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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Pre-surgical Procedure
- Before starting surgery, wipe everything with 70% ethanol to maintain sterile conditions. Thermal support using a heating pad is recommended.
- Anesthetize the mice by intraperitoneal injection of chloral hydrate (400 mg/kg). Confirm anesthetization by performing a toe pinch. Use of meloxicam SR at induction and buprenorphine upon recovery BID (bis in die, twice a day) for at least 24 h is recommended.
- Shave the hair with a surgical clipper. Fix a mouse in the mouse and neonatal rat adaptor using ear bars and a nose clamp.
NOTE: It is critical to make sure that mouse head is secured at this stage and does not move side-by-side.
- Apply vet ointment on eyes to prevent dryness while under anesthesia.
2. Stereotaxic Surgery for Guide Cannula Implantation
- Set the mouse and neonatal rat adaptor on the stereotaxic apparatus. Make an incision sagittally on the skin over the skull using a scalpel. Wipe off blood and the connective tissue on the skull using a damp cotton swab.
- Determine the coordinate for brain region of interest using brain atlas. Before starting measurements, make sure that midline is straight so that the drill bit can be moved A-P and remain on the midline suture the entire time.
NOTE: This article uses the coordinate (A/P: -3.1 mm, M/L: -2.5 mm, D/V: -1.2 mm, 12 degrees) to target posterior hippocampus.
- Level skull A-P
- Attach a drill on a manipulator of a stereotaxic frame. Lower the drill until it gently touches lambda and record its ventral coordinate. Repeat this procedure for bregma.
NOTE: When the skull is leveled, the vertical measurement of bregma is equal to that of lambda. If not, adjust the height of the nose clamp accordingly. After the skull gets leveled, record the anterior/posterior, lateral coordinate of bregma.
- Level skull left-right
- Move the drill from bregma to the coordinate (Anteroposterior [A/P]: -3.1 mm, mediolateral [M/L]: +2.0 mm), lower the drill to the skull and record the vertical coordinate. Then repeat this procedure for the coordinate (A/P: -3.1 mm, M/L: -2.0 mm).
NOTE: If the skull is leveled, these vertical measurements of two equidistant points from the midline are equal. If not, adjust the height of the ear bars.
- Drill a burr hole carefully at the target coordinate (A/P: -3.1 mm, M/L: -2.5 mm) to implant a guide cannula. If the diameter of a burr hole is not large enough for a guide cannula implantation, drill another hole that overlaps with the first one. Drill another hole on the right (contralateral side) parietal bone and insert a bone screw, which helps to secure dental cement in 1.10 (See Figure 1B).
- Cut a circular locking piece from the back side of lid of a 1.5 mL centrifuge tube by a razor blade and make a ''crown''. This crown is used to prevent dental cement from spreading to the skin. Place it on the skull so that the burr holes made in step 2.5 stay within the circle (See Figure 1).
- Set up a stereotaxic assembly by putting a guide cannula on the shorter arm of a stereotaxic adaptor and fasten it using a cap nut. Set the longer arm of the stereotaxic adaptor on the electrode clamp. Attach it on the manipulator of the stereotaxic apparatus (See Figure 1A).
- Rotate the dorsal-ventral (D-V) stereotax assembly on the manipulator arm by 12 degrees (See Figure 1B). Move the guide cannula to the burr hole made in 1.6. Insert the guide cannula slowly into the brain by lowering it by 1.2 mm.
NOTE: The angle of the probe is specific to the hippocampus; other regions may require other angles or no angle at all. Consult a brain atlas for precise coordinates. Dry the surface of the skull, because if not the cement will not stick and cement cap can become dislodged
- Add the dental cement within the crown to fully cover both the metal part of the guide cannula and the bone screw enough to secure them. Apply additional dental cement if there is a part of skull exposed.
- Wait until dental cement is completely dried (~12-20 min). Remove the stereotaxic adaptor from the electrode clamp. Remove the cap nut and replace the stereotaxic adaptor with a dummy probe and fasten the cap nut.
- Release the mouse from the stereotaxic apparatus and house the mouse alone in an individual cage.
