Handling Instructions

Injection Assembly:
Injection assembly consists of an infusion cannula attached to 18 inches of Tygon tubing. A 10uL Hamilton syringe is attached to the injection assembly.

Standard Coordinates:
  • Rats (>175g): AP - 4.3mm, ML +/- 1.2mm (+ for left side lesions and - for right side lesions), DV - 8.3mm.
Housing:
Animals can be group housed.

Staple Removal:
Staples should be removed 7-10 days post-operatively.

Notes:
  • Sham animals are identified with a right ear punch unless specified otherwise.
  • Default is left side lesion unless client requests right side.
  • Animals are ear tagged and apomorphine challenge rotational scores are provided.
  • Desipramine is administered to all animals 30 minutes prior to 6-OHDA infusion.
Drinking Supplements:
Adrenalectomized animals are maintained and shipped with 0.9% Sodium Chloride drinking water, ad lib. Certain species of mouse may require a corticosteroid to be added to the drinking supplement to increase survivability.

Housing:
Animals can be group housed.

Staple Removal:
Staples should be removed 7-10 days post-operatively.

Notes:
  • Sham animals are identified with a right ear punch and are maintained on normal drinking water unless specified otherwise.
Catheters:
The catheter material consists of a length of sterile 3.5 Fr polyurethane (PU) tubing with a PE10 tip. Fill volume of each catheter is 60µl.

Catheter Maintenance:
To increase the longevity of this preparation, the duodenal catheter and the stainless steel tubing should be flushed every two to three days with 0.2-0.5mL sterile distilled water. Never flush the bile duct catheter.

Animal Maintenance:
It is recommended the animals also receive SQ fluids and analgesics as needed for maintenance; wet feed may be added to encourage eating.

Expectations:
Surgery occurs within a week of shipment. Investigators should expect the bile duct preparation to be patent upon arrival and last for roughly one week post receipt, pending proper maintenance as described in the quality control section above. Bile duct catheterizations are not stable for long periods of time and should be used as close to the date of receipt as possible. There will be a yellow color to the catheters if bile is flowing into the return. A greenish color indicates salt build-up and flushing the duodenal catheter is required.

Sampling:
During our in-house bile collection procedures, bile flows at an average rate of 1ml/hour. Collection of bile is achieved by attaching the animal to a spring tether device. During collection, be sure to plug the duodenal catheter with a solid pin to prevent digestive contents from backing into the catheter. Collection can occur for extended periods of time (up to 12 hours). After sampling periods have concluded, the duodenal catheter is flushed with dH2O and the animal should be allowed to recover for a minimum of two days before sampling is repeated. Subcutaneous saline should be administered at a rate of 10mL/kg before and after each collection period. Analgesics are also administered as needed.

Housing:
Individually house animals to prevent cage mates from chewing on one another's catheters.

Staple Removal:
Staples should be removed 7-10 days post-operatively; do not remove staples around catheter port.
Catheter:
Catheter material consists of sterile 3.5 Fr polyurethane (PU) tubing with a 2 Fr PU tip. The catheter is sealed with a sterile stainless steel pin. 22 gauge blunted needles are required to access the port. Fill volume of the catheter is 40µl.

Lock Solution:
Heparinized Glycerol (500 IU/ml): 10.0 mL stock heparin (1000 IU/mL) + 10.0 mL 99% Glycerol solution (Sigma).

Catheter Maintenance:
To maintain animals over longer periods of time, catheters need to be flushed twice a week (once every 3-4 days). Catheters can be flushed by following the sampling procedure below minus the withdrawal of the whole blood sample.

Blood Pressure Monitoring & Sampling:
CAC are reliable for obtaining blood samples and blood pressure readings. Sample size and frequency should be minimized to essential time points to maintain the health of the animal. Pilot studies are always encouraged to be sure our CAC preparation performs up to user expectations. For blood withdraw, gather the following materials: Syringe assemblies (1cc syringe attached to 22G blunted needle), sterile saline and sterile lock solution.

