Tips for spinal cord slices with dorsal roots attached?

Hi everyone! I’m relatively new to the pain field and this has been an amazing resource for getting acquainted with techniques, papers, etc!

So far, I can prepare good transverse spinal cord slices for patching but I’ve been struggling with getting slices with intact dorsal roots/DRGs for suction electrode stimulation. Does anyone have any tips for getting slices with intact roots?

I mostly cut slices embedded in LMP agarose but getting roots this way seems to be up to luck and most of the time they’re short or mangled rootlets. I’ve also tried propping the cord up against an agarose block without embedding but in my hands the cord moves around a lot and it’s hard to “hold” the roots above or below the blade as it’s cutting. I’ve tried parasagittal slicing a few times with no success.

Any tips/tricks/help would be great!

Hi @james_maks
Thanks for your question and joining the forum and for your kind words. I’m so glad it is helpful to you. Please tell your colleagues to check out this site if they’re also in pain research.

I’m going to call in some folks who might have an answer for you:
@MGradwell @gcorder @liz @tberta @marvizon @alex_naka @cedric.peirs

Good luck! Keep us posted.

I have a small amount of experience with this preparation - played around with it but never quite got it working satisfactorily (so there’s surely someone on this forum better qualified to answer).

I found things worked best when I embedded in LMP agarose. In the moments during and immediately after embedding, you can gently manipulate the roots to be parallel to the cutting plane - but you have to be quick to avoid making a mess of the agarose. Also pays to be very attentive to the z-axis - sometimes it makes sense to shave off some throwaway slices in order to line up a good slice that has an entire root attached.

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Hi @james_maks, welcome to the spinal cord/pain field - hope you’re enjoying it!

I’m no expert in DR stim either but did have it working for a while (I now cheat and use genetic strategies). From memory;

  • keep the roots as long as possible when removing the cord. I would trim the root just before the DRG to keep it tidy.
  • gently pull the root away from the cord (it will run alongside before the entry zone)
  • I didn’t embed the cord to slice, though most do. For transverse slices, you can use a styrofoam block with a slight groove cut out for the cord to sit in. If done correctly, the cord should stay put and ‘stick’ to the styrofoam.
  • When slicing, I’d use a brush to guide the root above/below the blade as was necessary
  • A similar technique can be used for parasagittal slices if they better suit the neurons you’re studying.

Not sure if part of this community (@achamess), but I’d suggest Allen Dickie as a master DR stimulator! He definitely embeds too.


Hi @james_maks,

Like the others I have only a little experience with getting slices for DR stim but here are a few things I learnt.

I also prefer to rest my cord on a styrofoam block (like @MGradwell) I would add that in addition to cutting a small groove for the cord kind of sit in I would also add that wicking away any liquid when you place the cord on is extremely important, you can twist the end of a ‘kim wipe’ to make it pointy and gently touch the edges of the cord with it to get rid of excess liquid

gently push the roots away from the cord, as im sure you have seen the pia attaches the underside of the root to the cord so separating this is important for being able to reduce you slice thickness

Especially when you are figuring it out just try leaving a couple of roots attaches, and only on one side of the cord, as I’m sure you have experienced they are VERY sticky so if you are being too ambitious and trying to get every single one you will have problems, as you gain more experience you can up the number.

Be careful not to stretch the roots (it kills them) and there is nothing worse that finally getting a nice slice into your rig and finding out that the root is dead!

I hope this was at least a little helpful and goodluck!

oh, I should also add that Cedric Peirs was a huge help to me, another DR master for you and like Allen Dickie he is also very nice and approachable if you wanted to reach out directly.

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@ksmith5 @MGradwell @alex_naka
Thank you all for chiming in with this your very helpful posts. I’m sure many people will come across this over time and find the info here useful.

I don’t think Allen Dickie is on the forum, but @cedric.peirs is.

Thanks all!

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Thanks @alex_naka @MGradwell @ksmith5 for the tips. This is really helpful. And thank you @achamess for the welcome and tagging everyone!

I’ll try using a styrofoam block and keeping one or two roots attached to start and let you know how it goes.

I’ve tried positioning the roots in the embedded agarose before it solidifies but its hard to gauge when the blade is in the correct z-plane to cut around it. I’m sure it takes a lot of practice so I’ll just keep trying!

@MGradwell I’m hoping to use transgenic/viral approaches in the future myself but I wanted to try the suction electrode prep while I wait for mice to breed. Out of curiosity, which mouse lines are you using to target specific sensory populations?

Hi everyone,
The spinal cord-DRG preparation can be tricky when you do not know what you are doing, but is a piece of cake after practice if you follow the steps. I am currently editing a special issue on JoVe that will address methods to study mechanical allodynia, including how to use this particular slice preparation in transverse and sagittal planes.
Cedric Peirs.


