Microsoft word - telemetry unit implantation in rodents2.24.201
Stony Brook University
Institutional Animal Care and Use Committee (IACUC)
TELEMETRY UNIT IMPLANTATION IN RODENTS
1) Anesthesia is required for surgical implantation of the telemetry unit and associated
2) Anesthetic agents – animals will be anesthetized with one of the following agents for
Ketamine (90 mg/kg) and Xylazine (10 mg/kg), IP
Ketamine (75 mg/kg) and Xylazine (10 mg/kg), IP
Isoflurane via vaporizer, delivered to effect.
3) Use of any other anesthetic agents must be identified in the IACUC application.
B. Anesthesia Monitoring
Prior to and during the procedure the following parameters will be monitored at a minimum of 5 minute intervals:
• Respiratory rate • Response to noxious stimulus (ie toe pinch) • Spontaneous movement
C. Anesthesia Recovery Monitoring
1) During recovery from anesthesia, the following clinical parameters must be monitored
at a minimum of 15 minute intervals until the animal is ambulatory:
• Respiratory rate • Movement • Ability to maintain sternal recumbancy
2) To protect the animal from hypothermia they will be placed on a water recirculating
heating blanket or covered well, to conserve body temperature. Animals should never be placed directly on metal surfaces.
3) It is estimated that animals will recover within 30-60 minutes post-operatively.
A. Telemetry Unit Insertion Procedure – Femoral catheter/subcutaneous transmitter
All surgical procedures must follow the Guidelines for Rodent Survival Surgery at Stony
1) Shave and disinfect the ventral abdomen and the inner thigh region on the side of
proposed transmitter placement and drape surgical site. Disposable, adhesive drapes are preferred for rodents.
2) Open the skin over the femoral vessels. 3) Using blunt dissection, form a subcutaneous pocket up towards the area between the
caudal edge of the ribcage and the most cranial extension of the knee's range of motion. Ideally, the subcutaneous pocket should be just large enough for the transmitter body to be inserted into the pocket, with the tissue snug but not taut over the transmitter.
4) Once placed in the pocket, secure the transmitter housing by passing 5-0 sutures
through the tissues surrounding the pocket entrance and drawing together the entrance in a purse-string fashion.
5) Locate and expose femoral vessels. They are bundled together with the saphenous
nerve and can be found between the abdominal wall and the branching point of the caudal epigastric artery and vein.
6) Carefully isolate the femoral artery from the femoral vein and saphenous nerve (~10
mm if possible). NOTE: Care must be taken not to damage the nerve during isolation of the artery to prevent hind limb paresis or paralysis.
7) Pass three lengths of 5-0 non-absorbable suture underneath the isolated artery section
(proximal occlusion, artery ligature, and distal occlusion).
8) Irrigate the femoral artery with 2% lidocaine to dilate the vessel and prevent
9) Apply tension to the proximal and distal sutures to occlude blood flow and elevate the
10) Pierce the artery with catheter introducer (bent-tip, 22g. needle) cranial to the distal
occlusion suture and slip the catheter tip proximally into the vessel.
11) Temporarily release tension on the proximal occlusion suture and slide the catheter
12) Advance the catheter beyond the iliac bifurcation until the pressure-sensing tip is
13) Tie the middle, ligature suture around the artery so that it seals the artery wall around
the catheter stem. Release the proximal occlusion suture and observe for leakage. A small drop of veterinary tissue adhesive may be applied to catheter insertion point to seal the juncture.
14) Release the tension on the distal occlusion suture and use it to ligate the downstream
section of the femoral artery. Once the ligature is knotted around the artery, tie the suture ends around the catheter stem to stabilize the preparation.
15) To keep the catheter in the proper orientation, suture the catheter stem to surrounding
muscle tissue and loop the catheter subcutaneously to prevent kinking.
16) Close the incision with sutures or staples and seal the incision with sterile tissue
adhesive® to help prevent infection. Skin clips and non-absorbable suture materials should be removed 7-10 days post-operatively.
