomics, and genomics. For large animals, it’s nice if most of the body can be saved as a morphological voucher with clean tissue samples from non-taxonomically informative parts of the body unlikely to have exogenous contamination (i.e., avoid the gut) in the other fixatives.
If you come across aplacophorans and there are multiple specimens of the same species, please put some in formalin, some in ethanol, some in RNAlater, and directly freeze some. If you just get one of something and it is big, please put the middle 1/3 of the body in RNAlater and the head and tail in ethanol. If you get just one of something that is small but big enough to cut in half, please put the head in ethanol and the tail in RNAlater (the sclerites always point to the posterior). If you get something tiny that you would just smush if you tried to cut it, please just throw it in ethanol.
