TITLE: Implant Surgery: Chronic recoverable Neuropixels in mice
AUTHORS: Emily A Aery Jones

[DESCRIPTION]
This protocol collection explains how to build a low-cost, lightweight system to implant 1 Neuropixels 1.0 probe or 2 Neuropixels 2.0 probes into mice, record during freely moving behavior, then recover the probes for future use. This protocol explains how to implant Neuropixels probes, previously prepared by following the Assembly protocol instructions, in a mouse. See full collection for more details.

[BEFORE_START]
Test your assembly using a 3D printed skull prior to your first mouse surgery. 3D print a skull (https://github.com/emilyasterjones/chronic_NPX_mouse/tree/main/accessories/Mouse%20skull%20and%20brain), remove the lower jaw, and affix on a stereotax. Use dummy probes, broken probes, or 3D printed probe body replicas (https://github.com/emilyasterjones/chronic_NPX_mouse/tree/main/2.0_probes). Follow this protocol to completion. Confirm the sterics of your design and make adjustments to 3D printed pieces or how they are attached as necessary.
At least 1 day prior to surgery, singly house mouse in post-operative conditions to help mouse overcome neophobia of supporting materals. Prepare a fresh cage with food on the floor, half packet of gel food, half packet of gel water, and sprinkles/forage mix to encourage retaining weight during recovery. Add a nestlet and additional nesting material to support heat retention in single housing. Add enrichment such as chewing materials. Remove any overhead structures like huts or hoppers to prevent implant getting caught.
Machine headbar stereotactic adapters from files at https://github.com/emilyasterjones/chronic_NPX_mouse/tree/main/headforks
Plan your probe trajectory using this tool: https://data.virtualbrainlab.org/Pinpoint/

[GUIDELINES]
Durations are estimates given an experimenter with some stereotactic surgery experience. Once practiced, a skilled surgeon can complete this procedure in 3-4 hours (excluding set up and clean up), or 6-7 hours for dual 2.0 probes. Given the duration and difficulty of this surgery, it is recommended that experimenters with no surgical experience first become proficient in headbar attachment surgery and craniotomy surgery before attempting.
This surgery is adaptable for headfixed recordings. First, perform this surgery, stopping right before the Implant the probe section and skipping the Drill the craniotomy section, performing the Ending the surgery section as normal except skipping the dexamethasone. Train mice until performance criteria are reached. For second surgery, repeat this surgery, mounting directly in headbar stereotax adapters. Skip the shaving and scalp removal steps, skip the Level and mark skull section, drill craniotomy through the metabond, and skip the Mount skull components section except for mounting the well. Continue through Ending the surgery.
Images for dual 2.0 probes show beta probes, which are slightly wider and have dovetail caps on the back (brown side) of the probe. Dual 2.0 probes, once commercially available, will have thinner PCBs with dovetail caps on the front (green side). The assembly and surgery is identical.

[STEPS]
SECTION: Prepare surgical tools and field
1. Sterilize tips of metal instruments, ground screw, and headbar in autoclave 25 min  or hot bead sterilizer 5 s . Disinfect cotton swabs, toothpick, and a weigh boat under UV light 30 min . 

 

SECTION: Prepare surgical tools and field
2. Disinfect surgical field with 10% bleach. Lay absorbent padding across field. Lay down a sterile drape and place tips of metal instruments on this field. Load scalpel blade and drill bits and keep tips over sterile field.
 

SECTION: Prepare surgical tools and field
3. Draw injectables into syringes and place needles on the sterile field or cap them. Place iodine and ethanol each in a weigh boat with 3 cotton swabs. Place remaining tools and consumables and implant assembly on nonsterile pad or within easy reach.
 
  

SECTION: Prepare surgical tools and field
4. Turn on surgical heating pad, lights, scope, O2, and air flow.
SECTION: Prepare animal
5. Weigh mouse and record weight.
SECTION: Prepare animal
6. Set O2 flow rate to 1-1.5L/min and isoflurane to 3%. Place mouse in anesthetic chamber.
 

SECTION: Prepare animal
7. When mouse is immobile, remove from chamber and begin to shave. Shave back to front from behind the ears to just behind the whiskers, and side to side just above the eyes and from the outer edges of the ears. Return to chamber when moving and repeat until area is completely shaved. For final pass, apply Nair to entire shaved area, wait up to 1 minute, then wipe clean with a cotton swab. This ensures no follicles remain in the incision area.