  1. Place animal in a restrainer (small open topped boxes the size of a pipette container work well).
    Important: Always clamp the port with rubberized or smooth hemostats to prevent unintended blood flow and port damage when changing syringes and flushing the catheter.
  2. Clamp port and remove the pin from the catheter and set aside.
  3. Insert an empty syringe assembly (SA) into the port and release hemostats.
  4. Gently withdraw fill solution and blood; clamp port.
  5. Attach a second SA, release hemostats and withdraw sample (syringe may contain anticoagulant); clamp port.
  6. Release hemostats and slowly flush catheter with sterile saline ~ 100µl (or greater to match blood withdraw) clamp port.
  7. Add 40µl of sterile lock solution, clamp port and replace pin.
  8. If blood fails to flow in step 4, remove the empty SA and replace with a SA containing sterile saline and gently flush the catheter and repeat as outlined above.

Housing:
Individually house animals to prevent cage mates from chewing on one another’s catheters.

Staple Removal:
Staples should be removed 7-10 days post-operatively. Do not remove the staple around the port.

Notes:
  • During animal manipulation, it is important not to place undo stress on the catheter.
  • Using needles larger than 22 gauge will stretch the port and decrease the longevity of the preparation. Additionally, accessing the port by way of needles with bevels or rough edges will damage the port, again making sampling / use difficult.
Catheter:
Catheter material consists of sterile 3 Fr polyurethane (PU) tubing with a 1 Fr tip. The access port is sealed with a sterile stainless steel pin. 24 gauge blunted needles are required to enter the port. Fill volume of the catheter is 10µl.

Lock Solution:
Heparinized Dextrose (500 IU/ml): 10.0 mL stock heparin (1000 IU/mL) + 10.0 mL Dextrose.

Catheter Maintenance:
To maintain animals over longer periods of time, catheters need to be flushed twice per week (once every 3-4 days). Follow the sampling procedure outlined below, minus sample withdrawal to flush catheters. For best results, use the animals within one week of delivery.

Sampling / Test Article Administration:
CAC are reliable for obtaining blood samples and blood pressure readings. Sample size and frequency should be minimized to essential time points to maintain the health of the animal. Pilot studies are always encouraged to be sure our CAC preparation performs up to user expectations. For blood withdrawal, gather the following materials: Syringe assemblies (1cc syringe and microsyringe syringe attached to 25G blunted needle), sterile saline and sterile lock solution.


  1. Place mouse in a restrainer.
    Important: Always clamp the port with rubberized or smooth hemostats to prevent unintended blood flow and port damage when changing syringes and flushing the catheter.
  2. Clamp port and remove the pin from the catheter and set aside.
  3. Insert an empty syringe assembly (SA) into the port and release hemostats.
  4. Gently withdraw fill solution and blood to the point of seeing blood in the needle hub; clamp port.
  5. Attach a second SA, release hemostats and withdraw sample (syringe may contain anticoagulant) clamp port.
  6. Release hemostats and slowly flush catheter with sterile saline ~10µl) clamp port.
  7. Attach microsyringe and refill with 10µl lock solution – avoid overfilling the lock solution! Clamp port and replace pin.
  8. If blood fails to flow in step 4, remove the empty SA and replace with a SA containing saline. Gently flush the catheter with 10µl of saline and repeat as outlined above.

Housing:
Individually house animals to prevent cage mates from chewing on one another’s catheters.

Note:
  • Using needles larger than 25G will stretch the port and make future sampling difficult. Additionally, employing needles with bevels or rough edges will damage the port – decreasing the longevity of the catheter.
Housing:
Animals can be group housed. Barbering is common in male mice; enrichment is added to lessen the chance of occurrence.

Staple Removal:
Staples should be removed 7-10 days post-operatively.

Note:
  • Sham animals are identified with a right ear punch unless specified otherwise.
Cannula:
Either a cannula provided by the client or our standard cannula may be placed.

Coordinates:
Coordinates used are provided by the client and are calculated in relation to bregma. Some variation occurs depending on the size and strain of the animal.

Checking cannula placement after completion of studies is recommended.

Compound Administration:
If our standard cannula is used, Injector Cannulas will be provided with shipment. Injector cannulas are required to administer test articles via the guide cannula. Single injections should not exceed a rate of 1µl/min and a total volume of 5µl. Additional injections may be given after a washout period. Injector cannulas are typically attached to micro-syringes via a length of polyethylene tubing (PE 50). Injection is achieved by unscrewing and removing the dummy cannula and inserting an injector cannula until it locks in position. Connector assemblies from Plastics One can also be used for administration via infusion pumps.

Housing:
Ideally, animals are individually housed in solid caging to minimize chances cannulas will be dislodged in wire bars of caging and lids. Group housing is adequate as long as cage mates refrain from chewing on one another’s cannulas. With appropriate care, cannulas are expected to last at least three weeks.

