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Jugular Vein Catheter Design and Cocaine Self-Administration Using Mice: A Comprehensive Method

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Official websites use. Share sensitive information only on official, secure websites. This article was submitted to Motivation and Reward, a section of the journal Frontiers in Behavioral Neuroscience. The use, distribution or reproduction in other forums is permitted, provided the original author s and the copyright owner s are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms. Intravenous self-administration IVSA is a behavioral method of voluntary drug intake in animal models which is used to study the reinforcing effects of drugs of abuse. It is considered to have greater face validity in the study of substance use and abuse than other assays, and thus, allows for valuable insight into the neurobiological basis of addiction, and the development of substance abuse disorders. The technique typically involves surgically inserting a catheter into the jugular vein, which enables the infusion of drug solution after the performance of a desired operant behavior. Two nose- poke ports or levers are offered as manipulanda and are randomly assigned as active reinforced or inactive non-reinforced to allow for the examination of discrimination in the assessment of learning. Here, we describe our methodological approach to this assay in a mouse model, including construction and surgical implantation of a jugular vein catheter, set up of operant chambers, and considerations during each phase of the operant task. Keywords: intravenous, drug, self-administration, catheter, operant conditioning, addiction, mouse, jugular vein. Drug intravenous self-administration IVSA is a behavioral test that has become widely used and accepted in the study of the reinforcing effects of drugs of abuse using animal models, including rats, non-human primates, cats, and mice. It offers valuable insight into the neurobiological basis of addiction by mimicking real-world behaviors that are associated with recurrent drug use during the development and maintenance of substance abuse disorders. Drug IVSA involves surgically inserting an intravenous catheter, typically into the jugular vein, and uses the principles of operant conditioning, such that the subject learns to perform a desired behavior to receive drug delivery Panlilio and Goldberg, Typically, the drug IVSA test is conducted in operant chambers that offer two identical manipulanda—items in the conditioning chamber that can be manipulated physically—such as nose poke ports or levers. Manipulanda are assigned as active reinforced and inactive non-reinforced to allow both for the analysis of port discrimination in the assessment of learning, as well as for general activity levels. A reinforcement schedule allows the experimenter to modify task requirements for the type of research being conducted Platt and Rowlett, Since IVSA allows for volitional control of intake, it is considered to have higher face validity for addictive behavior than other types of tests, and has been used, for example, to verify that nicotine mediates reinforcement via activity at nicotinic acetylcholine receptors Fowler and Kenny, An alternative method is oral self-administration, which can be conducted in both two-bottle choice paradigms, as well as operant paradigms; however, there are trade-offs to consider for the two methods refer to Table 1. Advantages of the IVSA method include allowing the experimenter to remove possible confounding variables and accurate quantification of the amount of drug consumed. Along with increased accuracy, the IVSA allows for a rapid increase in drug levels in the blood and brain Kmiotek et al. The IVSA is also the gold-standard test for assessing the addictive propensity of novel compounds, and the significance of prior drug exposure can be investigated. In addition, infusions can be paired with discriminative cues to allow for later evaluation of drug-seeking behaviors following re-exposure to a reinforcer-paired cue Thomsen and Caine, , which is often used as a model of relapse Galaj et al. Abstinence, or time without access to a drug, causes a deprivation effect and can be used to assess drug-seeking when no drug is available or changes in administration upon renewed drug access. Using the drug IV self-administration assay, many factors, both intrinsic and extrinsic, have been shown to influence voluntary drug-taking. Intrinsic factors, such as exploratory and risk-taking behaviors in mice, are associated with an increased propensity to self-administer cocaine Dickson et al. Sex is also an important factor, since female rats self-administer more nicotine, for example, than male rats Galankin et al. Extrinsic factors, such as social learning and environment, also influence the tendency of an animal to self-administer drugs. A study in which rats had to drink a saccharin solution to get infusions of nicotine revealed that interaction with another rat drinking the solution increased the likelihood of stable drug-taking behavior Wang et al. There are mixed findings on the impact of diet, with studies showing that high-energy diets either do not change Bruggeman et al. The IVSA has been shown to be reduced following exercise, with rats reducing methamphetamine intake after just 1 day of access to an exercise wheel Aarde et al. The acquisition of the drug IVSA in rats may also be elevated by higher ambient temperature Aarde et al. In comparison to basic laboratory animal housing, enrichments, such as toys, obstacles, and