Surgical Records

Archive of surgical protocols and logs

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Comprehensive Rodent Surgery Checklist

Merged from Kemere lab protocols and Shay’s personal protocols

I. Pre-Surgery / Day Before (Ideally)

1. Sterilization

Rice:
- Autoclave: Double-bag tools. Place on autoclave tray(s).
  - Location: BRC Room 860G.
  - Setting: “Gravity” option (42-46 min).
  - Login: KEMERE / 740.
  - Autoclave Wiki Link
- ETO Sterilization (if needed for plastics/drives):
  - Content: Drive body with cone, cautery pen.
  - Location: BRC Room 860G.
  - Protocol: 12hr cycle + 12hr fume hood off-gassing.
  - Grab keys (fridge) and ampoule.
CU Anschutz:
- Autoclave: Double-bag tools. Place on autoclave tray(s).
  - Location: RC1N P18-7208 for the 7th floor one
- Ethanol/UV for electronics:
  - For VNS cuffs

Step	Action	Duration	Purpose
1. Gross Decontamination	Immediate soak in 1% Terg-a-zyme or Alconox solution.	10–15 mins	Breaks down proteins and biological films from the previous implant ONLY IF RECOVERED.
2. Mechanical Cleaning	Gentle scrub with a soft-bristled brush under a microscope.	As needed	Removes stubborn tissue fragments from cuff crevices.
3. High-Purity Rinse	Triple rinse with Deionized (DI) water or Sterile Saline.	2 mins	Removes residual detergents which can be neurotoxic.
4. Chemical Sterilization	Submerge in 70% Ethanol.	20 mins	Primary disinfection; disrupts lipid membranes of pathogens.
5. Drying Cycle	Air-dry in a Laminar Flow Hood (Class 100). Technically, my Grelife UV sterilizer also works but don’t do more than 20 minutes b/c heat and electronics may not play well together. The 20 minute cycle with the heat this provides has been validated to maintain electrical impedance of the VNS stimulation cuffs.	Until dry	Ensures no ethanol is trapped in insulation before UV exposure.
6. UV-C Irradiation	254 nm UV exposure in box. Grelife sterilizer that I left in the lab no need to flip sides with this.	20 mins/side or 15 minute with Grelife sterilizer.	DNA/RNA cross-linking; reaches areas not fully saturated by ethanol.
7. Sterile Storage	Place in autoclaved pouch or glass vial or sterilized glass petri-dish.	Indefinite	Maintains sterility until the next surgical procedure.

2. Instrument Staging

Bag 1 (Incision on scalp or cervical area):
- Scalpel handle + #10 or #15 blades
- Small scissors
- 4 Retraction clips/clamps (Rat or VNS only)
- 4 Hemostats (Rat only)
Bag 2 (Bone Work):
- Skull Scraper (Rat only)
- Bone wax (Rat only)
- Fine Tweezers (for bone picking)
- Blunt Tweezers (for holding screws)
- Micro-forceps (#5, #5/45)
Bag 3 (Cleaning)
- Cotton swabs (small and larger tipped)
- Cotton (small or ripped up into small pieces)
- Beaker (2) for screws/saline and ethanol
- Aluminum foil (Rat only)
- Sterile field sheets (2)
- Needle driver for suturing (Rat or VNS only)
- Paper towels (more the better you can touch anything/everything with a paper towel while being sterile)
Drill Bits:
- Kemrelab Rat protocol:
  - 2x FG2 (for 0-80 screws/big holes)
  - 2x FG 1/4 (small holes)
- Wellelab Mouse Protocol
  - TODO
Screws:
- Kemerelab Rat Protocol:
  - Skull screws (12x 0-80 3/32″) in glass jar with perforated top.
  - Ground screws (0-80 5/32″).
Implant Specifics:
- Silicon sheet (cut to size).
- Spatula (for acrylic).
- Cannula position map
- Vagus nerve surgery
  - Cuffs (have a backup ready)
  - Stimulator hardware
    - Wellelab: Pulsepal + A167 stimulus isolator (remember to charge beforehand) + Powerbank (remember to charge as well!)
  - Bag 1
    - Blunt dissection scissors
    - Blunt tweezers from Bag 2 should be placed here
  - Bag 2
    - Fine forceps (3 or 4 total with one bent/broken/blunt if existing!)
    - [ ] Fine hooks for vagus nerve and sternocleidomastoid (2)

II. Surgery Day Setup

1. Room & Equipment Prep

Clean: Mist Cavicide or 70% Ethanol on surgery table, stereotax, arms, scope, drill, and light source. Wipe down.
Heating Pad: Wrap in blue pad (diaper), tuck under stereotax, turn ON.
Stereotax: Clean earbars/nosecone with ethanol. Tighten loose screws.
- Kemerelab Settings: Left earbar ~8mm; Bite bar ~6.5mm.
Anesthesia: * Check Isoflurane level (> 3/4 full).
- Weigh/replace carbon trap (scavenger).
- Open O2 valve (Rice: 860B).
Aspiration/Vacuum Setup: Beaker with 10% bleach solution (20ml bleach + 180ml water).

