JoVE Biology

Survivable Stereotaxic Surgery in Rodents

1, 1, 1, 1

1Department of Pharmacology and Experimental Therapeutics, Tufts University

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    The monitoring of extracellular neurotransmitter levels in distinct brain regions of freely moving animals offers insights on the link between neurotransmitter release and behavior. In vivo microdialysis coupled with electrochemical detection provides excellent anatomical and chemical resolution; and information on how basal neurotransmission is altered by pharmacological or physiological manipulations.

    Date Published: 10/06/2008, Issue 20; doi: 10.3791/880

    Cite this Article

    Geiger, B. M., Frank, L. E., Caldera-Siu, A. D., Pothos, E. N. Survivable Stereotaxic Surgery in Rodents . J. Vis. Exp. (20), e880, doi:10.3791/880 (2008).


    The ability to measure extracellular basal levels of neurotransmitters in the brain of awake animals allows for the determination of effects of different systemic challenges (pharmacological or physiological) to the CNS. For example, one can directly measure how the animal's midbrain dopamine projections respond to dopamine-releasing drugs like d-amphetamine or natural stimuli like food. In this video, we show you how to implant guide cannulas targeting specific sites in the rat brain, how to insert and implant a microdialysis probe and how to use high performance liquid chromatography coupled with electrochemical detection (HPLC-EC) to measure extracellular levels of oxidizable neurotransmitters and metabolites. Local precise introduction of drugs through the microdialysis probe allows for refined work on site specificity in a compound s mechanism of action. This technique has excellent anatomical and chemical resolution but only modest time resolution as microdialysis samples are usually processed every 20-30 minutes to ensure detectable neurotransmitter levels. Complementary ex vivo tools (i.e., slice and cell culture electrophysiology) can assist with monitoring real-time neurotransmission.



    Two-month old average age C57BL/6J mice or equivalent or three-month old average age Sprague Dawley rats or equivalent are anesthetized with ketamine (60 mg/kg i.p. for rats; 100 mg/kg i.p for mice) and xylazine (10 mg/kg, i.p. for either species). Sedation is monitored using a gentle toe pinch withdraw reflex demonstrated in Walantus et al.(JoVE, 6, 2007) and Szot et al.(JoVE, 9, 2007).  Thermoregulation can be provided through a thermostatregulated heating pad (ALA Instruments Inc.) and monitored through a rectal thermometer Head is shaved of fur and cleaned with iodine before incision. After skin incision (2 cm long for rats, 1 cm long for mice) and removal of all soft tissue from the surface of the skull, placement of the guide cannula is determined in relation to bregma. A 6 mm hole is drilled through the skull with a battery-operated driller designed for rodent surgery (Fine Science Tools, Inc.). Care is taken so that the drill bit does not penetrate through meningeal membranes or blood vessels. Skull is implanted with bilateral 5 mm 21 gauge stainless steel guide shafts leading to the posterior nucleus accumbens, dorsal striatum or prefrontal cortex. The stereotaxic coordinates are established as per Franklin and Paxinos, 1997 (The Mouse Brain in Stereotaxic Coordinates, Academic Press) or Paxinos and Watson, 2006 (The Rat Brain in Stereotaxic Coordinates, Academic Press). Implants are secured by dental cement. A bolus of Lactated Ringers of the 0.9% saline is given at the end of surgery (5mls SC in rats and 1 ml SC in mice after fluids are warmed to normal body temperature) to prevent dehydration. Buprenorphine (0.1-0.5mg/kg SC) is administered twice daily and, then, on an as-needed basis, if animal appears to be in pain. Local antibiotic treatment (bacitracin ointment) and systemic antibiotic treatment (penicillin 100,000 IU/kg IM every 12 hours for the first 48 hours post-op) are administered if post-operative infections occur.