running wheels decrease the drug self-administration Ewing and Ranaldi, These findings have broad implications for the types of questions that can be answered using this important behavioral technique. Rodents are often used in IVSA studies, but there exists a historical preference for using rats over mice, likely due to feasibility refer to Table 2 for a comparison of the two species. In rats, the earliest veinous catheters for drug delivery emerged by the early s Slusher and Browning, ; Weeks, , while the first mouse catheters were described nearly 20 years later Barr et al. However, self-administration of substances in mice first relied on tail-vein injection without catheters Criswell, ; Criswell and Ridings, , and mouse self-administration using chronic indwelling catheters was only described later Carney et al. However, with the development of new and smaller materials and model-related motives for using mice to test the role of genetics in substance abuse, more labs are attempting mouse IVSA. In the current methodological description, founded originally on methods developed by Thomsen and Caine , we focus on cocaine IVSA using a mouse to model and measure the addiction-related behavior. Our primary goals in this manuscript are to increase the transparency of methodological choices in mouse IVSA, and likewise, increase the accessibility of the technique. We describe the surgical procedure we use and demonstrate and explain the importance of different phases of the behavioral test, including acquisition, extinction, measurement of a dose-response curve, and increasing the cost. We also discuss other methodological approaches that may be selected for different purposes. Details and images of an assembled catheter can be seen in Figure 1 , and tips for troubleshooting surgery and behavioral testing can be found in Table 4. Tubing-attached syringes will be used for checking catheter patency, surgical drug delivery, and catheter flushing, so it may be useful to prepare multiple syringes. Catheter design and care. A Set-up for attaching tubing to syringe for surgical drug delivery, catheter flushing, and checking catheter patency. B Descriptions of catheter components. C Custom acrylic base mold, in open position, with completed catheter in place to demonstrate mold placement. D Picture of assembled catheter. E Underside of assembled catheter. Allow to cure overnight. Glue must be fully cured for further steps. Make sure to perform any leak checks over the kimwipe so that the source of any leaks can be identified. Use a tubing-attached syringe to push distilled water through the partially assembled catheter. If there are no leaks, release the finger closing the tube, and ensure that liquid can flow through freely. Catheters can be removed from the hood after the curing step. Dip a cotton-tipped applicator in mold release and apply to the inside of each receptacle in the catheter base mold. Close the catheter base mold and tighten it with a screw. Be careful not to over or underfill. Place the mesh all the way down against the surface of the mold so that it touches the BCA mixture. Make sure that the mesh is centered around the cannula base. Make sure that the exposed surface of the BCA mixture is smooth. Use a permanent marker to mark the spot. Allow the silicone to cure overnight. Clamp the melted end quickly with another pair of hemostats to seal. Use a file or sandpaper to extend the bevel further up the shaft, leaving the needlepoint intact. Aim to remove about half the diameter of the shaft for 2 cm in length. Store at room temperature. Keep the solution sterile and make 5 mL of aliquots to be kept at room temperature. Store at room temperature in accordance with the expiration date on the label. An example of an operant chamber and details for set-up can be seen in Figure 2. One option is described in Figure 3. Example: 0. Operant box set-up. A An example operant box arrangement inside a light-attenuated chamber. A house light is located inside the upper left side of the chamber, out of view. B Closer view of a swivel and counterbalanced arm. C Closer view of manipulanda nose poke ports , active and inactive, positioned at the left and right sides of one chamber wall, with visual discriminative cues cue lights arranged above each port. D Closer view of tether with connector matching the catheter guide cannula threading. Tether length may need to be shortened and weight position changed on the counterbalanced arm so that the tether has an ideal amount of slack; it should not pull down the catheter or impede the mouse's ability to move or access all areas of the chamber floor. Infusion time calculation. Example formula for infusion time, based on mouse body weight, and desired infusion volume, which are used to calculate the desired flow rate for the set-up of syringe pumps. Discriminative cues lights, tones associated with the active port or lever should be activated with drug delivery. The inactive cue is not used. However, timeouts restrict the number of responses that can be made in a session, and if severe, may limit the ability to observe a normal range of drug-taking behavior. Timeouts may be particularly important when switching from a long period of low-moderate dose availability to a much higher dose 7. Alternatively, it may be more specifically used for the signal availability of the drug, remaining on outside of drug delivery and timeout periods. Label each syringe and place them near the surgery area with tips upright to keep sterile. Repeat anesthesia and shaving, if necessary. Monitor the mouse throughout the