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Buccafusco JJ, editor. Methods of Behavior Analysis in Neuroscience. 2nd edition. Boca Raton (FL): CRC Press/Taylor & Francis; 2009.

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Methods of Behavior Analysis in Neuroscience. 2nd edition.

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Chapter 10Contextually Induced Drug Seeking During Protracted Abstinence in Rats

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Over the past half century great strides have been made in the development of useful animal models for the drug abuse triad—self-administration, physical or psychological dependence, and withdrawal. In fact, the compulsion to self-administer cocaine even in the face of adverse consequences is not limited to human beings [1,2]. Practically, it is not that difficult to detoxify a drug addict, but the problem lies in the propensity for former addicts to relapse to drug-seeking behavior, a risk factor that does not appear to decrease in potency over time. Recently there has been an increasing focus on the issue of protracted withdrawal. This feature of drug addiction mirrors classical conditioning in that certain contextual cues or environmental stimuli associated with drug taking can readily initiate a form of withdrawal or craving in addicts that often leads to renewed drug seeking and relapse (for review, see [3–5]). Indeed, both the rat and human share common triggers of relapse, including the drug of abuse itself, stress, and stimuli or the environment conditioned to the drug of abuse [6]. Rodent models of human drug craving and relapse have used paradigms of extinction and reinstatement. Such models have shown predictive validity by demonstrating that clinically effective anti-craving drugs reduce drug-seeking behavior as a component of the model [7].


10.2.1. Animal Subjects

Male Wistar rats (Harlan, Indianapolis, Indiana, USA), weighing 300 to 380 g, are housed and tested in environmentally controlled rooms on a 12/12-hr day/night cycle, and they are maintained on standard rat chow and tap water (unlimited). All current animal protocols are approved by the Augusta Veterans Administration Animal Care and Use Committee. Initially each rat is maintained at about 85% of free-feeding levels for about 5 days. During this time the animals are acclimated to the operant chamber and trained for lever pressing. Once a rat is assigned to a particular operant chamber they are maintained there on a 24-hr basis except during the withdrawal phase (see below).

10.2.2. Use of Food to Shape Responding

Some investigators use food reinforcement to shape animals for lever responding. The advantage is that animals that are poor responders can be weeded out before they are prepared for i.v. self-administration of the drug of interest. The disadvantage is that the investigator must take care to insure that subsequent lever responding, whether during the i.v. self-administration phase or the post-withdrawal phase, is not reflecting the expectation or the habit of obtaining food rewards. Presently we prefer not to shape animals using food reinforcement, but for those that prefer this approach, and for those that plan to study food reinforcement as the sole reinforcing agent, we describe the method below.

We generally train animals on an operant food-reinforcement fixed ratio-1 (FR1) schedule during 2-hr daily sessions. Lever presses are reinforced by the automated delivery of a 45-mg food pellet. The lever is signaled active by the illumination of a stimulus light mounted above the lever. The only time the stimulus light is extinguished is during a post-reward 50-sec timeout period. Training and testing are accomplished in a Coulbourn Instruments (Allentown, Pennsylvania, USA) computer-controlled operant system that includes 16 operant chambers (represents an upgrade from current six-station system) with light cues and retractable levers. Each operant chamber is housed in a sound-attenuated and fan-cooled environmental compartment. Rats that maintain at least 100 responses for three consecutive sessions are surgically prepared for i.v. self-administration sessions as described below. The pellet feeder is removed from the operant chamber (to be replaced by i.v. infusion of morphine), but throughout the remainder of the study, the rats have unlimited access to standard rat chow and water.

10.2.3. Implantation of the i.v Infusion Line

The trained rats are anesthetized with sodium methohexital (65 mg/kg, i.p.) and under aseptic conditions a midline incision is made ventrally over the neck region. The jugular vein is exposed by blunt dissection and a small nick made to allow introduction of a non-thromogenic, softening, vascular implant tubing (Data Sciences, St. Paul, Minnesota, USA), which is filled with dilute heparinized (20 units/mL) sterile saline. The tubing is advanced about 2.5 cm and then it is tied off and secured to surrounding fascia. The tubing is tunneled under the skin to emerge at the nape of the neck. There the tubing is stabilized to a subcutaneous plastic anchor button and fixed to a water-tight swivel cannula mounted at the top of the operant chamber. The swivel is connected to the computer-controlled infusion pump. Two days later the patency of the venous infusion system is tested by the rapid i.v. administration of the short-acting anesthetic agent sodium methohexital (2.0 mg). Intravenous administration of methohexital leads to immediate loss of muscle tone and righting reflex in patent animals. The heparin-saline solution is maintained in the catheter until the start of morphine/cocaine self-administration (third day after surgery). It is important to maintain each rat in its previously assigned operant chamber.