    Following surgery, animals are individually housed with food and water available ad libitum. At least one week is allowed for recovery before microdialysis and euthanasia. Following recovery from surgery, the animals are transferred to a microdialysis cage and microdialysis probes are inserted and cemented in the guide shafts that have been installed during surgery. Probe insertion does not cause pain or discomfort because the probe is bypassing skin, muscle and meningeal tissue through the guide shaft. Therefore, probe insertion is done without anesthesia and any anesthesia-induced effects on neurochemistry or behavior are avoided. We let the probes stabilize for 12 hours and then we start sampling every 30 minutes for another 8-12 hours depending on the experiment. We monitor locomotor behavior of the animal through photocells or manual recording of movement by the experimenter. Microdialysate samples are injected into a High Performance Liquid Chromatography with Electrochemical Detection (HPLC-EC) instrument for neurochemical detection and analysis. We look for effects on basal neurochemistry and locomotor behavior. At the end of the experiment the animal is euthanized by an overdose of systemic ketamine (200 mg/kg i.p.) and xylazine (20 mg/kg, i.p.). Then the heart is perfused with 0.9% saline followed by 4% paraformaldehyde. The brains are removed, frozen and cut along the microdialysis probe tract to verify accurate probe placement.


    1. Set up the stereotaxic instrument and all the materials needed. Make sure the area and instruments are cleaned and sterilized.

    2. Shave off fur with electric razor. Go from the ears to just in-between the eyes, move razor in different directions to effectively clean area of fur. Apply povidine/iodine to shaved area but protect the eyes from it.

    3. Mount the animal onto the stereotaxic apparatus by placing the ear bars into the ear canal and tightening into place. First mount one ear bar in the ear canal, and then hold it in place and slide in the other ear bar. You know you are in the correct location when the head can no longer be moved side to side. Secure the mouth with the anterior mount of the stereotaxic and make sure that the head is level with a ruler. Put the ruler in a vertical position with respect to the stereotaxic instrument platform and check for a 90° angle between the ruler and the middle of the animal’s scalp). Confirm this through the stereotaxic instrument if it offers such capability.

    4. Make an anterior/posterior incision on the scalp with a sterile scalpel extending from the lambda to just in-between the eyes of the animal. Use sterilized hemostats to pinch off the skin and keep the incision open. Using several sterile cotton swabs, dry off the exposed skull surface.

    5. Put the guide cannula onto its mount, find bregma on the skull, and position the guide cannula right over this location. Write down the anterior/posterior and lateral coordinates. From bregma, find the correct coordinates needed for the placement of your probe with the aid of the stereotaxic atlas. Position the guide cannula to the correct coordinates by adding or subtracting from bregma. Bring your guide cannula down until it is touching the skull, and then record this ventral coordinate. Make a pencil mark with a sterile pencil at this location on the skull; this is where you will be drilling.

    6. Remove the guide cannula and sterilize your drill bit. Carefully drill a hole at the pencil mark until you get through the width of the skull. Check with the guide cannula to see if it would clear the hole without touching the sides. Keep drilling and checking until the cannula can clear in a straight path. Once the hole is made, use a sterile needle to gently punch the meninges in order to allow unobstructed insertion of the cannula.

    7. Next, using a hand drill, make six holes for skull screws: two anterior to the cannula hole, two lateral to the cannula hole, and two posterior to the sides. Sterilize six screws and place them onto the skull until they are tightly anchored on.

    8. Clean the guide cannula with ethanol and saline, mount, and lower it slowly to the proper ventral coordinate. Make sure that the sides are not touching and that it is going in perfectly vertical.

    9. Place the anchor screw medially and behind the posterior skull screws and hold it in place with tweezers. Mix a thin batch of liquid dental cement and cover the guide cannula, screws, and the rest of the skull with a sterile spatula. Make another batch, thicker this time, and completely cover the area and the cannula and anchor screw enough to secure it.