surgery. Clean incision sites with an alcohol pad or betadine and allow to air dry. Using an insulin syringe, administer ketoprofen 0. Pull up the skin on the back of the neck using forceps, and at the midline, just behind the base of the ears, create a tiny hole with small surgical scissors. Then, using the same motions with the hemostats, tunnel under the skin over the mouse's right shoulder toward the front of the chest. To locate the vein, look closely for jugular vein movement. Using forceps and small scissors, make a small incision from the sternum toward the ear over the vein Leaving them connected, place the syringe and catheter on the platform near the mouse's right shoulder. Very gently pinch the beveled end of the flushed catheter with the hemostats and pull it through the tunnel to the front of the animal Figure 4B. Turn the hemostat handle away from you and set it down behind the right shoulder of the mouse. Jugular vein catheter implantation surgery. A Placement of initial dorsal side incision. B Catheter placement after successful ventral to dorsal tunnel. C Isolated jugular vein. D Jugular vein with suture loops in place, ready for veinous incision. E Successful catheter placement in the jugular vein. F Positioning of catheter base on the dorsal side. With the first pair of forceps left in place, put the other set of forceps, closed, into the same opening and perform the same motion from the opposite direction. Repeat this back-and-forth, with the left and right forceps, several times on one side of the vein. Then repeat on the other side, until the vein is isolated and can be lifted. Be careful not to snag or grab the thin, clear wall of the vein, as this can tear easily, and cause bleeding. Keep the vein hydrated with saline from the rinsing syringe throughout this process. Using forceps, place them under the vein, one at the lower end below the bar and one at the upper above the bar. Turn each thread into a very loose knot around the vein by crossing the ends and pushing one end through the hole twice. Only pull the ends of each knot slightly, so they make relatively wide but secure loops around the vein Figure 4D. Using forceps, gently insert it under the top suture loop, running in the direction of the tail, before re-securing it with the hemostats. Be careful not to tear the vein. Open the artery scissors and orient them downward, placing the ball-tip snug to the underside of the vein and on the same side as the heart. Then close the scissors, which will result in a small cut on the top side of the vein. Use forceps or a needle holder to insert the end of the catheter into the vein incision 13 , pointing toward the heart. A successful vein entry will usually give no resistance Rest the sled on the stack of gauze and use it to guide the catheter into the vein If successful, slowly push the blood back into the animal, using care not to introduce much saline. Relieve the upward pressure on the metal bar, lowering it to rest on the mouse, and tighten the lower and the upper sutures to stop the bleeding Then, using forceps, pinch the skin around the jugular incision, avoiding the catheter. Jiggle the incision up and down gently to settle in the catheter. Suture together the incision, using a modified simple interrupted stitch. Using the long end of the suture, loosely wrap the end of the hemostats twice, then use them to grab the short end of the suture and pull both ends until just tightened at the skin. Check for blood flow with the syringe, as before, to be sure that the tubing position does not impede it. Apply triple antibiotic cream to both incisions. Once the mice are awake and mobile, they can be moved to a clean housing cage with food and water. Clean and re-sterilize the surgical area, then move tools to the platform to dry. Cefazolin should be mixed fresh daily see section 2. Apply triple antibiotic cream to incisions once daily for at least 3 days, and consider at least one saline injection, as described below, for hydration. The mice that are fully recovered from anesthesia and recovering as expected from surgery i. During self-administration, catheters should be flushed both before and after each session. On days with flushing and no testing, once daily is sufficient. Loss of righting reflex should be observed within 3 s of infusion, and mice not meeting this criterion should be removed from the study or may undergo surgery for new catheter placement. Mice suspected of dehydration should be given 0. Triple antibiotic cream should be applied to incisions that are red or oozing. For all these conditions, the facility vet should be consulted as needed. Sessions should be conducted at the same time over days. When deciding which phases to include in each experimental design, limitations in the duration of catheter patency should be a consideration. In this section, we provide general procedural guidelines for running operant sessions. Details and considerations for common specific phases of testing, along with example statistical analysis and results, are included in the Anticipated Results section. Mice may perform operant conditioning better with less stringent i. Reacquisition as described in the Anticipated Results section should be demonstrated before returning the mouse to the unfinished testing phase where they were left off. Experimental timelines for IVSA studies typically include multiple phases of testing, as mentioned above, which may vary based on the objectives of the experiment. For example, a reinstatement phase will provide insight into drug-seeking behaviors after withdrawal De Vries et al. One possible