10.2.4. The Morphine Regimen

Rats are permitted to self-administer morphine sulfate according to an FR1 schedule with a 50-sec timeout instituted after each infusion. An illuminated stimulus light signifies the beginning of the session and indicates that the lever is active (one press, one infusion). The light is extinguished and the lever is made inactive (presses elicit no infusion) for 50 sec after a reward is delivered. The pump is set to deliver morphine sulfate over a 5-sec period in 0.165 mL of saline. An experimental cohort usually consists of eight rats (usually provides adequate statistical power). The starting dose of morphine in a single infusion is 0.25 mg/kg. Animals are permitted to self-administer 0.25 mg/kg/infusion over the first three days. During the next three days animals self-administer 0.5 mg/kg/infusion, and during the next four days they self-administer 1.0 mg/kg/infusion. The only interruptions in the 24-hr access schedule are the brief periods needed to replace empty infusion syringes between the changes in drug concentrations. Over the final five days, the morphine levels in the infusion solution are tapered to 0.25 mg/kg/infusion (on days 10, 11, 12, 13, and 14 the morphine concentration is adjusted accordingly to 0.75, 0.75, 0.5, 0.5, and 0.25 mg/kg/infusion, respectively).

10.2.5. The Cocaine Regimen

The procedure for cocaine i.v. self-administration is similar to that for morphine except that the first dose of cocaine to be self-administered is 0.5 mg/kg/infusion, and the timeout period after each infusion is 280 sec. The timeout period of 280 sec is instituted so that the animal cannot self-administer more than 300 doses per day. During the next three days animals will self-administer 1.0 mg/kg/infusion, and during the next four days they will self-administer 2.5 mg/kg/infusion. In practice, the animals will maximally self-administer no more than about 150 doses in 24 hr (Figure 10.1). Over the final five days, the cocaine levels in the infusion solution are tapered to 0.5 mg/kg/infusion (on days 10, 11, 12, 13, and 14 the cocaine concentration is adjusted accordingly to 2.0, 2.0, 0.5, 1.0, and 0.5 mg/kg/infusion, respectively).

FIGURE 10.1. The self-administration of an escalating dose regimen of i.


The self-administration of an escalating dose regimen of i.v. morphine infusion by 21 rats under a contingent FR1 schedule of reinforcement with 50-sec timeouts. Access to the reinforcement response lever was available 24 hr per day. (a) The dose of morphine (more...)

10.2.6. Spontaneous Morphine Withdrawal Syndrome

On the final day of self-administration rats continue to have access to the morphine infusion lever until 1700 hr, at which time the i.v. infusion line is disconnected and plugged. Body weight and core temperature (rectal temperature measured by using a thermistor probe) are recorded (0 hr post-withdrawal) and the animals are placed back into their individual home cages. At 0900 hr on the following day (16 hr post-withdrawal) body weight and core temperature are again measured. Next the rats are placed in a standard open-field environment to assess abstinence signs associated with spontaneous morphine withdrawal by using a standardized withdrawal checklist [8,9]. These symptoms are scored by a “blinded” rater during 30-min observation periods. Withdrawal symptoms include withdrawal body (wet-dog) shakes, escape attempts (attempting to leap out of the cage), writhing, defecation/diarrhea, and chromodacryorrhea (reddish tears). This procedure is initiated three more times at consecutive 2-hr intervals, and the incidences of the scored symptoms over the four observation periods are totaled. In our previous studies, morphine withdrawal symptoms were shown to peak between 12 and 20 hr after withdrawal and they ended by about 100 hr [10,11]. Body weight and core (rectal) temperature are measured after the conclusion of the last observation period (1700 hr). Thereafter, body weight and temperature are measured at 900 hr and at 1700 hr each of the next two days (withdrawal symptoms will no longer need to be recorded), and body weight (which requires the longest period to recover, Figure 10.2) is recorded once daily for the next three days. After the last measurements, the animals are left undisturbed in their individual home cages for the remainder of the 6 wk post-withdrawal period.

FIGURE 10.2. The change in body weight in morphine-dependent rats after discontinuation of self-administration.


The change in body weight in morphine-dependent rats after discontinuation of self-administration.