    10. As the cement becomes thicker and before it solidifies, separate the skin from the cement cup and mold the cement cup with the spatula to make sure the cement cap is smooth all around and does not irritate the skin later.

    11. Allow the dental cement to completely dry before removing the animal from the apparatus. Remove the hemostats. Apply bacitracin all the way around the cement cap.

    12. Once the animal is off the stereotaxic instrument, inject it with 0.25 ml of penicillin IM (intra-muscular) followed by 1 ml of saline SC (subcutaneously).

    13. Place the animal in its own cage and monitor it until it becomes conscious before returning it to its room to recover.

    14. Monitor animals until they recover from anesthesia on the day of surgery and daily post-op, until the end of the experiment, for signs of infection and evaluation of pain/distress. This includes weekends and holidays. Low spontaneous movement, distress vocalization upon handling, hunched posture, diarrhea, swelling and discharge in the area of the headmount, and lack of feeding/drinking are all signs of pain and distress. Buprenorphine (0.1-0.5mg/kg SC) is administered twice daily, and then, on an as-needed basis, if animal appears to be in pain. Local antibiotic treatment (bacitracin ointment) and systemic antibiotic treatment (penicillin 100,000 IU/kg IM every 12 hours for the first 48 hours post-op) are administered if post-operative infections occur. If any of these symptoms persist following administration of buprenorphine, supplemental fluid, and antibiotic treatment within 12 hours of surgery, the animal is euthanized.


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    In vivo microdialysis is the tool of choice for measuring multiple neurotransmitters and metabolites in distinct brain sites of a living animal. However, it only monitors extracellular levels of neurochemicals and it does not offer the time resolution to monitor neurotransmitter exocytosis in real time. Through a version of the technique called "net-flux", the actual neurotransmitter concentration at a given site can be calculated, which in turn can give accurate measurements of neurotransmitter rate of reuptake through plasma membrane transporters.

    Microdialysis is ideal in illustrating differences in basal extracellular neurotransmitter levels between different groups of animals (i.e. different genotypes) and in deciphering the effects of drugs or other manipulations on neurotransmitter release.

    The introduction of assays alternative to HPLC-EC like capillary zone electrophoresis (CZE) coupled with fluorescent detection has increased the time resolution of in vivo microdialysis within a few minutes per sample.

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    The authors have nothing to disclose.


    Supported by DK065872 (ENP), a Smith Family Foundation Award of Excellence in Biomedical Research (ENP), F31 DA023760.


    Materials are described in the protocol document.


    1. Bungay, P. M., Newton-Vinson, P., Isele, W., Garris, P. A., Justice, J. B. Microdialysis of dopamine interpreted with quantitative model incorporating probe implantation trauma. J. Neurochem. 86, 932-946 (2003).
    2. Chen, K. C. Effects of tissue trauma on the characteristics of microdialysis zero-net-flux method sampling neurotransmitters. Journal of Theor. Biology. 238, 863-881 (2006).
    3. Geiger, B. M., Behr, G. G., Frank, L., Caldera-Siu, A. D., Beinfeld, M. C., Kokkotou, E. G., Pothos, E. N. Evidence for defective mesolimbic dopamine exocytosis in obesity-prone rats. FASEB Journal. 8, 2740-2746 (2008).
    4. Pothos, E. N., Creese, I., Hoebel, B. G. Restricted eating with weight loss selectively decreases extracellular dopamine in the nucleus accumbens and alters dopamine response to amphetamine, morphine and food intake. The Journal of Neuroscience. 15, 6640-6650 (1995).



    If I undertand the study correctly, it would be interesting to report the results of rate of reuptake for neurotransmitter, which is the main purpose of the empriment. 

    Posted by: AnonymousOctober 10, 2008, 6:14 PM

    Calculations of the reuptake rate of neurotransmitter can indeed be accomplished through net-flux microdialysis. However, the primary objective is the measurement of basal extracellular levels of neurotransmitters and their metabolites.

