The protocol overview begins with stereotaxic surgery to administer the combinatorial viral vector approach, using retro-Cre and inhibitory DREADDs, to reduce the excitability of prelimbic (PL) medial prefrontal cortex (mPFC) neurons projecting to the nucleus accumbens (NAc). After 4 weeks of viral expression, surgeries of intrajugular catheter implantation and minipump subcutaneous insertion are performed. Following surgical recovery, operant conditioning maintained by the cannabinoid full agonist WIN 55,212-2 is established, finalizing with the animals’ categorization into addicted and non-addicted mice and the characterization of phenotypic vulnerability traits. Subsequently, extinction and cue-induced reinstatement are tested to obtain parameters related to craving (Figure 2).

A. Stereotaxic surgery: DREADD approach with viral vector microinjection

1. Prepare adeno-associated viral vectors: AAV-hM4Di-DREADD [AAV8-hSyn-DIO-hM4D(Gi)-mCherry, 1.21 × 10¹³ gc/mL] and AAV-retrograde-Cre-EBFP (AAV pmSyn1-EBFP-Cre, 8.2 × 10¹² gc/mL).

2. Anesthetize mice by intraperitoneal injection (0.2 mL/10 g) of ketamine hydrochloride (75 mg/kg) and medetomidine hydrochloride (1 mg/kg) in 0.9% sterile saline, and place them in a stereotaxic frame for intracranial adeno-associated virus (AAV) injections.

3. Perform bilateral microinjections using a 26-gauge guide cannula and a 33-gauge internal injection cannula connected via PE-20 tubing to a 10-μL microsyringe; monitor infusion by displacement of an air bubble in the tubing.

4. For PL–NAc pathway inhibition, inject 0.2 μL/site of AAV-hM4Di-DREADD into PL at 0.05 μL/min and 0.4 μL/site of AAV-retrograde-Cre-EBFP into NAc core at 0.10 μL/min. Leave the cannula in place for 10 min post-infusion to allow diffusion, then withdraw slowly while maintaining body temperature at 35 °C on a heating pad; target coordinates (Paxinos and Franklin, 1997): PL, AP +2.10 mm, L ±0.3 mm, DV -2.3 mm; NAc core, AP +1.94 mm, L ±1.0 mm, DV -4.6 mm.

A1. Stereotaxic surgery (Figure 3)

Note: Sterilize the surgical material with a hot bead sterilizer before each animal surgery.

1. Injection cannulas circuit preparation.

a. Remove the microsyringe plunger (10 μL).

b. Attach the polyethylene tubing (50 cm) to the microsyringe.

c. Inject distilled water through the tubing, using a syringe, filling the tubing and the microsyringe. Verify that distilled water flows out through the back of the microsyringe.

d. Insert half of the plunger into the microsyringe.

e. Connect one of the sides of the bilateral internal cannula (33 gauge) to the tubing. Verify that the circuit is assembled correctly by pressing the microsyringe plunger and observing that water flows from the cannula without leakage.

f. Repeat these steps with a second microsyringe, which must be attached to the other side of the same internal cannula as the first microsyringe.

g. Place the microsyringes in the microinfusion pump.

h. Aspirate 1 μL of air into the tubing to form a bubble and monitor the microinjections.

i. Aspirate 4 μL of the viral vector (previously defrosted with ice).

2. Stereotaxic apparatus preparation.

a. Place and fix the bilateral guide cannula in the metal grip of the stereotaxic holder.

b. Insert the bilateral internal cannulae, attached to the tubing, into the bilateral guide cannulae. The bilateral guide cannula is used to keep the internal cannula straight and well-fitted into the holder, and to determine Bregma coordinates before the AAV is loaded. Note that only the internal cannula, filled with the viral vector, penetrates the brain.

3. Mouse preparation.

a. Anesthetize the mouse with a mixture of ketamine hydrochloride and medetomidine hydrochloride dissolved in sterile 0.9% physiological saline. Administer intraperitoneally (75 mg/kg and 1 mg/kg of body weight, respectively).

b. Shave the mouse’s head.

c. Place the mouse in the stereotaxic apparatus. Check that the head is fixed using the ears bars and the nose clamp.

d. Place a heating pad below the mouse to maintain proper body temperature during the surgery.