10.2.7. Reinstatement of Lever Pressing

After the 6-wk post-withdrawal period the subjects are returned to the operant chambers. The condition of the chambers is identical to that during the morphine self-administration phase, and each rat will always be returned to its original chamber. Again rats are allowed to lever press according to the 24-hr access schedule used during morphine or cocaine self-administration, including the (contingent) cue light. The animals have unlimited access to standard rat chow and water. In this case the i.v. line is not reconnected, and lever pressing will not result in a reward. Rats are maintained in the operant chamber for at least seven days.


10.3.1. Things to Prepare Before Surgery

  1. Order male Wistar rats, 300–324 g, from Harland, Indianapolis, Indiana, USA. Allow the animals at least 1 wk to acclimate after shipping.
  2. Connect the PE 20 tubing to the bottom of the swivel. This can be difficult. Insert a small 26-gauge needle in it first to stretch it a little. Then insert a piece of 22-gauge wire. This can help open the tubing. Attach the swivel to the tubing. By doing this before surgery it can be reattached easily to save time.
  3. The surgical instruments are sterilized in a glass bead sterilizer (Fine Scientific Tools, Foster City, California, USA). Before placing the instruments in the sterilizer they should be cleaned and scrubbed in rubbing alcohol. Insert the dry instruments into the beads at 250°C (it requires about 20 min for the sterilizer to reach 250°C) for at least 2 min. Allow the instruments to cool before starting surgery. The sterilizer does not need to be turned off between surgical procedures. Gloves, surgical gown, and mask should be worn during all surgical procedures.
  4. Dilute hospital-grade sodium heparin in sterile normal saline to 20 units/mL.

10.3.2. Installing a Permanent Intravenous Line

One hour before surgery rats are administered carprofen sterile injectable solution purchased in 50 mg/mL vials (Pfizer Animal Health, Exton, Pennsylvania, USA). Carprofen is an analgesic agent administered by subcutaneous injection at a dose of 5 mg/kg. Rats are anesthetized by i.p. injection with methohexital sodium (Brevital) 60 mg/kg. Brevital (purchased as 500 mg of the dry powder in 50 mL vial to provide a working solution of 10 mg/mL) is a short-acting anesthetic and in most cases the entire surgical procedure can be accomplished after a single injection. However, if the animal begins to recover (assess response to toe pinch), an additional 5 mg of Brevital should be administered. The fur should be clipped ventrally over the neck and on the area dorsally over the neck just behind the head after the animal is anesthetized. Next, the exposed skin in the surgical fields should be swabbed with Betadine antiseptic solution. A midline incision is then made in the skin over the ventrolateral aspect of the neck. The incision is about 2-cm long. A small pair of round-tipped scissors can be used to bluntly dissect the fascia without cutting. The right exterior jugular vein will be exposed. Care should be taken not to overly stretch the vein as it will become narrower and more difficult to cannulate. The vein can be isolated from the surrounding fascia by using curved forceps, but care must be taken not to puncture the vein. Place a small forceps under the vein to secure it. Place two pieces of 3–0 black braided suture at the top and bottom of the cleared vessel. Tie a knot on the cranial aspect of the vein to seal it off. Make the venotomy (incision) approximately 5 mm cranial to the site of crossover of the pectoralis major muscle. Before inserting the small animal vascular catheter into the vein, cut a bevel on the end to be inserted. Make sure that the tip is not too sharp. The catheter should be about 20-cm long. Fill the cannula with heparinized saline (20 units/mL). Insert a #7 Dumont tweezer into the vein and lift slightly, i.e., insert the right tip of the forceps into the vein and then grasp the vein. Slowly insert the small animal tubing into the vein for about 1.5 cm. Hold the vein and tubing with the Dumont tweezer until insertion is complete. (If nick in the vein is difficult to see, the vein can be “milked” by gentle rubbing and getting the blood to flow again. Application of saline to the cut end could also help.) Tie the vein and tubing in with the 3–0 suture that was placed there earlier. Both sutures should be tied to the vein to secure the catheter. A subcutaneous tunnel should be created from the neck area to the dorsum. A small incision (about 1.5 mm) is made in the skin just behind the head. Make it a pocket by inserting the scissors and opening them to clear the fascia away. Pass a trochar (point-sharpened 14-gauge stainless steel tubing) between the skin and muscle layers. (Note: Do not pass through the muscle.) around to the back of the neck behind the head and out the small incision. Pass the catheter through the trochar. Remove the stainless steel tubing. Pull the catheter through the anchor button and place the button under the skin. Suture the skin to the button with 3–0 black braided suture. Attach a connector made from 26-gauge stainless steel tubing to the catheter. Next, place a piece of PE 20 tubing filled with heparin saline to the connector. Place a piece of Vardex tubing about 4-in long over the spring support. (This is optional but keeps the rat from chewing into the spring support.) Feed the tubing through the spring support and attach the spring to the button. The tubing from the bottom of the swivel to the top of the animal should be long enough to include a loop in the top before attaching the catheter to the swivel (approximately 13-in long—you can always shorten it by cutting some from the end that joins to the connector). The loop should be small, about 2.5 cm in diameter. The loop absorbs tension on the catheter as the animal moves. If the loop is too large it can get caught on the swivel holder. Note: Coulbourn tethers have attachments at the bottom to fit their rodent harnesses. We do not use the harness and so the attachments are clipped off with a wire cutter. Tethers are cut to 32 cm. With a dull pair of scissors, squeeze between turns on the spring to create an offset. The output end of the swivel is twisted tight into the entry of the offset and the looped PE 20 tubing from the rat connects to the swivel output. This offset allows the rat to move more freely in the Habitest cage. Place a 5-in piece of Tygon micro-bore tubing, 0.2 in inner diameter (id) × 0.6 in outer diameter (od), on the input of the swivel. Fill the swivel and tubing with heparinized saline. Attach the swivel to the spring and then attach the looped tubing to the bottom of the swivel. Fill a 60-mL syringe with 30–40 mL of heparin saline. Insert a blunt 22-gauge needle into the end of the syringe and attach it to 0.2 in id × 0.6 in od Tygon tubing long enough to reach the top of the swivel. Place a 22-gauge stainless steel connector on the end of the tubing. Fill the tubing with heparin saline or drug. Connect it to the tubing from the swivel input. Infuse heparin saline at a rate of about 7.5 mL/day. Place the rat in the Habitest cage. Hook the swivel to the swivel holder. A weight at the back of the balance arm can be moved to make the line taut. Place the water bottle on the cage and place food in bottom of cage. (Note: Place 35 g of rodent chow per day.) Food consumption should be recorded on a daily basis. The Habitest Linc program for controlling the operant aspects of the task and for recording lever responses can be initiated at any time.