    Posted by: AnonymousOctober 10, 2008, 6:45 PM

    The authors have wonderfully demonstrated how to perform the stereotaxic experiment in rats. However they should have added few more words on how the ear bars should be adjusted so that it shows equal readings on both the sides before the opening of skull. In my experience I have noted that before opening the skull, one should make sure that both the vernier scales of the ear bars show almost correct readings in order to make sure the skull is on the right path, failure of which might lead to the miscalculation of the stereotaxic coordinates. Thanks and Regards, Rajesh S Omtri.

    Posted by: AnonymousOctober 12, 2008, 5:30 PM

    Correct placement of the ear bars is clearly a practice effect. We usually have one of the ear bars tight in position, then insert the tight ear bar in the ipsilateral ear canal, hold it in place and slowly insert the loose ear bar on the contralateral side before tightening it down. It is desirable that the skull is centered in between the ear bars. The skull surface must be always level (parallel to the platform of the stereotaxic instrument and at 90° to the guide of the microdialysis cannula) and skin at the incision surface should be flat and present no humps. These problems occur if the ear bars are inserted incorrectly (not in the ear canal). Correct ear bar placement can be identified by gently trying to move the head of the animal up and down and left to right. Before tightening the incisor bar, up or down movement but not lateral movement should be possible. Correct placement of the ear bars in the ear canal is the most important prerequisite for accurate stereotaxic placement. Emmanuel Pothos 

    Posted by: AnonymousOctober 12, 2008, 6:16 PM

    I am a scientist and I find it very hard to see such a gruesome procedure like this one. There should be a clear label on the content of videos as they can be quite disturbing, and they shouldn't be automatically broadcasted on the main website page. I wonder if the editors of this journal seriouly consider the possibility of risks the authors might face by being attacked by animal activists, and if the Principal Investigators of similar papers are held liable for exposing their students' identity to those groups while making these videos. This message dŒs not intend to diminish the value of the present work, but to bring this serious issue to the attention of the editors and the authors who appear on the video.

    Posted by: AnonymousOctober 20, 2008, 3:55 AM

    All procedures described in this article have been reviewed by the Institutional Animal Care University Committee at Tufts Medical Center and approved as compliant with federal and state standards of animal care. JoVE also conducted a veterinary review of the article before publication; nothing was "automatically broadcasted" as the viewer claims. Animals are anesthetized before any type of brain surgery, carefully monitored for appropriate depth of anesthesia and hydration during the procedure and diligently followed up through postoperative care with analgesic medication and antibiotics until full recovery. Stereotaxic brain surgery is one of the most sophisticated procedures in live and survivable animal surgery and it normally involves minimal pain for the operated animal because of the conditions set in place as described above. Stereotaxic brain electrode placement is a procedure that has been routinely used even for humans (i.e. Parkinson's disease patients) and such operations have been repeatedly broadcasted over the Internet from several hospitals for educational purposes. In some of these cases, the discomfort of the patient is so minimal that general anesthesia is not used and the patient is awake during surgery and able to respond to questions from the surgeons, who use the patient's response to assess the accurate placement of the electrode in the brain. The whole process in animals or humans is elegant, effective and high technology driven, not gruesome and painful. We appreciate the concern of the viewer about safety issues, but scientists have to take responsibility for their own work and it is not appropriate to publish it anonymously, being in this journal or elsewhere. Otherwise, the whole concept of the validity of peer-reviewed research and accountability of authors for their work is negated. There are numerous pieces of published work in different journals, including dissection videos, autopsies of animal tissue, images of animals etc. that can potentially be used by extremists to target the authors. Censoring scientific journals and scientists cannot eliminate this possibility.

    Posted by: AnonymousOctober 20, 2008, 8:30 AM

    Stereotaxic surgery should be performed under aseptic conditions. The surgeon shoud have a cap, mask, and surgery gloves. She should not be touching non sterile items while doing surgery, i.e. pens, cannula etc. Ophthalmic ointment is essential. Hemostats are not good  skin retractors as they damage tissue. There are antibiotics that can be given subcutaneously, which is easier and less painful to give.