2. Drug Preparation

Weigh Animal: Calculate dosages based on weight.
Buprenorphine (Ethiqa): * Dose: 0.03 mg/kg SC (or check bottle instructions).
- Note: Use between shoulders.
Lidocaine 2%: Draw with 23Ga (for rat) or 25Ga (for mouse) needle (Max 0.5ml/kg).
Meloxicam: For post-op analgesia (2 mg/kg SC. Typically going to be 5 mg/ml).
Fluids: 3-5 syringes of Lactated Ringer’s solution (warm if possible; 10ml/kg or 1ml/100g).
Topical: Ophthalmic ointment & 2% Lidocaine cream.

3. Shay logging methods

Convert markdown/html surgical template to PDF then edit PDF on iPad
- Spray iPad and Apple Pencil with 70% ethanol AFTER noting induction time in step below.
  III. Anesthesia & Positioning
  1. Induction:
- Place rat or mouse in chamber, Isoflurane @ 5 or 2.5%, O2 flow @ 1-2 L/min, respectively depending on species.
- Watch breathing: Ideal 60-70 bpm. If gasping, reduce Iso.
- Safety: Never leave room once animal is under or ask someone to watch the animal!
  1. Shaving:
- Transfer to nosecone (reduce Iso to 2-3% for rat or to 1-2% for mouse).
- Inject Ethiqa/Buprenorphine SC between shoulders now (Rice)
- Shave from mid-eyes to behind ears for cranial surgeries. Careful of whiskers. Shave from animal’s left shoulder area up to bottom of chin for vagus nerve surgeries whilst they are in a supine position. Be very gentle with shaving
  - For mice, especially with for the cervical shaving, Shay recommends to use Nair briefly to get rid of excess hair as that may sneak into the open surgical site.
    - Leave Nair on for about 15-20 seconds and clean with DI water/sterile cotton swabs 3. Stereotax Mounting (if needed):
- Apply Lidocaine cream to ear bars.
- Mount rat or mouse: Ear bars in canal crease. Ensure head does not move side-to-side.
- Secure bite bar (tongue to side) and nose cone. Tighten dorsal screw.
- Ensure head does not move up or down
- Check: Breathing (no gasping), Skull Leveling.
  1. Sterile Prep:
- Apply eye ointment + foil shades (for rat).
- Scrub scalp with Iodine (3x spiral from center out). Wipe with saline or ethanol.
- Inject Lidocaine SC at incision site (base of skull to eyes and retract slowly while injecting).
  - Inject Lidocaine 0.05 ml SC at cervical incision site. CAREFUL of not injecting too much or going too deep here as that could significantly hurt the animal based on location of the injection
- Spray glove/tools/drill with 70% Ethanol.

IV. Surgical Procedures

Cranial Procedures

1. Incision & Exposure

Incision: Midline from eyes to back of skull (~1.5 inch) for rats; For mice, just cut out the skin with fine scissors.
Retraction: Place 4 hemostats/clips. Ensure flat field. (FOR RAT ONLY)
Cleaning:
- Rat: Scrape tissue/periosteum with skull scraper from center out
- Mouse: Wipe skull with sterile cotton swab dipped in saline first and clear out any blood. Follow this with brief touch of ethanol on saline which will dry out the skull enough for any cranial implants according to Cristin (headbars specifically, for microdrives you will need anchor screws).
Goal: Bregma and Lambda must be clearly visible.
Hemostasis: Use bone wax or cautery if necessary. This should not be necessary in mice!

2. Calibration

Mount 32G (or finest available) needle or calibration tool.
Measure Bregma (B) and Lambda (L).
Level Skull: Z-difference between B and L should be < 0.3mm for rats, whereas, for mice Shay would shoot for < 0.1mm. Adjust bite bar if needed.

3. Anchors & Craniotomy

Drilling Screws (if needed):
- Rats:
  - Use FG2 bit. Low speed.
  - Pattern (for hippocampal target): 4 Anterior, 2 Posterior (avoid craniotomy sites).
- Ground Screw: Posterior location. Drill until deep red/blood seen. Screw in immediately (1-2 turns).
Metabond Base:
- Clean skull of blood.
- Apply Metabond (4 drops liquid : 1 scoop powder : 1 drop catalyst) around screws. Place liquids and powder in separate wells and mix to consistency that you need. Avoid craniotomy site.