SECTION: Prepare animal
8. When breathing has slowed to 1Hz, move animal to the toothbar. Switch isoflurane from chamber to nosecone. Wait until unresponsive to pedal reflex test, then lower to 1.5% isoflurane.
 

SECTION: Prepare animal
9. Place lacrilube ointment on eyes. Inject dexamethasone, buprenorphine, and saline.
 

SECTION: Prepare animal
10. Immobilize skull in earbars. Adjust nosecone AP and DV position as needed to ensure head is flat and centered. Set earbar on dominant hand side (at 5th tickmark), then hold neck scruff with dominant thumb and middle finger with pointer finger on the nose to manipulate the head. Position and secure earbar on nondominant side (at 5th tickmark). Tighten on both sides, then test by pressing down next to each ear. Confirm immobile and level before proceeding.
SECTION: Prepare animal
11. Sterilize scalp by passing iodine and ethanol with a cotton swabs alternately three times.
SECTION: Prepare animal
12. Cut and remove the scalp from the skull. Lift the middle of the scalp well above the skull and make a small horizontal cut with scissors. Then, make small circular cuts around the first incision. The anterior boundary is at the eyes. The posterior boundary is where interparietal and occipital bones meet (suture behind lambda suture). Whenever possible, move skin away, rather than cutting. Hold remaining skin open with serrefines. Tighten earbars, which are often slightly loose following incision.
 

SECTION: Prepare animal
13. Clean skull with scalpel. Scrape away any connecting tissue and muscle connections to the back of the skull. Superficial bleeding will occur; clear with cotton swabs.  

SECTION: Level and mark skull
14. Find bregma, lambda, 2mm left and right of midpoint along midline suture with drill. Record changes in DV between these points. Level midline axis by adjusting nosecone DV, then repeat measurements. If using Neurostar system, use Adjust for tilt and scaling feature to adjust coordinates.
SECTION: Level and mark skull
15. Make two marks to indicate the ML distance for the two MECs. Go to coordinates AP -3.9, ML +/-3.3. Bring the drill down to the surface of the skull and drill a superficial divot. Remove shavings with air duster, then fill holes with Sharpie. The craniotomy will be centered and behind this dot.
 

SECTION: Level and mark skull
15.1. 2.0 probes: Drill only at left MEC. Drill a divots at AP -3.9, ML -2.7 and -3.7. Fill with Sharpie. The ML extent of the craniotomy will be between these two lines.
SECTION: Level and mark skull
15.2. 2.0 probes: Drill divots at AP -1.5, ML +1 and +2 to target right dorsal hippocampus. Draw a line through each divot, extending anterior & posterior. The ML extent of the craniotomy will be between these two lines. Draw a line extending medial from the medial divot and lateral from the lateral divot. The AP extent of the craniotomy will be centered around these lines.
 

SECTION: Level and mark skull
16. Go to behind the transverse sinus, along the midline (approx. coordinates AP -6.15, ML 0) to drill a hole over cerebellum for the reference screw.

 

 

SECTION: Level and mark skull
17. Score the skull with the scalpel to increase the surface area for metabond to bond to in later steps.

  

SECTION: Drill craniotomy
18. Switch to cool light (e.g. from the scope) to better visualize the sinus. Using the air drill with the 0.5mm bit, shave down the skull in a ~1mm diameter circle behind (not including) your selected dot. Smooth down edges. Regularly stop to remove skull shavings with air duster and to clear and cool the area with saline to improve sinus visualization and to cool the brain to reduce swelling.
 


SECTION: Drill craniotomy
19. Once the skull is very thin above the craniotomy, use the fine-tip drill bit to carve a circular 0.5mm moat (or 1mm oval moat for 4-shank probes) around the area that you will insert the probe (posterior to the dot, right in front of the sinus). You can test whether the skull is thin enough for this step by pressing gently on the skull with bent forceps and seeing how much it gives. If it’s rock-hard, keep shaving it down.
 

SECTION: Drill craniotomy
20. Once the moat has broken through the skull all the way around carefully lift the piece of skull off the brain using bent forceps. Some bleeding will probably occur when you lift off the piece of skull. Stop with Sugi spears. Use the fine-tip drill bit to smooth down edges, and use bent forceps to look for hidden skull ledges to be removed. Try not to remove dura, and don't push the skull fragments towards the sinus.