timeline, including acquisition, extinction, dose-response, and increasing cost schedule phases, is provided as an example in Figure 5. In the following sections, we provide anticipated results reprinted with permission from Huebschman et al. Sessions were terminated early if the reinforcer limit was reached 1. Behavioral timeline for example data. Timeline for the example IVSA experiment, including acquisition, extinction, dose-response, and increased cost schedule phases of testing. Modified from Huebschman et al. Acquisition sessions began with the illumination of the active port cue light and a single 1. This dose and schedule were selected to induce stable drug-taking in a large percentage of animals. To limit the contributions of overtraining and variation in total drug intake during this phase, each mouse remained in acquisition only until a predefined set of learning criteria was reached and thus individual animals differed in the number of sessions during this phase. B Mice made significantly more nose-pokes at the active port than the inactive port during the last acquisition session. C The rate at which animals met the criteria and progressed to the next phase of testing. Of the 18 mice included in the acquisition phase, 12 met acquisition criteria within 15 sessions and progressed to later phases of testing. The rate at which these mice met the criteria is shown in Figure 6C. If mice struggle to acquire the task, and it does not interfere with the experimental objectives, interventions as in section 3. In the study shown here, overnight sessions were given to animals that had not yet met the criteria after 5 and 10 days. Extinction sessions were conducted in the same manner as acquisition, except that active nose-pokes had no consequence. Of the 12 mice that underwent extinction, 2 failed to meet the phase criteria within 15 sessions and were excluded from further analysis. The rate at which the animals met the extinction criteria is shown in Figure 7C. B Mice made significantly fewer nose-pokes on the last day of extinction compared to the first. In this study, active-port nose-pokes during extinction had no consequence. Depending on the objectives of the study, extinction sessions may be run such that active nose-pokes either do or do not trigger cue-light illumination. The former helps ensure that, during drug delivery phases, animals respond to the active port for the reinforcer itself and not to the associated cues. On the other hand, the latter is necessary to examine cue-induced reinstatement after the extinction phase, as it is the reintroduction of the cue-light that drives the reinstatement of drug-seeking behavior. Ideally, in either case, saline replaces drug infusion during extinction, a practice that also maintains any sound cues associated with pump operation. However, if necessary, infusions can be omitted. In the example provided here, we used a cutoff of 15 sessions, and mice that took longer were omitted from the additional testing phases. All mice met these criteria within 1—2 sessions. For dose-response testing, a single concentration 0, 0. Doses were presented in sequential order, with the starting dose counterbalanced across groups using a Latin square design. Post-hoc Bonferroni pairwise comparison indicates that responses on the active port for the 0. Dose-response and increasing cost schedule. A Average nose-pokes active port, large boxes; inactive port, and small boxes , excluding those made during time-out periods, during dose-response testing. Following dose-response testing, mice returned to the acquisition dose 1. Inverted U-shaped dose-response curves, as seen here, are typical of drug self-administration tasks, as rodents adjust their responses to maintain targeted minimum satiety Tsibulsky and Norman, Dose-response testing can provide valuable information, as left or right horizontal shifts in the dose-response curve indicate changes in sensitivity or tolerance Schenk and Partridge, On the other hand, vertical shifts have been associated with altered hedonic set points after prior drug experience or escalating drug intake Ahmed and Koob, Upward shifts are also observed in animals that display stronger responses on increasing cost schedules of reinforcement Piazza et al. Intravenous self-administration using operant conditioning techniques has proven a preferred way of studying substance abuse due to its relative resemblance to drug-taking behavior observed in humans. However, amongst rodent animal models, the rat has predominantly been used for IVSA, likely due to technical challenges associated with implanting and maintaining catheters in mice. However, as several labs have demonstrated, the IVSA can be performed in a mouse model and could be more accessible to labs requiring the method. In the Methods section, we describe our optimized approach for conducting cocaine IVSA in mice, including design details for a custom, back-mount catheter that can be prepared in-house, as well as detailed technical surgical advice tailored to jugular vein catheter implantation in this species. We show that these methods result in successful cocaine self-administration using data from the wildtype mouse group in our recently published manuscript, which can be consulted for additional details Huebschman et al. In the study, we moved mice through the acquisition and extinction phases of cocaine IVSA based on when they met the criteria set for learning, then we performed tests for dose-response and increased cost. Reflecting this level of intake, mice remaining in sessions six through 10 show, on average, from 20 to