10.3.3. Self-Administration Dose Calculations

Morphine Self-Administration

Day 1–30.25 mg/kg/day
Day 4–60.50 mg/kg/day
Day 7–91.00 mg/kg/day

Cocaine Self-Administration

Day 1–30.5 mg/kg/day
Day 4–61.0 mg/kg/day
Day 7–91.5 mg/kg/day

Sample Calculations for Preparing Stock Solution of Drugs for Self- Administration: For a rat with average body weight of 300 g and a required dose of 0.25 mg/kg/infusion:

  • Infusion pump is set to 0.165 mL/infusion
  • Dose (mg/kg/infusion)/0.165 mL/infusion × weight in kg = mg/mL 0.25 mg/kg/infusion × 0.3 kg/0.165 mL/infusion = 0.455 mg/mL (concentration of stock solution)
  • Prepare enough drug solution to fill all infusion syringes.

To calculate daily (24 hr) dose self-administered, divide the dose infused (mg/kg/infusion) by the number of infusions self-administered per 24 hr.

10.3.4. Ordering Information

Small animal vascular catheterData Sciences #277-0011-002
St. Paul, MN 55126-6164 USA
Button for DC95 tethersInstech #DC95BS
Tygon tubing 0.02 × 0.06VWR 63018-044 or Fisher 14-170-15B
Stainless steel tubing and wire
 14-gauge stainless steel tubingSmall Parts HTX-14
 26-gauge stainless steel tubingSmall Parts HTX-26
 22-gauge stainless steel tubingSmall Parts HTX-22
 22-gauge stainless steel wireSmall Parts J-SWX-022
PE20 tubing 0.015 × 0.043VWR 63019-025
Male Wistar rats 300–324 gHarlan, Inc. Indianapolis, IN, USA
Glass bead sterilizerFine Scientific Tools
FST 250 sterilizer
Rodent 22 g swivelBraintree Scientific RS-22G
(Need this or the ones from Coulbourn)www​
 Vardex interbraided tubing 0.25 id × 0.453 odNewage Industries
Southampton, PA 18966 USA
 Utility cutter short-angled blade scissorsFisher Scientific 14-277
 SuperCut iris scissors, straight, 4.5 inBraintree Scientific SC528