NOTE: The mouse should not be left unattended until it has regained sufficient consciousness to maintain sternal recumbency. Check the mouse daily until the day of microdialysis. The mouse receives nonsteroidal anti-inflammatory drugs (NSAIDs) such as carprofen if it appears to be in pain. Waiting 2 weeks is necessary for sleep-wake studies so the mouse habituates to the new environment, but other types of studies may require shorter recovery periods (e.g., 1-2 days).
3. Microdialysis Setup
- Quality-check of probes: Fill a disposable 1ml syringe with distilled water and connect it to the outlet (shorter port) of a probe using a byton tube. Cover the vent holes with the fingers and depress the syringe plunger gently to infuse water to the probe. Check that water appears from the probe inlet and there is no leakage on the surface of a microdialysis membrane.
- Activation of probes: Submerge the membranes of a probe in ethanol (70-100%) for two seconds. Then, infuse distilled water into the probe again with a syringe again.
- Preparation of perfusion buffer: In order to avoid adhesion of target molecules to the tubings,add BSA by diluting 30% BSA solution to 4% with artificial cerebrospinal fluid (aCSF: 1.3 mM CaCl2, 1.2 mM MgSO4, 3 mM KCl, 0.4 mM KH2PO4, 25 mM NaHCO3, 122 mM NaCl, pH=7.35), which closely matches electrolyte concentration in CSF, immediately prior to use. Filter the perfusion buffer through a syringe filter unit with 0.1 µm pore size.
NOTE: 4% bovine serum albumin (BSA) improves recovery for sticking proteins but can severely limit delivery of compounds, especially for compounds that have high BSA-binding. This is one advantage of using lower concentration of BSA (such as 0.15%) in certain instances. Note that BSA can aggregate very easily when agitated by vortex or stir plate. These aggregates can clog probes or membrane pores. Use caution when preparing a BSA solution to limit this aggregation
- Prepare two separate lines for inlet and outlet (See Figure 1C) and connect both lines with a connection needle. Fill a disposable 3 mL syringe filled with perfusion buffer and connect it with the inlet end of the tubing using a blunt-end needle. Fill the entire tubing with a perfusion buffer by running the syringe pump.
- Stop the syringe pump and replace the connection needle between inlet line and outlet line with an activated microdialysis probe from step 3.3 (See Figure 1C). Before this replacement, put the cap nut on the probe.
- Mount the roller tube in the outlet tubing on the roller pump. Start the syringe pump at 10 µL/min and then the roller pump at the slightly slower flow rate (9.5-9.8 µL/min). Make sure to remove all air bubbles in the entire tubing, which may influence the recovery when they enter the probe.
- Anesthetize the mouse from step 1.12 in the same way as in step 1.2. Put the mouse collar around its neck. Remove the cap nut and dummy probe and slowly insert a microdialysis probe from 3.4 through the guide cannula and fasten the cap nut.
- Place the mouse in the cage connected to a free-moving system and tether the mouse with the collar. Keep running both syringe pump and roller pump at the indicated flow rate in step 3.5 for at least 1 h.
NOTE: To achieve ISF collection from awake animals, this article uses a free-moving system, where the cage itself responds to an animal's movement to keep tubings from twisting. Alternatively, liquid swivels available from various companies can be also used.
- Stop the roller pump first and then the syringe pump. Set the desired flow rate. Run the syringe pump 20% faster than the roller pump.
NOTE: For instance, if you run at 1 µL/min, operate the syringe pump at 1.2 µL/min. Optimal flow rate should be determined empirically for each molecule.
- Collecting ISF samples: place the free end of outlet tubing on the refrigerated fraction collector.
NOTE: Appropriate sample volume varies depending on assays used for analysis.
- After the completion of the experiment, remove the probe. (Anesthetize the mouse as needed.) Handle mouse recovery the same way as in step 2.11. Analyze the collected ISF by methods such as high-performance liquid chromatography (HPLC) or enzyme-linked immunosorbent assay (ELISA).
- To wash the entire tubing after microdialysis, connect inlet and outlet tubings again by replacing the probe with the connection needle and run the diluted bleach in the entire tubing and then flush it with water. Dry and store it for repeated use.
NOTE: Other types of tubings are acceptable, however, BSA in perfusion buffer can clog the tubings with small diameter, thus these tubings are considered as single use. The tubings can be worn down or clogged after multiple uses, so make sure that the flow rate is consistent every time before use.