4. Stereotaxic surgery.

a. Perform a vertical cut in the middle of the head with a scalpel.

b. Scratch the skull carefully to visualize the bregma accurately.

c. Place the cannulas above the bregma and annotate the bregma coordinates.

d. Calculate medial-lateral, dorsal-ventral, and anteroposterior coordinates of your target area where the injection has to be performed.

e. Move the cannulas to the specific injection point by adjusting the stereotaxic apparatus using the medial-lateral, dorsal-ventral, and anterior-posterior readings.

f. Mark the injection point on the skull using a pencil.

g. Following the pencil mark, drill holes into the skull using a manual drill.

h. Clean the injection area with physiological saline.

i. Insert the bilateral cannula into the holes and readjust the dorsal-ventral coordinate.

j. Place the bilateral cannulas precisely at the calculated coordinates.

k. Mark the two sides of the air bubble on the tubing to monitor the injection.

l. Perform viral vector injection.

i) Microinfusion pump parameters [example for prelimbic (PL) and nucleus accumbens core (NAc core) areas]: Target volume: PL 0.2 μL; NAc core 0.4 μL. Infusion rate: PL 0.05 μL/min; NAc core 0.1 μL/min. Infusion time: 4 min.

ii) Leave the cannula in place for 10 min after infusion to prevent reflux.

iii) Withdraw the cannula slowly for 10 min.

m. Clean the injection area with physiological saline.

n. Sew the incision with suture thread.

o. Apply blastoestimulina on the sutured incision to promote proper healing.

p. Inject subcutaneously atipamezole hydrochloride (2.5 mg/kg of body weight) dissolved in sterile 0.9% physiological saline to reverse the anesthesia.

q. Inject intraperitoneally gentamicin (1 mg/kg of body weight) dissolved in sterile 0.9% physiological saline.

r. Inject meloxicam subcutaneously (2 mg/kg of body weight) dissolved in sterile 0.9% physiological saline.

s. Inject glucose serum (0.8 mL) subcutaneously.

t. Place the animal on a heating pad until the animal is awake.

u. During the following 3 post-surgery days, check the mouse and inject 0.8 mL of glucose serum every day.

5. Cleaning.

a. Clean the tubing and cannulas with ethanol, then rinse with distilled water and air-dry.

b. Clean all the surgical material with soap and water.

Note: After bilateral intracranial injection, 4 weeks are required for proper expression of the viral vector.

Note: Frequently verify the correct displacement of the air bubble within the polyethylene tube to prevent injection errors caused by cannula obturation or tube fugues.

B. Jugular vein catheterization and osmotic minipump implantation

Note: Jugular vein catheterization and minipump implantation procedures are performed after 4 weeks of viral expression, as shown in the experimental schedule (Figure 2).

1. Anesthetize mice by intraperitoneal injection (0.2 mL/10 g body weight) of ketamine hydrochloride (75 mg/kg) and medetomidine hydrochloride (1 mg/kg) dissolved in 0.9% sterile physiological saline.

2. Implant indwelling intravenous silicone catheters in the right jugular vein as described previously [14]: adapt 6-cm silicone tubing (inner diameter 0.3 mm, outer diameter 0.6 mm) to a 22-gauge steel cannula bent at a right angle and embedded in a dental cement disk with nylon mesh underneath. The step-by-step procedure for jugular vein catheterization (Figure 4) is as follows:

a. Anesthetize the mouse with a ketamine/medetomidine solution (0.2 mL/10 g, intraperitoneal).

b. Apply ophthalmic gel (e.g., xylin) to both eyes to prevent corneal drying during surgery.

c. Shave the chest and back, then disinfect the skin by wiping with 70% ethanol followed by iodine solution over the same area.

d. Identify the interscapular region and make a 1–2 cm skin incision parallel to the spine. Using scissors, gently separate the skin from the underlying musculature to create a subcutaneous pocket for the catheter.

e. With curved forceps, further blunt-dissect the subcutaneous space toward the right shoulder, where the external part of the cannula of the silicon tube will be positioned.

f. Turn the animal onto its back and locate the heartbeat. On the upper right thoracic region, grasp the skin overlying the expected course of the right jugular vein with forceps, make a small skin incision, and bluntly dissect toward the subcutaneous tunnel previously created from the back.

g. Pass the forceps from the thoracic incision through the subcutaneous tunnel to the dorsal incision and pull the catheter through so that it lies over the right shoulder. Confirm catheter integrity by gently flushing with heparinized saline (0.1% heparin in saline).

h. Expose the right jugular vein by carefully removing overlying fat and muscle fibers.

i. Once the vein is visible, insert fine pointed forceps (e.g., Dumont) beneath the vein from the right side, and a second pair from the left side; rotate the forceps to free the vein from surrounding tissue.

j. Pass a bent silk suture under the vein from left to right, then cut the loop to obtain two separate threads. Tie the caudal (lower) thread loosely around the limb to keep it in place. With the cranial (upper) thread, apply a firm single knot to transiently occlude blood flow.