Note:
  • Mice should be lightly sedated prior to cannula manipulation.
Catheter:
Catheter material consists of sterile polyurethane (PU) tubing. The body of the catheter is made of 3.5 Fr tubing and the insertion tip is 2 Fr tubing. The access port is sealed with a sterile stainless steel pin. 22 gauge blunted needles are required to access the port. Fill volume of the catheter is 80µl.

Lock Solution:
Heparinized Glycerol (500 IU/ml): 10.0 mL stock heparin (1000 IU/mL) + 10.0 mL 99% Glycerol solution (Sigma).

Catheter Maintenance:
To maintain animals over longer periods of time, catheters need to be flushed twice a week (once every 3-4 days). Catheters can be flushed by following the sampling procedure below minus the withdrawal of the whole blood sample.

Blood Pressure Monitoring & Sampling:
FAC are reliable for obtaining blood samples and blood pressure readings. Sample size and frequency should be minimized to essential time points to maintain the health of the animal. Pilot studies are always encouraged to be sure our FAC preparation performs up to user expectations. For blood withdrawal gather the following materials: Syringe assemblies (1cc syringe attached to 22G blunted needle), sterile saline and sterile lock solution.

  1. Place animal in a restrainer (small open topped boxes the size of a pipette container work well).
    Important: Always clamp the port with rubberized or smooth hemostats to prevent unintended blood flow and port damage when changing syringes and flushing the catheter.
  2. Remove the pin from the catheter and set aside.
  3. Insert an empty syringe assembly (SA) into the port and release hemostats.
  4. Gently withdraw fill solution and blood; clamp port.
  5. Attach a second SA, release hemostats and withdraw sample (syringe may contain anticoagulant) clamp port.
  6. Release hemostats, slowly flush catheter with sterile saline ~200µl (or greater to match blood withdraw) and clamp port.
  7. Add 80µl of sterile lock solution, clamp port and replace pin.
  8. If blood fails to flow in step 4, remove the empty SA and replace with a SA containing sterile saline and gently flush the catheter and repeat as outlined above.

Housing:
Individually house animals to prevent cage mates from chewing on one another’s catheters.

Staple Removal:
Staples should be removed 7-10 days post-operatively. Do not remove the staple around the catheter port.

Note:
  • During animal manipulations, it is important not to place undue stress on the catheter.
  • Using needles larger than 22 gauge will stretch the port and decrease the longevity of the preparation. Additionally, accessing the port by way of needles with bevels or rough edges will damage the port, again making sampling / use difficult.
Catheter:
Catheter material consists of sterile 3.5 Fr polyurethane (PU) tubing. The access port is sealed with a sterile stainless steel pin. 22 gauge blunted needles are required to enter the port. Fill volume of the catheter is 80µl.

Lock Solution:
Heparinized Glycerol (500 IU/ml): 10.0 mL stock heparin (1000 IU/mL) + 10.0 mL 99% Glycerol solution (Sigma).

Catheter Maintenance:
To maintain animals over longer periods of time, catheters need to be flushed twice a week (once every 3-4 days). Catheters can be flushed by following the sampling procedure below, minus the withdrawal of the whole blood sample.

Dosing/Sampling:
FVC’s are utilized primarily for sterile test article administration. Administration is achieved by bolus dosing or constant infusion. Rates of infusion should not exceed 1ml per minute. To a lesser extent, femoral vein catheters can be utilized for obtaining blood samples. 200-300 gram rats can tolerate 1000 to 1500µl of total blood withdrawal in a 24 hour period. Sample size and frequency should be minimized to essential time points to maintain the health of the animal. For test article administration or blood withdraw, gather the following materials: Syringe assemblies (1cc syringe attached to a 22G blunted needle), sterile saline and sterile fill solution.

  1. Place animal in a restrainer (small open topped boxes the size of a pipette container work well).
    Important: Always clamp the port with rubberized or smooth hemostats to prevent unintended blood flow and port damage when changing syringes and flushing the catheter.
  2. Clamp port and remove the pin from the catheter and set aside.
  3. Insert an empty syringe assembly (SA) into the port and release hemostats.
  4. Gently withdraw fill solution and blood; clamp port.
  5. Attach a second SA, release hemostats and withdraw sample or administer test article (syringe may contain anticoagulant) clamp port.
  6. Release hemostats and slowly flush catheter with sterile saline ~200µl (or greater to match blood withdrawal) clamp port.
  7. Add 80µl of sterile lock solution, clamp port and replace pin.
  8. If blood fails to flow in step 4, remove the empty SA and replace with a SA containing sterile saline and gently flush the catheter. Continue as outlined above.