greater than 25 active port responses per 3-h session. We note that mice shown in this study were required to meet slightly more stringent acquisition criteria within seven days, while mice shown in our study took up to 10 days. As such, response levels are difficult to directly compare, but as expected, our mice average lower responses on the last day of extinction. The authors report around 16—18 sessions on average to extinction criteria for their control groups, while our mice met extinction criteria, on average, in 8 days. The importance of the extinction phase with cues cannot be overemphasized, as it tests whether self-administration can be supported by exposure to a cue that was previously associated with drug delivery. Drug reinstatement studies, which necessitate extinction without the discriminative cue, should therefore include the criterion of increased responding when cocaine is again made available after extinction Thomsen and Caine, Interestingly, many studies have also capitalized on this interesting observation of operant sensation seeking in mice, using it as an additional tool to understand behavioral addictions more broadly Olsen and Winder, ; Dickson and Mittleman, Returning to our representative results, our dose-response findings indicate a peak in wildtype mouse responses at the 0. This finding resembles wildtype mice published previously. These comparisons highlight that our results are within the normal limits found in the literature—or deviate in an expected way, such as due to more stringent extinction criteria—but we stress that a range of results can be observed across the literature for all self-administration phases or tests and may relate to differences in the mouse strain, timeout periods and maximum allowed responses, position and salience of discriminative cues, presence of unintended cues such as syringe pump sounds , differences in chamber cleaning practices, etc. The drug self-administration assay has high face validity and is a valuable tool for studying the reinforcing effects of drugs of abuse and substance use disorders. It can be adapted to investigate a wide range of drugs and drug-related behaviors, including initial drug-taking, changes in sensitivity or tolerance, and persistence or reinstatement of drug-seeking. While drug IVSA in mice presents challenges, there are tremendous advantages to being able to use the mouse research model in this assay, including the availability of genetic options, reduced need for the colony and behavioral testing space, lower cage and supply costs, and the ability to maintain continuity with prior or associated studies performed in mice. Our hope is that this detailed protocol for catheter construction, surgery, equipment setup, and basic experimental planning may allow for more researchers to bring the advantages of the mouse model to bear on the problem of understanding the substance abuse. Surgery photographs were taken by GV. Behavioral data were collected by JH. All authors contributed to the article and approved the submitted version. The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest. All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher. We thank Dr. Morgane Thomsen for her generosity with her time and knowledge, which provided the foundation for our methodology. We thank Sadaf Ali for critical assistance with catheter development and Dorothy Foster for help with behavioral testing. If the step for pulling back blood with the syringe is consistently improved by less than maximum catheter insertion, cutting catheters to 1. Measure across just the open space of the inner syringe at the widest point to find the diameter. To check yourself, see your pump manual's Appendix and compare to similar syringes. Look closely for the exit point of blood from the vein when the cut is made will help. If the first few attempts are not successful, targeting the catheter to enter the vein slightly higher than the perceived cut can sometimes increase success. If the catheter cannot be pushed in fully and produces no blood at pullback, this sheath is often the cause. Double check that the cut made to the vein by the artery scissors resulted in bleeding. If blood flow is diminished, minimally loosen the ties one at a time and recheck the flow. Additionally, drug-taking behavior under food-restriction may have limited translational relevance to human drug use. This section collects any data citations, data availability statements, or supplementary materials included in this article. As a library, NLM provides access to scientific literature. Front Behav Neurosci. Find articles by Gia Valles. Find articles by Jessica L Huebschman. Find articles by Elsbeth Chow. Find articles by Corinne Kelly. Find articles by Yuhong Guo. Find articles by Laura N Smith. Smith laura. Received Feb 21; Accepted Apr 21; Collection date Considerations for choosing intravenous vs. Advantages and disadvantages of intravenous vs. Open in a new tab. Considerations for mouse intravenous self-administration compared to other methods. Special attention to humidity and hydration also improves outcomes in mice Thomsen and Caine, Being natural prey animals, fear management e. Magnetic twist ties can be used to elevate and help secure the anesthesia nose cone. The mouse jugular vein cannot be located. The pectoral muscle lays like a pink blanket tucking in the jugular vein, which peeks out from the top. Once the skin incision is made midway between sternum and the ear, use forceps to gently slide top layers of tissue toward the animal's left side, which often reveals the dark red vein. If not