    Posted by: AnonymousFebruary 2, 2009, 4:18 PM

    There is not such a thing as sterile stereotaxic surgery in living animals. The mere presence of a living animal on the table with its fur, bodily fluids etc. negates sterile conditions. Doing the procedure under a culture hood with negative air flow is also not advisable as it limits access to the animal from all angles, it makes it more difficult for the animal to maintain appropriate temperature due to the air flow and it contaminates the hood area, which is counterintuitive particularly if the hood is used for cell cultures. The most appropriate actions are to sterilize the components used for the surgery (i.e. cannula and skull screws) prior to use, sterilize all insrtruments before surgery and during surgery as needed and maintain as clean of an environment as possible in the incision area by shaving away the fur and treating with povidine prior to the incision. Gloves should be used, face mask and cap will not hurt but none of the above will ensure sterile conditions. There is a variety of skin retractors available, we have not found that hemostats are worse than others in damaging tissue. Antibiotics given subcutaneoulsy are acceptable, but not as long lasting as those given intramuscularly. In any case, the easiest antibiotic to administer is bacitracin, right around the headcup of the animal. Emmanuel  

    Posted by: AnonymousApril 22, 2009, 5:43 PM

    Suggestion for dental cement:  My lab uses a UV dental acrylic that is much easier to handle.  The acrylic sets when exposed to a UV light in about 10 seconds, and we do not need to use bone screws to secure the cap.  However, I'm guessing that the UV acrylic is more expensive.  Its available from Pentron. Oh, and don't forget eye lube.

    Posted by: AnonymousNovember 1, 2008, 10:56 PM

    We have tried in the past to use dental cements that their manufacturer claims do not require head screws.  We were not convinced. In many cases the cement head cup came off as one piece as we were trying to implant the microdialysis probe. Using sterile head screws is the best way to ensure that the cement cup will be securely attached to the skull. Any other method shaves off about ²0 min from each surgery but it increases the probability that the cup will come off and waste the entire procedure. Suppliers do tend to charge more for cements that supposedly work without head screws so in the long run this is not cost effective.  Eye lube as an eye protectant is indeed a very good precaution for this procedure. 

    Posted by: AnonymousNovember 1, 2008, 11:12 PM

    Nice demonstration of stereotaxic surgery in rats. I think that the best way to control that the skull is perfectly flat (parallel to the platform) would to check the height coordinates at the bregma and at the lambda using the canula as recommended in the stereotaxic atlas. That might not be a problem for ICV canulation since the ventricle are quite big but for canulation in a specific structure or nucleus it is critical.  I usually use only 4 screws but I guess 6 are necessary for a microdialysis probe. Also, do you calculate the coordinates from the surface of the skull or from the dura ?

    Posted by: AnonymousDecember 2, 2008, 11:30 PM

    We calculate steotactic coordinates from the skull surface.

    Posted by: AnonymousDecember 3, 2008, 12:11 AM

    how can i download (Survivable Stereotaxic Surgery in Rodents) thanks

    Posted by: AnonymousDecember 18, 2008, 5:18 AM

    Hi.  Please contact us at

    Posted by: AnonymousApril 17, 2009, 11:09 AM

    ı can not understand that why u r doing such as these trials for understanding brain mechanism, cuz I believe that if somethings can not explained naturally, we also can not understand exactly

    Posted by: AnonymousJanuary 7, 2009, 7:54 AM

    Hey, it is obvious that you are not Dr. Ayla Arslan, then who are you? it seems like you are one of her students using her name as a nick as she always recommend JoVE during her Biopsychology lectures. :)))))))

