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Temporal Strategies in Visual Cortex
"Personnel Working on Study" Jeramy L
Kulesa Phone: 612-625-8847
Thomas J Nelson E-mail
Address: nels5223@umn.edu
Xu Yang: (No info)
Information on Study (Taken from IACUC documents) View the IACUC Animal Usage Form 6 Rhesus monkeys The non-human primates will be killed with a drug overdose of Thiopental (150 mg/kg IV) or Pentobarbitol (120 mg/kg IV).
We will perform chronic neurophysiological recording experiments from trained, behaving macaque monkeys. These experiments involved controlled fluid intake during behavioral training and neurophysilogical recording, as well as recovery surgeries in which head-restraining posts, eye monitoring coils, and recording chambers are implanted. The animals are euthanized at the end of the experiment or sold or made available to other investigators requiring primates. The following chart indicates the experimental sequence for the animal. Magnetic Resonance Imaging of Cerebral Cortex: MRI images afford a high resolution map of the animals’ brains that can be used to guiding the placement of recording chambers and electrodes. These maps reduce the number of recording sessions for a given animal, and in some cases can obviate the need to euthanize the animal for histological verification of recording sites after we have finished our experiments. The procedure itself is noninvasive and can be considered diagnostic. Animals will be administered atropine (0.05 mg/kg), sedated with ketamine (15 mg/kg). A catheter will be inserted in a leg vein, and the animal will be intubated to maintain an airway. The animals will then be anesthetized with propofol (2 mg/kg IV), placed in a primate chair, and secured via a surgical implanted plastic headpost (see below). Anesthesia will be supplemented by continuous IV drip of propofol (0.5 mg/kg/min). Anesthesia level will be verified by continuous monitoring of pulse oxygenation, pulse rate, expired CO2, and respiration rate. The animal is wrapped in a chemical heating pad to maintain body temperature, which is monitored by a rectal thermister. After data acquisition, animals are returned to their cages and monitored until they are sitting upright. Behavioral Training: Our experiments require that the animals direct their attention to specified positions in the visual field while holding their gaze on a fixed location. The duration of the training period depends on the tasks and the characteristics of the animal. It lasts up to 16 months, with the animal trained 5 days a week. Animals are first conditioned to get into and out of a primate chair and maintain a relaxed, normal posture while in the chair. The primate chairs are adjustable along several dimensions to allow comfortable sitting posture. Each animal is completely unrestrained within it standard operant condition techniques with positive reinforcement (delivery of fruit juice or water for correct responses). Depending on the task, a correct response is wither a hang (level release) or eye movement made at an appropriate time. If the animal will use a lever, we normally train it to do the basic task without eye position requirements before implanting a seleral search coil and headpost (below). This typically takes 2-4 months. After the surgery, we wait at least two weeks before we begin to train under head restraint. The head post attaches to a mounting bracket on the primate chair and ensures the head stability that is needed for monitoring eye position and recording from individual neurons. The duration of head restraint is initially brief and is gradually increased during the process of training. Animal adapt readily to head restraint. The head post is coupled directly to the skull and therefore genereates not pressure on the skin. If the animal will make an eye movement for its response, the search coil and headpost are implanted before any task training. Once the animal is conditioned to head fixation and eye position can be monitored, it is trained to perform its task while holding its gaze on a small spot in the center of the video display. This fixation control allows us to know where the visual stimuli fall on the animals’ retinas and thereby create controlled, repeatable stimuli. With tasks involving eye movement responses, the animal to trained to move his eyes from the fixation spot to a specific target on the screen. Once the animal is proficient at the task, we being to collect neurophysiological data. Neurophysiological Recording: When training is completed, animals undergo a surgery in which a recording cylinger is implanted on the skull (details in section R8). When recording from surface regions on the cortex we use microelectrodes which penetrate the dura. For those recordings a small craniotomy (2-4 mm) is made inside the camber under ketamine anesthesia the day before the start of recording. Delaying the craniotomy makes it possible to collect as much data as possible before the dura mater thickens to a point that makes microelectrode penetration impossible, which occurs over a 3-6 weeks. In order to maximize data collection during this period there will typically be six recording sessions per week. When the dura becomes impenetrable, we either remove its overlying granulation tissue (under ketamine anesthesia), make a new craniotomy, or extend the existing craniotomy. For this procedure, which typically take about 15 minutes, the animal is sedated with Ketamine (10 mg/kg IM). We typically make 3-4 craniotomies within each chamber and remove granulation 1 or 2 times from each craniotomy. When it is necessary to record from deep brain structures, a larger craniotomy is made at the time of surgery to allow maximal flexibility in electrode placement. For such recordings, a guide tube is used to bring the electrode to within 10 mm of the target. The guide tube is constructed from stainless steel hypodermic tubing and advanced through a grid within the recording chamber. The portion that punctures the dura mater and enters the brain is 28 gauge. When a guide tube is being used, both the electrode and the guide tube are soaked in disinfectant before insertion. Data are collected using metal microelectrodes that are advanced into the brain using a hydraulic microdrive. The microelectrodes are advanced