10.3.5. Coulbourn Instruments—Required Equipment

DescriptionPart Number
Cue, single high bright light—ratH11-03R
House lightH11-01R
Balance armH-29-01
Panel, blank metal assortmentH-90-00R-M-KT01
Pump, infusion—programmableE73-02
Habitest power base w/lidH01-01
Habitest LincH02-08 (for two rats)
Environmental control boardH03-04
Test cageH10-11R-TC
Non-shock floorH10-11R-TC-NSF
Catheter protection and tethersA73-59R
Rat harness to fit 150–500 g ratA71-21R-350/500
Graphic State software, version 3.03GS3.03
PCI-3 computer interface cardPCI-3-Kit
Computer controller requirementsSys Control 1F P4, 3.2 GHz, 1GB RAM, 250 GB HD, 17-in LCD, DVD-RW (May use Windows XP with PC1 slot and turn-key package)

Note: To self-administer food pellets, a feeder and the wires to connect them to the board are required. There may be other cables and accessories required depending on the chosen system configuration. This info can be obtained from Coulbourn Instruments (


10.4.1. Morphine Self-Administration

These data reflect the trial of different morphine self-administration regimens. Twenty-one rats initially participated in the first experimental series, and each animal was trained to self-administer 0.25 mg/kg doses of morphine. These data are presented in Figure 10.1. Animals generally maintained their level of lever responding (about 100 responses/24 hr) that carried over from the earlier lever training experiments where food pellets provided the response motivation. That the animals transferred this behavior to morphine-reinforced responding was insured by the removal of the food hopper (the space was covered by a flat insert indistinguishable from the surrounding wall), and by the continuous availability of food in the operant chamber. Also, in separate studies in which rats shaped on food rewards were transferred to saline self-administration, the lever responding that carried over extinguished over the same time period [12]. Note that it is possible that rats can learn to self-administer without the use of prior shaping with food reinforcement. This is because with 24-hr access, inadvertent lever responses occur with sufficient frequency to help encourage the behavior. Should you use this approach, you should be prepared to encounter some animals that fail to lever respond sufficiently during the first few days to the extent that they are removed from the study. As indicated in Figure 10.1A, lever responding and the daily dose self-administered increased over the first 3 days. Lever responding became relatively constant over the next 6 days during the self-administration of the 0.5 mg/kg and the 1 mg/kg doses. Though the numbers of animals remaining in the study decreased dramatically over the last 4 days (the self-administration phase was terminated at different times to enhance the variability in the total amount of morphine consumed prior to withdrawal), there was a dramatic decrease in responding when the infusion dose was increased to 2 mg/kg. The level of responding recovered even for the two rats that self-administered 4 mg/kg/infusion. The daily dose of morphine was maintained fairly constantly during the self-administration of the 0.5 mg/kg and 1 mg/kg doses per infusion (Figure 10.1D). This was evident in the observation that after day 6 (the last day of 0.5 mg/kg/infusion), when the dose of morphine was increased to 1 mg/kg/infusion, responding decreased slightly so that the average total dose self-administered could be maintained at about 70 mg/kg/day. This profile of responses, and the amount of morphine self-administered, was again apparent in transitioning from the 1 mg/kg to the 2 mg/kg doses per infusion. Two animals continued to self-administer 4 mg/kg morphine per infusion with additional fall-off in the number of lever responses. By varying the number of days of self-administration, and allowing some animals to self-administer high concentrations of morphine, we were able to obtain a broad spectrum of total morphine doses as well as the associated total number of lever responses. For morphine self-administration, we allow rats to self-administer only up to 1 mg/kg/infusion. If the animals maintain a good level of responding (> 70 responses per day) they will become dependent on the drug after about 4–5 days self-administering this dose.

At the completion of the self-administration phase of the study, animals are returned to their home cages and allowed to undergo withdrawal. Figure 10.2 shows the change in body weight as a function of time after withdrawal. There was a characteristic sharp decrease in body weights averaging 16.2 g measured 36 hr after withdrawal. Thereafter, body weight gradually increased to near control levels by 84 hr post-withdrawal. The animals continued to gain weight through the last observation period at 6 days after withdrawal. More importantly, the withdrawal-associated decrease in body weight was shown to be linearly related to the total dose of morphine self-administered [12]. These data therefore relate the quantity of morphine consumed during the dependence phase to magnitude of the expression of this withdrawal symptom. In general, other withdrawal symptoms were not as dramatic or intense as with opiate antagonist-precipitated withdrawal, and the most prevalent of the visually observed symptoms was withdrawal body shakes. Of the other symptoms, defecation and diarrhea were noted most often, though these are not necessarily characteristic withdrawal symptoms.