SECTION: Drill craniotomy
21. When the craniotomy is clear and finished, mark the sinus with a Sharpie and extend the dot anteriorly with a Sharpie, to improve visualization once the well is added in next steps. Apply KwikCast to seal the craniotomy from glue in subsequent steps. 

SECTION: Drill craniotomy
21.1. 2.0 probes: repeat process for hippocampal craniotomy. Drill a 1mm wide (medial-lateral axis) oval centered on the dot. No sinus to worry about.
  

SECTION: Mount skull components
22. Affix screws in the drilled hole. Grasp with the bent forceps and place it on top of the hole. Use a screwdriver to slowly screw it in. Screw to the point where you can’t pull it out with your bent forceps (about 3 rotations; no need to screw it all the way into the skull).
SECTION: Mount skull components
23. OPTIONAL: Attach the probe to the stereotax (see next section) and tax to the ideal probe insertion location relative to bregma (e.g. AP -5.1, ML +/-3.3 for MEC). Tax up, remove the probe holder, and store out of the way. You will cover bregma with a headbar in this section.
SECTION: Mount skull components
24. Select a well size that will allow you to access the full craniotomy with an angled probe, visualize the sinus and center Sharpie marks, but not be too large. Apply a thin layer of UV cure glue, position around the craniotomy, and cure with UV light. This allows rapid curing so the well can be held in place and won't move around during gluing. Once secure, apply and cure a second layer around the outside of the well to make sure it can hold liquid.
 
  

 

SECTION: Mount skull components
25. Place headbar into holder and screw it in. Lower onto the skull. Position just behind the eyes. Center the marker line over the central suture. Lift up 1/2 turn.
  
  

SECTION: Mount skull components
26. Prepare metabond in a mixing well that has been chilled to -20C. Add 1 scoop of metabond powder, 4 drops of solution, and 1 drop of catalyst. Mix well and apply with precision applicators. Let metabond run under the headbar, then lower it down 1/2 turn to put it back on the skull. Use metabond to cover the ground screw (including the soldering junction) and the entire skull. Make a bridge over the headbar with metabond. Apply a thin layer to allow it to cure completely quickly.
 



SECTION: Mount skull components
27. Allow metabond to cure completely for 10 min .

 

SECTION: Mount skull components
27.1. While waiting: Mount rod to Sensapex holder, then to stereotactic arm. Angle the stereotax to 8-10 degrees (for MEC) and tax to just above the target location, either by eye or using coordinates from the earlier measurements from the dot.
SECTION: Mount skull components
27.2. While waiting: Remove rod from stereotax and affix probe. Slowly and carefully dip probe into IPA 3 times to sterilize, then DiD 10 times to dye. Alternatively, to be more cautious, dip using the stereotax. Rotate the stereotax arm 180 degrees so it hangs over the bench rather than the mouse. Place the IPA and DiD tubes into a holder, then mount the probe above them and use the stereotax to dip down into them while visualizing with the scope. Store on a clamp out of the way.
 

 

SECTION: Mount skull components
28. Release headbar from headbar holder and mouse from earbars. Mount headbar stereotactic adapters into earbar slots. Move nosecone anteriorly and ventrally until headbar is positioned snugly in headbar stereotactic adapters. Affix with headbar screws. Record nosecone position for explant surgery.
 

SECTION: Insert the probe
29. Remove Kwikcast and wash craniotomy with saline. Run bent forceps around the rim of the craniotomy to ensure no dried tissue covers the brain; if so, remove it.
SECTION: Insert the probe
30. Mount probe in Sensapex holder to stereotax (at 10 degrees from vertical for MEC) with wings facing rostrally and lower to brain surface. Record DV start coordinate. Make sure entry is unimpeded, advancing 800um quickly (e.g. 0.5mm/s) to pierce dura. If the probe bends, stop immediately and try a different insertion point.
 

 

SECTION: Insert the probe
31. Drive probe at slowest speed (e.g. 3.3microns/s) to final depth (e.g. 3200 angled depth for MEC, 2500 depth for hippocampus). Optionally: insert 200 microns beyond target depth, then retract 200 microns to ensure the probe will not hit skull when the brain settles. If the probe bends during insertion, retract until bend is relieved and either stop driving the probe or try another insertion. Driving to this depth takes 10-15 minutes.
 
