Overview: Sorting is a manner in which we can take the field samples we have gathered and isolate the arthropods from the debris they are mixed in with. Samples can be gathered in different ways, such as through beating which would have less debris than when gathered through leaf litter. Depending on the method used to gather the samples and ratio of debris to arthropods in the sample, it may be easier to remove the debris or the arthropods themselves. This is done through examining the poured out sample through a microscope and finding the arthropods in the sample. Once all the arthropods are gathered, we take a photo to store this information. Following this, the arthropods are collectively weighed and their weight is recorded. These now isolated arthropods are lastly transferred into a lysis buffer to property store them for later use and examination.
- Prepare workspace: Make sure to work in a well ventilated area. Put on gloves and use 10% bleach to sterilize your work area.
- Gather supplies: petri dishes, forceps, small beaker for bleaching forceps, lighter, ethanol lamp, ethanol squirt bottles, sharpie, 15 mL tube, lysis buffer, and p1000 micropipette and tips.
- Select sample from freezer and record sample ID in appropriate spreadsheet. Label clean 15 mL tube with sample ID.
- Remove debris from sample: Pour sample into petri dish. Use ethanol squirt bottle and forceps to make sure entire sample is extracted from the tube. Add enough ethanol to petri dish for specimens to be submerged, if necessary. Place petri dish on white laminated paper on top of an ice pack under the stereoscope/microscope. Adjust and focus the microscope as needed. Use sterilized forceps to remove any debris (plant or other matter) from the petri dish. You can use another petri dish or a kim wipe to put the debris temporarily before discarding in the trash. Alternatively, you can move the arthropod specimens onto a new, clean petri dish, leaving the debris behind. This is especially helpful when working with dirty samples like leaf litter.
- When the sample is free of debris, move petri dish and label on white background to the camera station.
- Take photo of sample: Place petri dish and label on white background under the camera. Place the ruler next to the label at a right angle.
- Look through the viewfinder to make sure the sample is centered and the camera is focused. Use forceps to spread out the specimens, minimizing overlap. Take photo with the camera.
- Weigh sample: After taking a photo, strain sample through nylon mesh strainer over discard container/beaker (ethanol can be discarded down sink drain). Let sample rest to drain for about 1 minute. You can dab a kimwipe to absorb excess liquid from outside the bottom of the strainer. Tare scale with a new clean weigh boat. Dump the samples from the nylon mesh in the clean weigh boat. Use sterilized forceps to gently transfer any remaining specimens to the weigh boat. Make sure the scale is tared.! If you forgot to tare the scale before transferring the sample or if the scale turned off, tare the scale now using a different clean and dry weigh boat. Place the sample on the scale and record wet weight (mg) in the spreadsheet. (Make sure you are only recording the weight of the sample, not the sample + weight boat.)
- For samples (usually Malaise) with oversized specimens: Pull off 1-2 legs from each oversized specimen (depending on the ratio of body size to a normal body size) to leave with the sample and record the weight without the bodies under Wet Weight Column (the first weight will go under “Wet Weight with Bodies). Put the bodies in a new 15mL tube with 95% EtOH from the aliquots. Label the tube and cap and store in the -20 freezer on the top shelf.
- Clean nylon mesh strainer between samples: Agitate nylon mesh strainer in 10% bleach for 1 min between samples. Transfer to DI water and agitate for 1 min. Transfer to second DI water wash and agitate for 1 min.
- Transfer sample and lysis buffer to tubes: Add 1 ml of lysis buffer (in labeled 50mL tubes, not in freezer) with p1000 micropipette to the labeled 15 mL tube. Transfer sample from weigh boat to labeled tube with clean forceps. Make sure there is enough room in the tube for the lysis buffer to generously cover the specimens. Add more lysis buffer as needed according to the table below. There should be about double the lysis buffer relative to the approximate volume of specimens in the tube. This does not have to be exact and it can be adjusted later if needed.
- Use a new tip for each sample when adding more lysis buffer – be careful to not contaminate the stock of lysis buffer. If the lysis buffer has been contaminated, discard tube.
- Samples in lysis buffer can be stored in the refrigerator in a tube rack until further processing.
Sample Weight (mg) | Volume Lysis Buffer (mL) |
| 0-50 | 1 |
| 50-100 | 3 |
| 100-200 | 6 |
| 200-400 | 8 |
| 400+ | 10 |
| split into two tubes, label “1 of 2” and “2 of 2” |
Cleanup: Turn off stereoscope lights. Cover stereoscope. Extinguish ethanol lamp flame. Turn off cameral and cover lens. Make sure completed samples in lysis buffer are labeled and placed in the freezer. Put freezer packs back in the freezer. Put petri dishes, forceps, used weight boats, mesh strainer, squirt bottle tips, and beakers in bucket with 10% bleach solution for ~2 minutes. Then, rinse in first DI water bath, Rinse again in second DI water bath. Let dry on paper towels next to the wash bins. Nylon mesh used for straining should be sterilized in 10% bleach and two DI water rinses. “Dirty” ethanol from straining samples can be carefully discarded down sink drain.