k. Choose a straight segment of the vein for catheterization and puncture it with a 20 G 1½ needle. Remove the needle and insert the catheter guide fully through the needle tract into the lumen (Figure 4A).

l. Advance the silastic catheter tubing into the jugular vein until the silicone bead reaches the vessel entry. Verify correct placement by observing blood backflow into the catheter.

m. Tighten the lower ligature to secure the catheter and slowly inject 0.2 mL of heparinized saline (0.1% heparin) to fill the catheter with fluid (Figure 4B).

n. Tie additional knots with both upper and lower silk threads to firmly anchor the catheter, then add two extra knots on the left and two on the right to stabilize the assembly.

o. Ensure that approximately 1.5 cm of catheter tubing lies within the right jugular vein and that the remaining tubing runs subcutaneously to the dorsal cannula exiting at the midscapular region.

p. Close the thoracic skin incision with sutures and apply a thin layer of antibiotic ointment (e.g., Bactroban).

q. Infuse 0.025 mL heparinized solution (0.3 mg/kg) through the catheter and cap the external port to prevent air entry. Position the tulle of the catheter under the dorsal skin, suture the skin over it to secure the tubing, and close firmly (Figure 4C).

r. Apply topical healing ointment (e.g., Blastoestimulina) to all sutured areas. Administer atipamezole (2 mg/kg, i.p.) to reverse anesthesia, followed by gentamicin (1 mg/kg, s.c.), meloxicam (2 mg/kg, s.c.), and 0.9 mL of glucose-containing saline subcutaneously for hydration (Figure 4B).

s. Allow animals to recover for 3 days before initiating intravenous self-administration sessions.

t. Evaluate catheter patency after the final self-administration session and whenever behavior deviates markedly from previous performance. Assess patency by injecting thiopental (Tiobarbital, 0.5 g) through the catheter at 1.25 mg/kg; if clear anesthetic signs are not evident within 3 s, exclude the mouse from further analysis. For a 12-day protocol, catheter patency should be maintained in >80% of animals. Loss of catheter patency is minimized through careful handling and delicate manipulation of the catheter assembly throughout the experiment. Indicators of catheter blockage or failure include a sudden loss of infusion-contingent responding, resistance during catheter flushing, backflow or leakage at the access port, and subcutaneous swelling observed during infusion. Animals showing evidence of catheter malfunction should be excluded from further testing and subsequent analyses. Final confirmation of catheter patency is performed at the end of the experiment using a thiopental test; however, routine monitoring throughout the study should be conducted to detect potential issues early and prevent data loss.

3. Insert the catheter tubing 1.1 cm into the right jugular vein and secure with a suture; tunnel the remaining tubing subcutaneously to the cannula, exiting at the midscapular region. Suture all incisions and coat with local analgesic (Blastoestimulina).

4. Provide post-surgical care consisting of intraperitoneal antibiotic (gentamicin, 1 mg/kg), subcutaneous analgesic (mixture of glucose serum and meloxicam, 2 mg/kg), and subcutaneous anesthesia reverser atipamezole hydrochloride (2.5 mg/kg), all dissolved in 0.9% sterile physiological saline. Allow mice to recover for 3 days with follow-up analgesia before starting self-administration sessions.

5. At the end of the self-administration sequence, assess intravenous catheter patency by thiopental sodium test; remove any mouse from the study if clear signs of anesthesia are not observed immediately after injection.

6. Prepare Alzet osmotic minipumps filled with clozapine-N-oxide (CNO; 5 mg/mL in 0.9% saline) or vehicle, and implant them subcutaneously in the lower back during jugular vein catheterization surgery to deliver CNO at 0.25 μL/h for 15 days [15]. The step-by-step osmotic minipump implantation protocol (Figure 5) is as follows:

a. Anesthetize the mouse with isoflurane and fix its front and hind legs with tape.

b. Shave the mouse’s lower back.

c. Clean the shaved lower back with ethanol.

d. Make a horizontal cut (1.5 cm), taking as a reference the beginning of the hind legs.

e. Separate the skin from the back muscle, introducing blunt-tipped scissors through the cut.

f. Introduce the osmotic minipump previously filled with CNO (diluted in 0.9% sterile saline; 5 mg/mL) or saline subcutaneously in the middle of the back. The pump cap is in the direction of the animal’s head.

g. Close the cut with surgical clips (approximately 4).

h. Apply iodine to the area.

Note: Correct subcutaneous implantation of the osmotic minipump and meticulous wound closure were performed to minimize the risk of pump displacement, extrusion, or postoperative infection. Under these conditions, incorrect implantation was rarely observed. Animals with suspected pump displacement were excluded from subsequent behavioral testing, and correct pump positioning was verified post-mortem when necessary.