17) Administer the appropriate post-operative analgesic and monitor the animal for return
B. Telemetry Unit Insertion Procedure – Abdominal aorta catheter/ peritoneal cavity
This is the procedure of choice when use of femoral arteries is prohibited due to study needs and/or when accurate core body temperature is required. All surgical procedures must follow the Guidelines for Rodent Survival Surgery at Stony Brook University.
1) Shave and disinfect the ventral abdomen from the xiphoid process to the pelvis and
drape surgical site. Disposable, adhesive drapes are preferred for rodents.
2) Make a 4-10 cm midline abdominal incision to allow good visualization of the
peritoneal cavity from the renal arteries down to the aortic bifurcation.
3) Expose the contents of the abdomen using a spring retractor. 4) Hold back the intestines using saline moistened gauze sponges to allow good
visualization of the descending aorta on the dorsal body wall.
5) Gently dissect the aorta from the surrounding fat and connective tissue using sterile
6) Clear excess tissue from the aorta to allow for good hemostasis following
7) Carefully insert an occlusion ligature (2-0 non-absorbable suture) between the aorta and
the vena cava, just caudal to the left dorsal muscular branch. Form a loop under the aorta that will allow occlusion of blood flow. Care should be taken not to damage the vessel as this may result in thrombosis or fibrosis of the catheter tip later.
8) Restrict the blood flow using the proximal ligature to elevate the vessel in preparation
for catheterization. NOTE: The aorta blood flow must not be restricted for greater than 3-4 minutes to avoid hind-limb paralysis due to ischemia.
9) Puncture the aorta just cranial to the aortic bifurcation using a 21-gauge needle bent 90°
at the beveled end. Make sure the concave surface of the bevel faces down, and do not place more than the beveled needle tip into the blood vessel.
10) Slide the tip of the catheter under the needle using the needle as a guide (or use a
catheter introducer), and pass the catheter cranial until the entire thin-walled section is within the vessel.
11) Thoroughly dry the puncture site and surrounding tissue to assure complete bonding of
the tissue adhesive. Apply one drop of sterile tissue adhesive to the puncture site and place a cellulose patch on the glue. Lift catheter to allow complete glue penetration.
12) Replace intestines into their original position and moisten with sterile saline. 13) Simultaneously close the body wall and secure the transmitter into place by closing the
abdominal incision. Incorporate the “suture rib” on the telemetry unit into the closure using non-absorbable sutures (3-0 or 4-0) in a simple interrupted pattern.
14) Close the incision with sutures or staples and seal the incision with sterile tissue
adhesive to help prevent infection. Skin clips and non-absorbable suture materials should be removed 7-10 days post-operatively.
15) Administer the appropriate post-operative analgesic and monitor the animal for return
C. Telemetry Unit Insertion Procedure – Carotid catheter/ subcutaneous transmitter
This procedure may be used as an alternate to catheter placement in the descending aorta or
femoral artery, depending on study objectives. All surgical procedures must follow the
Guidelines for Rodent Survival Surgery at Stony Brook University.
1) Shave and disinfect the ventral throat area and drape surgical site. Disposable, adhesive
2) Make a midline incision from the sternum to the jaw (about 2.5 cm). Retract the
salivary glands to expose the muscles of the trachea.
3) Locate the carotid artery along the left side of the trachea and carefully isolate the
4) Pass two lengths of non-absorbable suture material underneath the isolated section of
5) Position one suture just proximal to the bifurcation of the external and internal carotid
6) Position the other suture close to the clavicle and apply tension to elevate the artery and
7) Pierce the vessel just below the point of ligation with a bent-tipped 25g needle and
insert the catheter. Advance the tip at least 2 mm so that the catheter notch is near the carotid bifurcation. Tie in the catheter using the two sutures.
8) Use blunt tipped dissecting scissors to form a subcutaneous pouch to hold the
transmitter. The location of the pouch will depend on the species and vary from the neck (large animal) to the lateral flank (mouse). Insert the transmitter into the pouch and secure with sterile tissue adhesive.