    Posted by: AnonymousOctober 31, 2009, 6:14 PM

    Hi                 I am khalid a Ph.D scholar in deptt of pharmacy,university of Peshawar Pakistan.Its really great contribution to science and i eally enjoyed and learnt alot from the movie of Survivable Stereotaxic Surgery in Rodents thanks indeed and keep up this great work. khalid rauf

    Posted by: AnonymousFebruary 26, 2009, 10:56 AM

    Wow, you guys really do not knwo what you are doing. Why would you use the archaic acrylic dental cement when you could use Glass Ionomer Luting Cement? Why didnt you anesthiatize with O² delivered isofluorine? Why did you not sue a stereotaxic drill? Why was the cement applied so sloppy? Why do you not use a digittal display for the coordinates, it ensures much more precise surgeries.

    Posted by: AnonymousMarch 12, 2009, 9:38 PM

    Hi Dave,

    How dŒs the Glass Ionomer Luting Cement compare with the Light-cured Dental Adhesive Resin listed in this journal by Okamura lab?

    Look forward to hearing from  you.



    Posted by: AnonymousMarch 15, 2009, 12:35 PM

    In our hands, dental acrylic is the only cement that ensured headcups stayed on for several weeks when used in combination with 6 skull screws. Emmanuel

    Posted by: AnonymousApril 22, 2009, 4:04 PM

    Do you use any preanesthetic medications?

    Posted by: AnonymousSeptember 19, 2009, 12:51 PM

    Usually not, if the animal suffers from CRD (chronic respiratory disease) you can pretreat with atropine to facilitate breathing. However, CRD is an indication of substandard conditions in the animal colony (infrequent change of bedding, poor air flow etc.). If you have animals with CRD, consult with your veterinarian to improve your facility and check on your source for laboratory animals, whether commercial or another lab, for facility conditions as well.

    Posted by: AnonymousOctober 18, 2012, 6:29 PM

    I am an undergraduate at the University of California, Santa Barbara doing an Honor's thesis project on the rat dorsal Raphe nucleus. In my project, I need to implant a cannula into the dRN, but am concerned about profuse sagittal sinus bleeding if I go through the midline. I noticed in other papers they often go into the DRN at about a 30 degree angle, in order to avoid this issue and also to avoid the cerebral aqueduct. As the angled cannula is a more complicated procedure, for me it would be easiest to place the cannula at the midline, and I'm wondering what's the best way to deal with these issues, such as how bleeding is stopped or slowed down, how it can be avoided, how many animals I can expect to lose, etc. Any advice would be much appreciated!

    Posted by: AnonymousOctober 27, 2009, 3:54 PM

    The angled approach is the best, but if you encounter sagittal sinus bleeding make sure to put in place large cotton tips from a sterile bag, press gently for a few minutes to slow down blood flow and leave on until blood has clotted. Then very carefully remove cotton tip to avoid breaking the blood clot. Although this bleeding would be fatal in humans, it usually is not fatal in rats. Emmanuel

    Posted by: AnonymousOctober 18, 2012, 6:34 PM

    nice job.
    just some comments:
    the membrane (after cuting the skin to expose the skull) should be carefully and totally removed - this decrease the chances of the acrylic fall off.
    if you do a small cut, 1 or ² screw would be enough.
    another important thing, regarding guide cannula is that it should be obstruct after surgery so that no reflux happens and nothing enters for this hole - if this happens you can loose all you surgery. If, when you try to put a needle inside for drug injection (p.ex) and it dŒsnt enters, you can use a some H²O² to open it (in case of blood coaguation)


    Posted by: AnonymousDecember 11, 2009, 4:52 PM

    We use obdurators to seal off guide cannulas post-op. We avoid using only 1-² screws no matter what the size of the incision, this is clearly inadequate anchoring for the headcup and it will come off in a matter of a few days at best. It really pays off to anchor the headcup with as many screws as you can. Emmanuel