    

SECTION: Secure the probe
32. Load blue & clear Dowsil components into 1.5mL Eppendorf tubes for easy access. Mix 20uL each of blue & clear Dowsil gels in a 0.5mL  Eppendorf tube. Draw up slowly as it is viscous. After mixing well, draw  up 20mL, and apply gently to the inside wall of the well. If it spills  out, wait to harden then scrape away.
SECTION: Secure the probe
33. Plug ground/reference pin into screw pin. Apply UV cure acrylic to the joint and cure. Bend the wires inward so they’re flush along the back of the probe.
  
  
SECTION: Secure the probe
34. Build up dental cement on the skull, covering the screw. Using thicker cement, build up from the headbar to the towards wings, covering up to the notch. Do not get any dental cement on the body piece. Build a cement bridge between the wings and from the wing to the screw. Apply cement to fill the gap between the well and the headbar on the left edge. Your goal is to create a ring of dental cement around the probe so that once the outer tape layer is applied, the bottom ring of dental cement will prevent anything from entering the probe zone from below while the tape secures anything from above.
  
  
SECTION: Secure the probe
34.1. 2.0 probes: do the above steps except don't cement the screw. Wait for cement to fully cure. Test this by pressing the dental cement with forceps, it should be hardened and have no give when fully cured. Go to step 37 to remove the Sensapex holder, then repeat steps 28.1, 28.2, and 30-35 for the anterior probe, including cementing the screw. When maneuvering the second probe, make sure you don't collide with any part of the first probe.
 
  
  
 


SECTION: Secure the probe
35. While waiting for the dental cement to cure: apply a thick layer of UV cure glue and affix the dome just behind the well and screw. Make sure there is enough room for the Sensapex holder to open in the next steps. Cure the glue, then apply additional layers to secure it, pulling the neck muscle away from the glue so it securely attaches to the skull. Complete the dental cement ring by using dental cement to attach the dome to the left wing and to the screw.
 

 

SECTION: Secure the probe
36. Release the Sensapex holder from the probe and tax up.
  
  
  
  

SECTION: Secure the probe
37. Using electrical tape to close, snap the flex cable holders on, tab slot facing towards the body piece. Connect the dome antennae to both holder pieces with dental cement.
   

SECTION: Secure the probe
37.1. 2.0 probes: flex holder 1 (with tab slot) faces caudally and goes around the posterior probe. Flex holder 2 (no tab slot) pieces go around the front of the posterior probe and the back of the anterior probe.
SECTION: Secure the probe
37.2. 2.0 probes: place a piece of electrical tape between the edge of the dome and the anterior probe to shield the ground/reference wires. This is to prevent the copper tape added in the next step from connecting the ground/reference to the animal's skin, which would introduce noise into the recording. Alternatively, to reduce weight, fold the wires between the probes and inside the dome so the copper tape won't contact them, then skip this step.
  

SECTION: Secure the probe
38. Wrap in a single piece of copper tape. Make sure the bottom margin doesn't touch skin, but is low enough to not have any gaps above the dental cement. Press to secure to dome, body piece, and sides of flex cable holder. Make slits in front of and behind the flex cable holder and fold the tape in. Press in the remaining larger piece. Put a thin piece of electrical tape along the top margin to prevent it from releasing. Fold over flex cable into tab slot and secure with a similarly dimensioned piece of electrical tape.

 
  
2.0 probes: close the gap between the anterior and posterior probes with a thin piece of electrical tape. Flex cables are folded in an S-bend: down through the flex holder rectangular opening, then up through the tab slot.
  

SECTION: Ending the surgery
40. Inject saline, rimadyl, and baytril. Turn off isoflurane and remove mouse from stereotax. Weigh mouse and record weight. Place mouse on heated pad in a recovery cage. Monitor mouse until walking normally. 
SECTION: Ending the surgery
41. Daily for 3 days, weigh mouse and administer rimadyl, baytril, and dexamethasone. Keep free feeding for at least 3 days post-op to allow weight to stabilize. Monitor for any signs of pain or failure to recover. If needed, follow humane endpoint procedures as detailed in your animal protocol.
SECTION: Ending the surgery
42. Recording can begin 1 day post-op. Number of detectable neurons is highly variable for first 5 days, then remains steady. 

SECTION: Secure the probe
39. Close posterior skin margin with 1 or 2 sutures and apply antibiotic skin ointment to improve skin healing. Cut the back nails of the mouse to reduce the damage they'll do when the scratch the incision.