4. Specific Procedure Branches

Option A: Microdrive Implant

Craniotomy:
- Use FG 1/4 bit for rats. Use finer available drill bits for mice
- Drill target coordinates (see reference below).
- Remove bone island carefully (don’t push down!).
- Remove Dura (bend 27G needle tip to make incision into dura and lift).
  - There are a million different ways to do a clean duratomy. Just be careful and avoid blood vessels lines and have a plan of incision and peeling of dura before you get started. Adjust as needed and ask for tutorials from multiple people. Not one way is correct or better on this! You’ll find your own best practice.
Insertion:
- Mount drive to arm. Align vertical.
- Lower until bundle touches cortex surface.
- Drive: Lower to target depth (e.g., -0.8mm for cortical touch or specific coord).
Sealing:
- Apply Silicone Sheet or Kwik-Sil or Duragel around edges.
  - For larger sized electrodes (i.e., not tetrodes), silicone sheet or duragel is recommended as Kwik-Sil will adhere to the electrode making it not drivable; thus, defeating the purpose of a microdrive.
- Build metabond followed by (in mice) acrylic column from bottom up to secure drive to skull/screws.
  - This takes practice but can be done very cleanly and look pretty!
- Grounding: Connect ground wire to ground screw pin. Cover with acrylic.
- Tip: Use UV curing acrylic! Makes life much faster/easier and avoids potential exothermic reaction issues caused to the brain when occurring on the skull surface for extended durations.

Option B: Stereotaxic Injection (Hamilton)

Prep: Clean syringe with acetone/distilled water.
Test: Push “Run” to verify flow.
Target: Go to coordinates.
- Technique: * Rats: Lower 2mm past target, retract 1mm (create pocket), or specific depth.
  - Mice: TODO. Ask Ziying. He’s done this.
Inject: * Rate: ~0.05 - 0.2 ul/min.
- Wait time: 5-10 mins post-injection before retracting.

Option C: Aspiration & GRIN Lens

Durotomy: Remove dura over aspiration site.
Aspiration: Vacuum (10% bleach trap) to remove cortex to desired depth.
Implant: Lower dummy/GRIN lens to target. Cement in place.
Vagus Nerve Surgeries

1. Incisions

2. Field of view

V. Closing

Kwik-Sil: Apply to any exposed brain/craniotomy gaps.
“Dust and Squirt”:
- Acrylic powder + Liquid monomer.
- Cover all screws and ground wire connections.
- Warning: Do not let acrylic run into eyes (your’s or the rodent’s!)
Suturing:
- Simple interrupted or running sutures for skin around the implant.
  - See suturing resources for further details!
- Vetbond on knots (if not confident/needed)

VI. Post-Operative Care

1. Immediate (Unconscious)

Turn off Isoflurane, O2, Flush O2.
Remove mouse/rat from ear bars gently.
Apply fresh eye ointment.
Place in recovery cage on heating pad.
Administer Fluids: Lactated Ringers SC (5ml or 0.2ml for rats or mice, respectively).
- This isn’t necessary but it aids recovery regardless of when fluid function returns.

2. Recovery (Conscious)

Monitor for “Righting Reflex” (rodent can flip itself over).
Return to clean cage with food/water.
Analgesia: Meloxicam (Day 0, 1, 2 post-op).

3. Cleanup

Discard sharps/biohazard.
Wash tools (Updated Protocol for VNS/Cranial Instruments):
- Critical Rules: No Dawn/dish soap. No overnight soaking in standing liquids.
- Prep: Mix fresh 1% Tergazyme in warm DI water daily (enzymes expire after 8 hours).
- Immediate Clean: Rinse with DI water $\rightarrow$ Soak in fresh 1% Tergazyme (15-20 mins max) $\rightarrow$ Scrub hinges with nylon brush $\rightarrow$ Triple rinse with DI water $\rightarrow$ Apply water-based surgical lubricant (e.g., Surgilube) to hinges $\rightarrow$ Flash dry with lint-free wipes (Never air dry).
- Delayed/Overnight Clean (Moist Pack): Coarse spray tools with fresh 1% Tergazyme $\rightarrow$ Seal in Ziploc bag $\rightarrow$ Next day: Execute “Immediate Clean” steps above.
Wipe down stereotax.

Reference

Coordinates & Diagrams

Rats

Perirhinal Cortex (PRh):

AP: -4.08 mm (range 3-9.6)
ML: ±4.57 mm (15° angle)
DV: -7.45 mm

CA1:

AP: -3.6 mm
ML: ±2.5 mm
DV: -2.6 mm

VHC

AP: -1 mm
ML: ±1 mm (closer the better but obviously this is slightly difficult with the suture lines)
DV: 4 mm

Diagram: Skull & Screw Placement | Craniotomy | Ο | | :—: | :—: | | Ground/Ref Screw | Ø | | Anchor Screw | x |

Useful Guides

If you’re just getting started, Shay strongly recommends going through all of these!