Hi james_maks,
It is quiet tricky to explain how to position the roots for slicing by text only, so hopefully Cedric Jove video will be more helpful.
I share ksmith5 opinion about not being very ambitious. A days with three good roots is a good day.
Anyway, here are some of the things I do that may be helpful :
Before I embed the cord in LM agarose, I put it on a piece of humidified kinwipe in such a way that the dorsal horn and the roots are touching the paper. I then try to position the roots to be as much perpendicular as possible to the cord (be careful not to stretch them). Then I pour the agarose over it. Once the block is solidified, I remove the paper carefully before slicing. Some of my colleague here are putting a really thin piece of paper and are keeping it during cutting.
An other thing is that I use a toothpick or a insect pin to try to tease some of the bigger roots into smaller one (but this may be tricky and you way kill them).
I also adapted the thickness of the slice to the roots so I can cut a a whole segment without touching the root (300µm to 500µm). But I am using fluorescence to pick my post synaptic neurons, so it is not a problem to patch deep.

I hope this helps
Good luck !

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Yes! @cedric.peirs this will be so good!!!

We’ve got the all-star crew of spinal recording people here! :smiley:

Thanks @cedric.peirs and @Amaury for your comments. @cedric.peirs That JOVE will be an important resource Thanks for making it happen.

We did this routinely in my lab. I generally agree with the advice given by @ksmith5 and @MGradwell. Specific instructions can be found is some of my papers:

Marvizon, J C, V Martinez, E F Grady, N W Bunnett, E A Mayer. Neurokinin 1 receptor internalization in spinal cord slices induced by dorsal root stimulation is mediated by NMDA receptors. J Neurosci 17: 8129-8136 (1997).

Marvizon, J C, E F Grady, E Stefani, N W Bunnett, E A Mayer. Substance P release in the dorsal horn assessed by receptor internalization: NMDA receptors counteract a tonic inhibition by GABAB receptors. Eur J Neurosci 11: 417-426 (1999).

Adelson, D W, L Lao, G Zhang, W Kim, J C Marvizón. Substance P release and neurokinin 1 receptor activation in the rat spinal cord increases with the firing frequency of C-fibers. Neuroscience 161: 538-553 (2009). PMC2692762

Chen, W, J A McRoberts, J C G Marvizón. μ-Opioid receptor inhibition of substance P release from primary afferents disappears in neuropathic pain but not inflammatory pain. Neuroscience 267: 67-82 (2014). PMC3998911

Here are some tips:
-The vibratome that you use to cut the spinal cord is critical. We use a Integraslice 7550PSDS with a microscope attached. Previously we used the standard TPI Vibratome, but we had to modify it to decrease the forward movement and increase the vibration amplitude.
-Tilt the microscope so you can examine the dorsal surface of the spinal cord. You can also mount the spinal cord so it is not completely vertical. You will also need a good light source.
-We glue the spinal cord to an agar block using Loctite. The consistency of the agar is critical - it should be hard enough not to move with the spinal cord. Both the spinal cord and the agar have to be dry before applying the Loctite to the agar. The we glue to it the ventral surface of the spinal cord.
-Select the dorsal roots that you are going to cut while cleaning the dura. Cut away all the others. Completely eliminate the ventral roots.
-aCSF bubbled with O2 / CO2 should be added to the spinal cord immediately after gluing it to the agar. We keep the aCSF ice-cold and bubbled throughout the cutting process.
-The cyanoacrylate glue Loctite should work almost instantly. We apply it in a thin layer using the wooden part of a cotton-tipped applicator cut at a slanting angle.
-After you glue the spinal cord to the agar block, separate the roots from the spinal cord by gently pulling. Then grab the end of the first root with fine forceps and sink it in the agar below the cutting plane, so that the root is pulled down away from the blade.
-Carefully aim the blade just above the point of entry of the root by looking through the microscope.
-Cut with minimum forward speed and high transverse speed (vibration).
-Before making the second cut, grab the tip of the root (which was sunk into the agar) and place the whole root on top of the agar block. The root should be stretched a little. Otherwise, you would cut it with the second cut.
-Move the blade down 400 - 450 micrometers.
-Made the second cut. It the root is wide (L4 or L5), you will cut through part of the root, but the remaining should be undamaged. If this is a problem, select L3, L2 or L1.
-Carefully inspect the entry zone of the root through the microscope for damage. Reject the slice if this is not good.
-Detach the slice from the agar by grabbing the strip of Loctite attached to the ventral surface.
-Keep the slice on a nylon net inside a beaker with aCSF bubbled with O2 / CO2. The bubbles will attach to the root and pull the slice to the surface. To avoid this, place a ring with a net over the slice or carefully thread the end of the root through the net.
-Do not use the end of the root (which you have been handling) for stimulation. In my experience, hook electrodes or electrodes on a side chamber work much better than suction electrodes.

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