9) Close the skin incision with sutures or staples. Skin clips and non-absorbable suture
materials should be removed 7-10 days post-operatively.
10) Administer the appropriate post-operative analgesic and monitor the animal for return
D. Bio-potential Lead Placement for ECG/EMG recording
This ECG/EMG procedure may also be performed for some studies following the placement of catheter and transmitter by one of the methods described above. All surgical procedures must follow the Guidelines for Rodent Survival Surgery at Stony Brook University.
1) For ECG/EMG lead placement, the skin overlying the lead placement sites are shaved
and disinfected when the animal is prepared for catheterization as described above. The ECG sites are the right cranial and left medial thorax; the EMG site is study dependent.
2) A small incision is made in the skin overlying the lead placement sites. The lead wires
are tunneled subcutaneously to their placement sites and attached to the muscle using 4-0 non-absorbable sutures. In mice, the leads may simply be tunneled but not sutured to the muscle.
3) Close the incision with sutures or staples. Skin clips and non-absorbable suture
materials should be removed 7-10 days post-operatively.
4) Administer the appropriate post-operative analgesic and monitor the animal for return
B. Equipment malfunctions/recovery
If the catheter, bio-potential leads, or transmitter cease to work, attempts will be made to diagnose the problem. If it can be determined that the malfunction can be corrected by simple adjustments to the positioning or the placement of the equipment, a subsequent surgery may be conducted to accomplish this goal. This will occur only if the additional survival surgery is deemed justified by the clinical veterinarian, after consultation with the principal investigator. A decision to perform recovery surgery will be made based on the value of the research animal due to historical data already obtained and the adequacy of the battery charge remaining on the transmitting unit. All surgical procedures must follow the Guidelines for Rodent Survival Surgery at Stony Brook University. Following euthanasia, all equipment will be removed, cleaned with enzymatic detergent, soaked overnight in 2% activated glutaraldehyde, and thoroughly rinsed in sterile saline before re-use. Non-functioning units or those with depleted batteries will be returned to the vendor for reconditioning.
C. Post-operative Analgesia
Analgesia will be provided under the direction of the Clinical Veterinarian but will include at least a single post-operative dose of Buprenorphine (Mice – 0.01 - 0.05 mg/kg SQ, Rats – 0.1-0.5 mg/kg SQ) or Ketorolac (4 mg/kg SQ). Additional doses will be administered as needed until the animal is not showing any signs of pain (inappetance, poor grooming, hunched posture, etc.).
D. Adverse Effects
1) Potential adverse effects from this procedure are minimal but may include:
• Anesthetic- related respiratory distress • infection of the subcutaneous pocket, abdominal cavity or catheter insertion
• dehiscence of the surgical site • seroma formation around the transmitter • hind limb paresis or paralysis related to ischemia or nerve damage • hemorrhage due to leaking of the vessel around the catheter insertion site • post-operative pain as evidenced by:
• decreased activity • decreased food and water intake • rough hair coat, hunched posture • weight loss • vocalization when the incision is touched • isolation from the individual from the group
2) Adverse effects can be prevented or limited by trained surgeons using aseptic
surgical techniques and the proper choice of suture materials, as described above.
E. Clinical Monitoring and Management
1) Animals will be monitored daily until the skin sutures or staples are removed (7-10
days), and then once to twice weekly until completion of the study.
2) Post -operative monitoring sheet must be used (request DLAR for monitoring sheet
if needed) and this will assist investigators with pain assessment.
3) If any adverse effects are seen, the Clinical Veterinarian will be informed
immediately and appropriate treatments administered. Animals not responding to treatment will be euthanized.
F. Early Endpoints
1) Animals that are showing signs of pain or distress including inappetance, hunched
posture, poor grooming, weight loss of more than 15%, or difficulties locomoting will be removed from study and reported to the clinical veterinarian. If deemed untreatable, these animals will be euthanized.
2) Animals showing swelling or discharge from the incision site or dehiscence will be
reported to the clinical veterinarian. If deemed untreatable, these animals will be euthanized.
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