successful, use forceps to break the overlying clear fascia, then repeat above. If not successful, puncture the visible layer of tissue discretely and continue to pull aside top layers as you look for the vein. While clearing tissue from the jugular vein, the vein tears. Immediately staunch the bleeding with firm but gentle pressure to the vein or by lifting the surgical bar. If the bar is not in place, try to make a small space to insert it while continuing to control any bleeding. It is possible to place a catheter in a torn vein if it is not severed. To avoid tearing the vein, remove fatty tissue by gripping it away from the vein and slowly pulling in a motion parallel to the vein not perpendicular. Do not grip the vein directly or pull the clear surface of the vein. Opaque tissue fibers that remain close to the vein can be separated and cut against the surgical bar. Death occurs when using catheter sled. Vulnerability to this issue differs by mouse size and background strain. The sled should be inserted minimally and not more than 5 mm. A stack of gauze near the mouse's head should be used to keep the winged end of the sled propped at the level of the vein or slightly above. The catheter cannot be fully pushed in or pushes back out. The catheter is inserted between the vein wall and a sheathe that surrounds the vein. Double check that the vein cut resulted in bleeding. If not, or if there is a tiny amount of blood, make a slightly larger cut at the same position. If bleeding indicates the cut is already adequate, shifting the catheter or sled introduction point up or down the vein length from the perceived cut can help. Attempts at vein entry are not successful. Minimize manipulation of the vein at every opportunity, as it tends to shrink the vein. Target the catheter or sled to enter the vein slightly higher or lower than the perceived cut. Check whether bleeding indicates a sufficient cut and consider improving the cut. If the sled has not been used, try it. If too much time has passed, blood may have clotted. Use rinsing syringe of saline to clear and moisten vein. If saline does not help, a new cut may be required. Surgery time and mortality need to be reduced. Improve technique by inserting the catheter without a sled. While cutting the vein, look closely for the exit point of blood, then move swiftly to introduce the catheter tip firmly at that site, using a lateral and downward motion toward the mouse's vein and feet. If not successful, a sled can still be used. Blood was flowing with syringe pullback, but now there is no blood flow with pullback. Vein suture ties may be too tight. Check blood flow just after securing the ties. If flow is diminished, minimally loosen the ties one-at-a-time and recheck, repeating until flow is restored. If blood flow was confirmed after vein sutures were tied but stops after the mouse is supine, the tubing may be constrained. From the back incision, seat the catheter base and slightly rotate it left or right to adjust the direction of the tubing as it exits the base, while checking blood flow. If blood remained visible in the tubing after a blood flow check, it may have coagulated. Ideally, pushing a tiny volume of saline into the line should move blood easily. If not, gently alternating between pushing and pulling the syringe plunger may loosen the clotting. Avoid pushing much saline into the animal as it can result in death. Mice die or become ill during recovery. Give post-surgery and recovery administrations of USP-grade saline 0. Separate mice recovering from anesthesia from cage mates not anesthetized for at least 24 h. If mice become ill despite antibiotic use, ask your facility vet to test for presence of harmful, systemic bacteria and advice on antibiotic choice. During operant conditioning Mice are slow to acquire task. Ensure mice are not sitting in chambers for long after sessions have ended. Check catheter is patent. If using levers, consider changing to nose poke ports. Should timeout periods or response maximums be used? Timeout periods following drug delivery and limitations on the maximum number of responses allowed in a session may restrict the normal range of drug-taking behavior. Mice are slow to extinguish. Sessions may be lengthened to 5—6 h temporarily. If extinction sessions include the discriminative cue reinstatement not planned , then response requirements can be increased temporarily i. Catheter malfunction or loss of catheter patency during experiment. Remove the catheter, implant a new one into the jugular vein on the mouse's left side, and allow recovery. If the study uses criteria to determine completion of each phase, run each phase as before, repeating any that were already run. Note: this approach may not be appropriate for all study designs. Cocaine concentrations used in the current study to achieve the per infusion doses listed. Similar articles. Add to Collections. Create a new collection. Add to an existing collection. Choose a collection Unable to load your collection due to an error Please try again. Add Cancel. Sterile Blunt Needles, 30 Gauge, 0. Tubing for syringes, drug delivery lines, catheter caps. See Figure 1C. Juglar vein catheter implantation surgery. Hardened Fine Scissors 24 mm cutting edge; length 9 cm. See Figures 4D,E. Metal dust caps thread must match guide cannula on catheter. Habitest Modular Test System—Mouse, including test cage, wall panels, house and cue lights, nose poke ports, flooring, power base and control board, counter-balance arm. Anesthesia cone will not stay properly positioned.

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