    Posted by: AnonymousOctober 18, 2012, 6:46 PM

    Oh! another thing...
    would be really good to use local anesthetic with vasoconstrictor before cutting the scalpe.
    this will minimize animal nociception and will avoid excessive bleeding.
    but not toooooo much, other wise wont be god to animals, and we also see some increase of infeccion

    Posted by: AnonymousDecember 11, 2009, 4:57 PM

    The authors and our attending veterinarian would like to add the following information to the article, which was not otherwise clearly stated or shown and may be of help to readers and viewers:
    1) After anesthesia and prior to surgery, eye lubricant was applied to protect the corneas of the animals.
    ²) Prior to inserting the ear bars into the rats' ears, lidocaine gel was applied to provide analgesia.
    3) All rats did receive an initial dose of buprenorphine following the surgeries and then were given subsequent doses on an as needed basis. This was not clear in the video or text.
    4) The dose of penicillin given was 100,000 IU/kg.
    All of the above measures were approved in our IACUC protocol for this procedure and our attending veterinarian has reviewed and verified these additional comments.

    Posted by: AnonymousApril 28, 2010, 3:07 PM

    is it better that ketamine is better than halothan as a anesthetic agent

    Posted by: Ravi S.May 14, 2010, 2:16 AM

    my just quest is how the hole in the skull is closed/filled? Did you leave it for natural tissue growth or use gelatin or something else.thanks

    Posted by: AnonymousMay 14, 2010, 9:48 AM

    In our case the guide cannula leaves very little space to add anything else. Some of my colleagues are using bone wax or gelatin for larger openings. Emmanuel

    Posted by: AnonymousOctober 18, 2012, 6:41 PM

    No flash please

    Posted by: AnonymousAugust 8, 2010, 1:31 PM

    Overall a nice video, but there are a few things that should be done to improve aseptic technique. The eyes need to be lubricated prior to shaving to protect them from the hair and from dessication. To reduce infections a surgical drape should be used, along with a surgical mask. Lastly, pointing with a sterile instrument would have been better.

    Posted by: AnonymousNovember 5, 2010, 6:34 PM

    It is a very nice presentation, I would like to add a little in it . When the animal id fixed with ear bars and the scale on ear bar and the scale of sterotaxic base should be equidistant
    See in video your demo point 0²:51.


    Posted by: AnonymousMay 24, 2011, 3:35 AM

    Im a 4th year Psych Honours student doing a project that needs me to implant canulas into the infralimbic. I just did my second practice surgery today, and it was terrible. The cannulas were mislocated, it took me an hour to put in 4 bone screws, and they went through the skull, and the dental cement ran into its eyes, and Im just glad that rat was put down before it woke up because there is no way it would have survived. I've always been sort of clumsy, and I have to do 35 of these, and half my year is gone and I dont have time to come up with another project.
    So yeah, Im freaking out right now,

    Posted by: AnonymousMay 30, 2011, 8:58 AM

    Hi Nadia,
    It seems you do not have adequate training and supervision to perform this procedure. It is essential that a member of your laboratory team with extensive experience in stereotactic surgery, if not the primary investigator directly and the head veterinarian for your institution's animal facility, should go over things with you multiple times and actively do the procedure with you before any further attempts. You have to ensure the animal's welfare, lack of pain and recovery during and after the procedure if you wish to be anywhere close to acceptable standards. In my opinion, brain stereotactic surgery is an advanced procedure that should be used only with the maximum of caution and the best of training for undergraduate projects. If the animal facility or your faculty supervisor do not have the time or the skills to train you properly, then it would be best to choose something else for your thesis project. Best, Emmanuel

    Posted by: AnonymousMay 30, 2011, 10:46 AM

    Hi everyone,

    Just a quick question, after attaching the ear bar I have realized that it takes me quite a while after making an incision in the skull to expose the bregma and lambda. Any suggestions?

    Posted by: Jin P.November 5, 2012, 6:41 PM

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