transdurally in a procedure that does not require full aseptic methods or anesthesia. The brain itself does not have sensory endings, and the microelectrodes are smaller than fine hypodermic needles (~100um O.D.). Electrodes are advanced until the action potentials from single neurons can be well isolated. On each day of recording the animal is put in its chair and brought to the animal prep area in the laboratory, where its head post is secured to a mating bracket. The external surfaces of the recording chamber cap and the surrounding bone cement are cleaned with a disinfecting solution (Novalsan), and the cap is removed by personnel wearing a surgical mask and gloves. Only sterile instruments and solutions enter the chamber. The interior of the chamber is rinsed with sterile saline and disinfectant is applied with sterile swabs. Secretions and debris in the chamber are removed by suction. A microdrive carrying a microelectrode that has soaked in Novalsan for 1 hour is fitted over the chamber, which is then filled with sterile mineral oil and sealed. The animal is placed in the recording room and the microelectrode is slowly advanced through the intact dura mater and into the brain. When the day’s recording is completed, the microelectrode is retracted from the brain and dura mater. In the animal prep area the chamber is drained of oil and the microdrive is removed. The chamber is rinsed with sterile saline and disinfectant and a final sterile saline rinse, and closed with a sterilized cap. When all data have been collected from one recording cylinder, we typically implant another over a different region of cerebral cortex or the other cerebral hemisphere, and this is used for further recordings for about another 3 to 6 month period. Almost all animals received are implanted with 2 cylinders. It is less common for us to implant more than 2 cylinders, because there is generally no suitable scientific target or no room for access. General Surgical Procedures and Post Surgical Treatment: The monkey is injected with atropine (0.05 mg/kg IM) to avoid congestion, followed in 10-15 minutes by a dose of Ketamine (15 mg/kg, IM) An intravenous catheter is placed aseptically in the saphenous vein. The catheter is fixed to the surrounding tissue and managed with a dry dressing. The larynx is sprayed with lidocaine and the monkey is intubated to maintain a clear airway. Hair is removed from the surgical site with electric clippers. Cenftriaxone (roccephin) will be administered (100 mg/kg, IV) while the animal is being stereotaxically secured and prior to any incisions. The animal is placed in a steroeotaxic apparatus on the surgery table to provide coordinates for the placement of skull implants and to assure stability during surgery. Isoflurane anesthesia is started at this time and continued throughout the procedure. Because of health concerns, we use barbiturate anesthesia in place of isoflurane during the recovery surgeries on the animal subjects of pregnant investigators. The level of anesthesia is monitored using pinch reflex, corneal reflex, jaw tone, respiration and the heart rate and rhythm, expired CO2, O2 saturation, and mucous membrane color. Throughout the surgical procedure the animal is administered intravenously 5% dextrose in lactated Ringer’s solution at a rate of about 5-10 ml/kg/hr. Body temperature is kept at 37-38 deg C using a thermostatically controlled heating pad or warm air blanket. The surgical sites are then scrubbed with Betadine. Surgical attire includes a scrub suit, shoe covers, cap, mask, sterile gown and sterile gloves. All instruments, solutions and drapes are sterilized, and sterile procedures are used throughout. At the end of the surgical procedure the animal is extubated. When the monkey regains consciousness it is returned to a recovery cage in a separate room, where it is monitored continuously until it is able to sit up. A heat lamp is provided. Antibiotics and analgesic are given while the animal is still sedated. Sutures and staples are removed 10-14 days after the operation. Collegen-based postoperative skin treatments (Medifil or Collasate) are applied as needed to encourage recovery around the implant margins. Head Post Surgery: An crescent-shaped incision is made in scalp so that a flap of skin can be retracted. The retracted flap is wrapped with saline-soaked gauze. The skull is then scraped clean of periosteum. A plastic (PEEK) or titanium headpost whose base is composed of 5 plates is placed on the skull. Each plate has 2 to 3 screw holes. Holes are drilled and tapped in the skull and screws are inserted through the plates and into the bone. Once the plates are secured, the leads from the scleral search coil are routed subcutaneously from the zygomatic incision and attached to a connector. This connector is then secured to the head post. The skin incisions are then closed using either nondigestible sutures (Ethicon) or stainless steel staples. When all procedures are completed, the animal is removed from the stereotaxic holder and anesthesia is discontinued. Recording Imaging Chamber Implant: The monkey is placed in the stereotaxic apparatus, and a skin flap is made to expose the region at which the stainless steel or titanium chamber will be attached. The fascia are dissected from the muscle sheath, and skin and fascia are retracted and placed under saline soaked gauze. The chamber is then placed the desired coordinates and angle, and the location marked on the skull. One to three stainless steel or titanium orthopedic plates (Synthes, veterinary) with 3 to 6 screw holes are then bent to fit the skull in orientations that leave one end adjacent to the chamber. The last cm of each plate is bent to rise vertically from the skull along the sides of the chamber. Holes are drilled and tapped and screws are inserted through each plate and into the skull. Using the stereotaxic carrier, the chamber is placed in the desired position and the dental cement is applied to bond it to the plates. The chamber is then capped. The fascia and skin are sutured using digestible sutures for the fascia and nondigestible sutures or stainless steel staples for the skin. The monkeys is then removed from the stereotaxic instrument and anesthesia is discontinued.
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