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Dissection Counter and Fridge Etiquette 

**SUPER IMPORTANT** After the dissection, clean off the counter with multipurpose cleaner & a sponge, then warm tap water, and finally ethanol (wiping it down with paper towels in between steps). This prevents accumulation of (hard to remove) organic material on the counter. 

If you are the last one to use the counter, be sure to spray and wipe down the counter and tools with warm water and 70% EtOH

Make sure you do not keep the saline bottles out of the fridge for longer than necessary. Please also refill them when they are close to empty and wipe down the bottles when they look dirty. 

Wipe off any saline you spill - salt builds up quickly! (on counter and in fridge) 

When leaving preps in the fridge, label with (tape and) Sharpie with your name and date. Do not leave partially dissected preps in the fridge for >1 day. 

Please also replace empty paper towel or glove or kimwipe boxes, refill transfer pipet boxes, replace old sandpaper, etc. 

Carcass Freezer

The carcass freezer gets emptied by the technician or undergrads when necessary. Let someone know if it is getting full and needs to be emptied. 

How to do the STNS Dissection 

To watch the Cancer borealis STNS dissection: the video is located in the Marder Lab Google Drive (–if you don't have access yet please see the Lab Manager.

To watch the Homarus americanus STNS dissection:

Additional Tips: 

Gross Dissection

  • Holding the rongeurs so that the convex side is facing away from you can help with breaking parts of the carapace off. 

Fine Dissection

  • Adjust the magnification on your oculars and distance between them to your liking.  
  • Be aware of your posture when dissecting (keeping a straight back minimizes discomfort).  
  • Pin down the stomach tight + flat against the black dish to ensure good optics. 
  • Adjust light sources and change out saline as often as necessary. 
  • Make sure to keep track of the left + right sides of the dissection - you will not be able to access desheathed STG cells if the prep is pinned upside down. 
  • When removing the prep from the dish, you can start by separating the lower end nerves from the tissue, then move upward to the ions. 
  • Make sure to clean your microdissection tools after you finish the fine dissection. (You can use tap water and EtOH to keep them clean and prevent rusting.) 
    • These tools are expensive, so protect them. You can learn how to sharpen them, but they can be hard to repair if bent/blades get notched. 
  • To avoid dry spots, you can use sandpaper to sand down the Sylgard dish before conditioning it with excess stomach tissue. (If you plan to record intracellularly from the cells, make sure not to sand down the dish too much in the middle where the STG will be. This helps maintain good optics in later steps.) 
  • You can cut pins ahead of time and store them in a small Sylgard dish for later use. 

For step-by-step guides of the C. borealis gross and fine dissections with pictures refer to the following pdf files:

How To Sharpen/Repair Micro-Dissection Tools 

**Please ask someone for a general introduction to these tools before using them for the first time!

Use the sharpening block and mineral oil to grind away parts of the tools that are notched, misaligned, blunt, etc. As you do so, frequently use a kimwipe to wipe off the oil and examine the points. (The oil sometimes bends light in ways that makes you think the tools have been completely fixed when that is not the case.) 

Do your best to grind sparingly - the more steel you grind away, the smaller your point of contact (for forceps), and then the tools don't work as well. 

If you are trying to repair forceps that became bent at the tip, use round nose pliers to gently and carefully bend the tip(s) back into alignment and then sharpen them together. 




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