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Leech Dissection

pbrodfue's picture

Just starting this project

Leech Dissection Protocol

1. Remove leech from bucket with large forceps or finger.

2. Put leech in dissection tray which has been filled with cold physiological saline. Let it sit for approximately 5 mins or until it stops moving

3. Pin through the tail with needle and then through the head. Make sure to stretch animal out as far as possible. It it does not strecth easily, do not force it. You can always stretch if farther after you cut the body wall.



Setting up AxoClamp - 2A

1. After turning on the equipment, be sure the BRIDGE is set to 0 (on both the inner and outer dials).

2. Turn the CAPACITANCE NEUTRALIZATION dial counterclockwise, as far as possible.

3. To test the resistance of the electrode, lower the electrode into the saline. You will know it is in the saline because the voltage reading (which when out of saline is very large) will get very close to zero. Then turn the inner knob of the INPUT OFFSET dial until that voltage reading is at exactly zero.

4. Flip the switch located between the power buttion and the DESTINATION dial from EXT. to CONT. Voltage should read around 50 or 60 mV.


Protocol for Neurobiotin Prep

-When stimulating a DP

1. Dissect a chain of 3 ganglia, with the posterior terminal ganglion desheathed, and the DP for the center ganglion uncovered. Put the well around the desheathed ganglion. (Ex: if you cut out ganglia 9, 10, and 11, desheath 11 and put a well around it, and stimulate DP(10)). Be sure to transfer the tissue in a pipette so that air does not hit the exposed cells. Pin the tissue out in a dish.

2. Wick the normal saline out of the well and fill the well with neurobiotin (20 uL is more than enough).

3. Stimulate the DP every 30 seconds for 15 minutes.

4. After stimulation, wick out the neurobiotin, and replace it with normal saline; allow the tissue to sit for approx. 1.25 hours (the tissue can sit in the dish it is currently pinned out in, or can be pinned out tightly in a petri dish).

5. After letting it sit, pour off the normal saline; pour on 4mL of paraformaldehyde dilution (the stock is 4%; dilute to 2% by mixing 2mL of 4% paraformaldehyde and 2mL of PBS). Let the tissue fix for at least 2 hours.

6. Pour off the paraformaldehyde; fill the dish with PBS; let the tissue sit overnight in the fridge.

7. Pour off the PBS; pour on the antibody (to prepare antibody: 5mL of PBS-X + 50 uL of Cy-3 [each tube of Cy-3 is 50 uL]).

8. Pour off the antibody; wash for 1 hour in PBS.

9. Dehydrate the tissue: pour on 30%, 50%, and 70% ethanol, respectively, for 5 mins each wash. Then pour on 95% and 100% ethanol, respectively, 2 times each, 5 minutes each wash.

10.Cut the tissue out of the petri dish, removing the connective tissue.

11. Clear in methyl salicylate: fill one well of a multi-well plate with methyl salicylate and put tissue in the cell. Let it sit for 5 minutes.

12. Mount the tissue on a slide, and leave to dry overnight (To mount the tissue, adhere 2 coverslips to the slide about half a coverslip's width apart, using methyl salicylate. Then, place a drop of methyl salicylate in between the coverslips where you want to place the tissue. Place the tissue on the slide, and add methyl salicylate as needed. Place another coverslip over the tissue and overlapping the other 2 coverslips using forceps to minimize bubble formation).

13. Visualize the slides with the Confocal Microscope.

-When Stimulating Cell 204

1. Dissect a chain of 4 ganglia. Desheath the most anterior ganglion and the ganglion 2 posterior to that one. Put the well around the anterior terminal ganglion, and stimulate 204 in the other desheathed ganglion. (Ex: If you cut out ganglia 9 through 12, desheath 9 and 11, put the well around 9 and stimulate 204(11).

2. Wick the normal saline out of the well and fill the well with neurobiotin (20 uL is more than enough).

3. Find 204, check the resistance of a newly-pulled electrode (SEE "Setting Up Axoclamp - 2A" above).

4. Lower electrode into cell.

5. Set the stimulation on the MASTER-8: FREE. 4. ENTER. Set the current with the dial below the switch marked 4. Set the current so that there are approx. 15 spikes/sec on the graph (this is around 2 nA).

6. Stimulate for as long as possible, or about 30 minutes. Be sure to keep an eye on your prep in case the electrode floats out of the cell.

7. For the rest of the protocol, follow #4-13 from above.


Protocol for DNQX Prep

1. Dissect the central nerve cord from M2 to the Tail.

2. Uncover DPs from ganglia 7, 10, and 13 if putting a well around M4; uncover DP (4, 7, 13, 14) if putting well around M10; uncover DP (4, 7, 10, 13) if putting well around M15. Pin the tissue loosely in a dish with normal saline.

-Making a well: if you are required to make a well with Vaseline, it is best to have the opening of the   hypodermic needle facing down. As you push the syringe, push down lightly so that there are no openings in the wall. Before pinning out the tissue, test the well for holes by filling the dish and wicking out the well; if the well refills, it has a hole. 

3. Suck some saline into the electrodes. Then take up two DP nerves, one in each electrode (preferably the most anterior and the most posterior, but be sure they are sufficiently active).

4. Stimulate the more anterior DP by flipping the MODE switch for that channel to STIM and then pressing 8 on the MASTER-8. Immediately switch the MODE switch to REC when the MASTER-8 reads zero.

5. Stimulate once a minute until 10 swim bouts are recorded.

6. Stop the recording. Wick out the normal saline from the well. Fill the well with DNQX (add 20 mL of normal saline to 200 uL of 100 uM DNQX--which is one tube--and let stir for a while). Wick out the DNQX and fill the well again. Repeat this 3 more times (washing any residual normal saline from the well), the final time leaving the DNQX in the well. Let sit for 5 minutes.

7. Repeat steps 4 and 5

8. Wick DNQX from the well and fill the well with normal saline. Let sit for 5 minutes. Then repeat steps 4 and 5.

9. Save the data (Filename: the date) as both a "Chart" file and an "Igor" file.

*If you cannot get the tissue to swim, it can be helpful to bathe the entire prep in a dilution of serotonin (100 mL NS in 500 uL 10-3 M); then wash the entire prep with NS for ~10 mins. Then, try getting the prep to swim again.