Note: Check the correct placement of the minipump on the following days after surgery. If necessary, perform a smooth massage at the minipump placement site to facilitate relocation and prevent skin wounds.

C. WIN 55,212-2 self-administration training

1. Apply an operant drug self-administration procedure based on previously established paradigms [7–10,12,16,17].

2. To minimize acute behavioral suppression and potential aversive or dysphoric-like effects associated with the first exposure to WIN 55,212-2 (e.g., hypolocomotion and anxiety-like responses), mice receive an intraperitoneal injection of WIN 55,212-2 (0.1 mg/kg) 24 h before the first self-administration session [12,18].

3. Train mice to self-administer WIN 55,212-2 intravenously under a fixed-ratio (FR) 1 schedule for 5 consecutive daily sessions, then switch to an FR2 schedule for an additional 5 sessions; conduct all sessions at the same time each day.

4. Start each session with an automatic priming infusion of WIN 55,212-2 via the catheter, followed by two 55-min active periods separated by a 15-min drug-free interval (total session length, 125 min) [11,12,16]. Before connecting the mouse, we press the infusion pump button to produce the sound as a conditioned stimulus.

5. Signal session onset by turning on the house light for 3 s; present cue lights and infusion pump noise as conditioned stimuli during each drug delivery. Implement a 10-s time-out after each infusion, during which the cue light is off, and responses at the active nose poke do not yield drug. Record all responses at active and inactive holes, including those during time-outs.

6. During the drug-free period, deliver neither drug nor cues, and indicate this phase by activating the house light. Terminate the session when either 50 reinforcers have been delivered or 125 min have elapsed, whichever occurs first.

7. Define acquisition of self-administration when all three criteria are met: (i) response rates show at least 80% stability across three consecutive sessions (variance ≤ 20%), (ii) ≥75% of total responses occur at the active hole, and (iii) mice obtain a minimum of five reinforcers per session. Return mice to their home cages after each session. Animals are assigned to home cages in the same room as the operant chambers.

C1. WIN 55,212-2 self-administration training (Figure 6)

1. Twenty-four hours before operant training, an intraperitoneal injection of WIN 55,212-2 (0.1 mg/kg) is performed. Operant conditioning is performed during viral vector expression, which is chronically activated by CNO.

2. Sessions 1–5 (FR1): During these sessions, operant conditioning under the FR1 schedule of reinforcement is applied.

3. Sessions 5–7 (FR2): During these sessions, operant conditioning under the FR2 schedule of reinforcement is performed.

4. Sessions 8–10 (FR2): FR2 sessions to evaluate the criterion of persistence of response and the phenotypic traits of impulsivity and sensitivity to reward.

5. Session 11: Progressive ratio test to evaluate the motivation criteria.

6. Session 12 (FR2): FR2 session before shock test.

7. Session 13: Shock test to evaluate the compulsivity criterion.

8. Sessions 14–23: The extinction phase is applied to evaluate the first parameter related to craving by removing the cue-light, the infusion of WIN 55, 212-2, and the pump noise.

9. Session 24: Cue-induced cannabinoid-seeking test to evaluate the second parameter related to craving.

Note: In cases of failure to acquire self-administration, verify that the operant chamber is functioning properly, ensure that the infusion system is unobstructed, confirm that the animal is correctly connected to the infusion line, and check that the tubing length permits free and unrestricted movement within the chamber. It is also important to assess whether excessive anxiety or stress may be interfering with task engagement; if so, additional habituation and gentle handling may help reduce potential confounding factors. In cases of excessive inactive responding, confirm correct nose-poke functioning and accurate response registration. These measures facilitate distinguishing technical malfunction from behavioral factors and improve protocol reproducibility.

D. Three addiction criteria

1. At the end of training, evaluate addictive-like behavior using three DSM-5–based criteria (persistence, motivation, compulsivity) (Figure 7) to compute an addiction score [6,17,19].

2. Persistence of responding (drug seeking): Quantify the number of non-reinforced active nose pokes emitted during the 15-min drug-free period on each of the 3 days preceding the progressive ratio (PR) session.

3. Motivation: Assess motivation for the drug using a PR schedule in which the response requirement for each successive infusion follows the series 1, 2, 3, 5, 12, 21, 33, 51, 75, 90, 120, 155, 180, 225, 260, 300, 350, 410, 465, 540, 630, 730, 850, 1000, 1200, 1500, 1800, 2100, 2400, 2700, 3000, 3400, 3800, 4200, 4600, 5000, and 5500; define the breakpoint (motivation value) as the last ratio completed, with the PR session lasting up to 4 h or ending earlier if mice cease to respond at all nose pokes for 1 h.

