The worms will often still move a lot when it is in a squeeze preparation (see Observation), and this can make it hard to make detailed observations. Worms can be anesthetised with a solution of MgCl₂ in distilled water (tap or mineral water will often also do). For marine samples I usually use a 7.14% solution (i.e. 7.14g per 100ml), and I place the worm into a 50:50 (or even 60:40) mixture of MgCl₂-solution and sea water. In contrast, freshwater species are much more sensitive to MgCl₂, and so in these I will often use only a few percent of solution (too high concentration will stop the beating of the cilia, and make the epidermis crumply and less transparent, which is not nice for observation). What work better in freshwater is Phenoxypropanol (see below).

Once you have the worm in a nice squeeze preparation go through a series of magnification steps to document more and more detailed structures of the worm. I always start taking pictures with the 4x objective, even if the worm is very small. This makes it easier later to reconstruct the magnification at which the different pictures were taken. I try to take pictures of the following structures: habitus, eyes, brain, pharynx, pharynx glands, testis, sperm in testis, ovary, forming eggs, eggs and/or sperm in female antrum, eggs, vaginal cilia, shell glands, false seminal vesicle, seminal vesicle, copulatory stylet, prostate glands, vesicula granulorum, male pore, male pore cilia, sperm, epidermis, epidermal cilia, sensory cilia, rhabdites, rhammites, adhesive glands, and food items.

For taxonomy the most important structure is of course the copulatory stylet, but also the structure of the female antrum and the sperm morphology is important. However, the sperm morphology is a little tricky to document, because you would like to document the sperm from the seminal vesicles, which are ready to fertilize, rather than the forming sperm form the testis. This means that you have to somehow make sure what you are looking at. 

With a bit of practice you will be able to recover worms from the squeeze preparations to do some additional things with it. To do this place a drop of water (usually the stuff the worm was held in before you looked at it) next to the cover slip. The capillary force will draw in the water and the strength of the squeezing will be reduced. Generally the worm will start moving around again below the cover slip and you can then lift the cover slip with pair of fine watchmakers forceps.

What I then usually try to achieve is to amputate the tail plate, and to make another squeeze preparation with this, while I'll place the frontal part of the worm into 100% Ethanol or RNAlater for a DNA and/or RNA sample.

Example Workflow

1. Clean the slide and the binocular plate
2. Take new 100µl pipette tip
3. Place the worm in the centre of slide in about 40µl while observing it
4. Anesthetize the worm
   • In freshwater use Phenoxypropanol and water. A concenration of 0.015% is a rough guide, but concentrations vary with species. Use less and add more if needed.
   • In saltwater add 7 % MgCl2 in 1:1 or lower concentration
5. Clean cover slip and place small plasticine feet on its corners (see Observation)
6. Place coverslip on slide with fine tweezers (try to move the slide to orient the worm with the ventral side towards the observer, it facilitates observation of female structures)
7. Take pictures of the worm under the Microscope. Use all Magnifications (4x,10x,20x,40x,100x). I use DIC only for 20x,40x and 100x
   • 10x orient the camera, pictures of all layers and maybe short movie, document distribution of rhabdites
   • 20x DIC, movie, sensory hairs
   • 40x special focus on testis, ovary, stylet, seminal vesicle and antrum
   • 100x same as for 40x, plus focus throught the area around the eyes
8. Add peripheral drop to relax squeeze preparation, remove cover slip with tweezers and recover worm
9. Cut off the tail plate with one continuous rolling of the scalpel
10. With small pipette suck up anterior part in 1µl and quickly transfer to PCR tube with EtOH (for DNA samples) or RNAlater (for RNA and DNA samples). Don’t touch EtOH with tip or worm will get stuck!
11. Check if worm actually is in PCR tube, and place a mark tube was filled
12. Change tip and clean a new slide
13. Transfer the tail plate with 1µl onto the clean slide (it is extremely impprtant that there is no dust to achieve a thin preparations that shows the sperm in 2D)
14. In freshwater species add a small quantity of 10‰ ASW. Otherwise the osmotic shock will change the shape of the sperm
15. Place coverslip without feet to smash tail plate, releasing the sperm
16. At 40x take picture of stylet, locate the sperm (it canbe hard to find them at 100x)
17. At 100x take pictures of sperm (if you also have phase contrast, use it)
18. Add some drops of Lactophenol on the side of the slide (don’t get it on your fingers!)
19. Label slide
20. Label EtOH PCR tube on 3 places and tape the sides with scotch tape (EtOH can dissolve the permanent marker)
21. Enter information into datasheet, archive the images

Tips and Tricks

• Use enough fluid to place the worm on the slide to avoid smashing it
• Clean everything nicely to avoid something cutting the worm or obstructing the view
• Gently lift the coverslip with tweezers to turn the worm
• Orienting the worm with the ventral side towards the observer facilitates imaging the antrum and received sperm
• Have as many informative structures on a picture as possible
• Think about orienting the worm in a way that pictures could be stitched together later for pulication