10.4.2. Context-Induced Post-Withdrawal Lever Responding

After completing the 6-wk withdrawal period, rats were placed back into their original operant environment and allowed to make lever responses with no reward consequences (even though the illuminated stimulus light continued to signal an active lever). These data are presented in Figure 10.3. Returning the animal to the operant chamber resulted in a doubling of the average 24-hr lever response rate that was measured on the last day of the self-administration schedule. Average responding decreased over the next two days, but it rebounded during test days 4 and 5. By day 7 the response rate extinguished to that measured during the last day of self-administration. The differences among the means for each day of testing relative to the pre-withdrawal level of responding was statistically significant (P < 0.01).

FIGURE 10.3. Context-induced reinstatement of lever responding 6 wk after discontinuation of morphine self-administration in dependent rats.


Context-induced reinstatement of lever responding 6 wk after discontinuation of morphine self-administration in dependent rats. The number of lever responses (per 24 hr) during the 7-day lever reinstatement period with lever responding on the last day (more...)


10.5.1. The Drug Abuse Cycle and Pharmacological Intervention

Animals that self-administered morphine according to our standard protocol were shown to be physically dependent on morphine. This was evidenced by the appearance of characteristic symptoms of abstinence, and more specifically by the precipitous decrease in body weight that was maximal on the second day after morphine withdrawal. After monitoring body weight for 6 days, the rats were returned to their home cages to complete the 6-wk protracted abstinence period. Harris and Aston-Jones [13] reported that the preference for a morphine-paired environment in formerly dependent rats was maintained from 2–5 wk post-withdrawal. In fact, increased drug-seeking behavior after a protracted term of abstinence was noted for several drugs of abuse [14]. Likewise, in the present study, returning formerly dependent animals to the self-administration environment resulted in an initial doubling of the 24-hr response rate as compared with the last few days of self-administration. Extinction of enhanced lever-pressing activity came slowly, but self-administration levels were attained by day 7.

The 24-hr access model described here provides a step closer in relevance to real-world conditions than studies that rely on one- or two-hour daily test sessions that often occur during the animals’ sleep cycle. Another important feature of the paradigm is that treatment interventions can be studied within the same model for each component of the drug abuse cycle: self-administration, dependence, withdrawal, protracted withdrawal, and renewed drug-seeking behavior. Figure 10.4 illustrates the days of administration for each treatment intervention (arrowheads) across the three regimens. The regimen outlined in the uppermost graph of Figure 10.4 permits the evaluation of treatment intervention administered during the self-administration phase on subsequent acute and protracted withdrawal behaviors. Note that the first treatment intervention occurs on day 9—the last day of the highest concentration self-administered. This paradigm is designed to examine the direct effect of treatment intervention on self-administration behavior. At the conclusion of day 10, self-administration is maintained over the following 5 days. Treatment intervention can continue to be administered up until the start of withdrawal at the end of day 14. This paradigm of interdiction after the development of physical dependence is more clinically relevant than beginning the treatment intervention simultaneously with the start of self-administration, though the latter paradigm can be used to assess the effects of treatment interventions designed to inhibit self-administration behavior. When treatment intervention is initiated at the start of the self-administration period, if lever responding is decreased, the expression of downstream withdrawal symptoms will automatically be reduced. Thus the regimen outlined in the upper graph of Figure 10.4 circumvents this limitation.

FIGURE 10.4. The phases of the drug-abuse cycle as provided by the 24-hr access model for self-administration.


The phases of the drug-abuse cycle as provided by the 24-hr access model for self-administration. Arrows indicate the days in which a treatment intervention is administered.

The regimen outlined in the middle graph of Figure 10.4 permits the evaluation of treatment intervention during the acute withdrawal period on acute and protracted withdrawal behaviors. The regimen outlined in lowermost graph of Figure 10.4 permits the evaluation of treatment intervention during the protracted withdrawal period on protracted withdrawal behavior (contextually induced lever responding). Therefore, insight can be gained into the specificity of each treatment intervention on the component of the drug abuse cycle, and information can be obtained regarding the role of each preceding component on the expression of the subsequent components of the cycle.


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Copyright © 2009, Taylor & Francis Group, LLC.
Bookshelf ID: NBK5230PMID: 21204337


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