4. Compulsivity (resistance to punishment): After a stabilizing FR2 self-administration session following the PR test, place mice in a different operant box and run a 50-min FR2 session in which each infusion is paired with foot-shock punishment; after the first active response, deliver a foot shock (0.18 mA, 2 s), and after the second response, deliver both foot shock and drug plus cue light. If the second response does not occur within 1 min of the first, restart the sequence and use the total number of shocks obtained as the compulsive-like behavior measure. Prior to punishment testing, footshock intensity was calibrated and maintained constant across sessions to ensure consistent aversive contingencies. Excessive shock intensity may produce nonspecific suppression of responding, whereas insufficient intensity may fail to elicit measurable punishment sensitivity. Shock calibration and grid cleanliness were verified before the shock session to rule out technical malfunctions; particular attention was paid to thoroughly cleaning the grid floor to ensure optimal electrical conductivity.

E. Establishment of mouse subpopulations

1. After completing the three behavioral tests, classify mice as addicted or non-addicted according to the number of addiction-like criteria fulfilled.

2. Define a mouse as positive for a given addiction-like criterion when its behavioral score is at or above the 75th percentile of the saline control distribution.

3. Consider mice meeting 2 or 3 criteria as addicted and categorize them as vulnerable.

4. Consider mice meeting 0 or 1 criterion as non-addicted and categorize them as resilient.

F. Behavioral tests to evaluate addiction-like phenotypic traits

1. Evaluate two additional phenotypic traits as markers of vulnerability to addiction: impulsivity and sensitivity to reward (Figure 8).

2. Impulsivity: Quantify impulsivity-like behavior as the number of non-reinforced active responses emitted during the 10-s time-out period following each WIN 55,212-2 infusion, using the 3 days preceding the PR test for this measure (higher counts indicate greater difficulty in inhibiting responding once initiated).

3. Sensitivity to reward: Assess sensitivity to reward as the total number of reinforcers earned during 2-h sessions across the last three consecutive FR2 self-administration sessions maintained by WIN 55,212-2, where higher reinforcer counts reflect greater reward sensitivity.

G. Extinction and parameters related to craving

1. Include in the extinction phase only mice with patent catheters that satisfied all acquisition criteria. After thiopental testing, allow mice to rest for 1 day and assess locomotor activity in a 2-h session using individual activity boxes (10.8 × 20.3 × 18.6 cm) equipped with infrared sensors for horizontal activity and an infrared plane for rearings.

2. For extinction, run 2-h daily sessions for 10 consecutive days in the same operant chambers used for self-administration, during which neither WIN 55,212-2 priming infusions nor response-contingent WIN 55,212-2 infusions or associated cues are delivered after active nose pokes. Define extinction as performing active responding <35% of the mean active responses previously done during the last 3 days of WIN 55,212-2 self-administration, maintained for 3 consecutive extinction sessions; only mice meeting this criterion proceed to craving-related tests (Figure 9).

3. Resistance to extinction (craving-related measure 1): During the first 2-h extinction session, record the total number of active nose pokes; higher values indicate greater resistance to extinction and increased drug seeking when access to WIN 55,212-2 is prevented.

4. Drug-seeking behavior (cue-induced reinstatement, craving-related measure 2): The day after extinction criterion is reached, perform a single reinstatement session in the same chamber under conditions identical to acquisition, except that active responses are never reinforced by drug; conduct a 120-min FR2 session in which each active nose poke produces all conditioned cues (cue light, pump noise, priming-injection light) without WIN 55,212-2 delivery [16].

H. Histology verification

1. Verify viral expression.

a. Perfuse animals using intracardiac perfusion with 4% paraformaldehyde (PFA) in 0.1 M Na₂HPO₄/0.1 M NaH₂PO₄ buffer (PB), pH 7.5, delivered with a peristaltic pump at 30 mL per min for 2 min. Animals are previously profoundly anesthetized by intraperitoneal injection (0.2 mL/10 g of body weight) of a mixture of ketamine/medetomidine.

b. Extract the brains and post-fix with 4% PFA for 24 h. Transfer them to a solution of 30% sucrose at 4 °C.

c. Make coronal sections (30 μm) using a freezing microtome and store them in a 5% sucrose solution at 4 °C until the immunofluorescence study.

d. Perform an immunofluorescence study using specific antibodies for your viral vector fluorescent reporter used.

e. Visualize the stained sections of the brain with a confocal microscope to evaluate histological verification of correct injection placement.

f. Make representative immunofluorescence pictures (Figure 10).