Housing:
Individually house animals to prevent cage mates from chewing on one another’s catheters.

Staple Removal:
Staples should be removed 7-10 days post-operatively. Do not remove the staple around the port.

Notes:
  • During animal manipulations (dosing / weighing), it is important not to place undue stress on the catheter.
  • Using needles larger than 22 gauge will stretch the port and make sampling difficult. Additionally, sampling by way of needles with bevels or rough edges will damage the port, again making sampling difficult.
Catheters:
Intestinal catheter material consists of a length of sterile 3.5 Fr polyurethane (PU) tubing for rats and 3 Fr PU tubing for mice. The access ports are sealed with a sterile stainless steel pin. For the rats, 22 gauge, blunted needles are required to access the port and 24 gauge are required for the mice.

Lock Solution:
Sterile distilled water (dH2O).

Catheter Maintenance:
To increase the longevity of this preparation the intestinal catheter should be flushed every two to three days with 200-500µL sterile distilled water.

Test Article Administration:
Intestinal catheterizations are reliable for administration of test articles, provided the catheter is flushed two to three times per week with sterile distilled water. To access the port, collect the following materials: syringe assemblies (1cc syringe attached to a 22 or 24G blunted needle) and sterile distilled water.

  1. Clamp the port with rubberized or smooth hemostats to prevent unintended material flow and port damage when changing syringes and flushing the catheter; remove the pin from the catheter.
  2. Insert a syringe assembly (SA) containing dH2O and release hemostats.
  3. Gently infuse solution to be sure the catheter is patent; clamp port.
  4. Attach a second SA; release hemostats and administer test article.
  5. Clamp port; attach a new SA and slowly flush catheter with sterile dH2O: ~100µl.
  6. Clamp port and replace catheter pin.

Recommended maximum infusion rate:
2000µl per minute.

Housing:
Individually house animals to prevent cage mates from chewing on one another’s catheters.

Notes:
  • During animal manipulation it is important not to place undue stress on the catheter.
  • Using needles larger than recommended above will stretch the port and decrease the longevity of the preparation. Additionally, accessing the port by way of needles with bevels or rough edges will damage the port, again making sampling / use difficult.
Catheter:
Catheter material consists of sterile polyurethane tubing. Insertion length is determined by animal’s target size. Access ports are trimmed to 25mm and can be accessed with 25G blunted needles. Fill volume of the catheter is approximately 5µl. Maintenance flushing of the catheter is not required.

Compound Administration:
This catheter is useful for the introduction of material (novel test compounds, cells, etc.) but is not designed for sampling Cerebrospinal Fluid. Remove the stainless steel pin and replace with a Hamilton microsyringe. Test article administration via the catheter should not exceed the recommended rate of 10µl/min and a total dose volume of 30µl. Additional injections may be given after a washout period.

Housing:
It is important to individually house animals in order to minimize chances of catheter disruption by cage mates chewing on one another’s catheters.

Notes:
  • Catheters are inserted to match the age / size of the animal at surgery. Accordingly, Taconic recommends use of ITC animals as close to receipt as possible.
  • Checking catheter placement and integrity after completion of studies is recommended.
Catheter:
Catheter material consists of a sterilized polyethylene tubing with a silicone rubber intra-vascular tip. The access port consists of a 20mm length of PE50 tubing (0.023" ID) that is sealed with a sterile stainless steel pin. Fill volume of the catheter is 5µl. Access the port via a 23G blunted needle.

Lock Solution:
Heparinized Dextrose (500 IU/ml): 10.0 mL stock heparin (1000 IU/mL) + 10.0 mL Dextrose.

Catheter Maintenance:
To maintain animals over longer periods of time, catheters need to be flushed twice per week (once every 3-4 days). Follow the sampling procedure outlined below, minus sample withdrawal to flush catheters. For best results, use the animals within one week of delivery.

Sampling / Test Article Administration:
JVC are reliable for administration of test articles and some blood sampling, provided the mice are 25 grams or larger. To access the port, collect the following materials: syringe assemblies (1cc syringe attached to a 23G blunted needle), a microsyringe, sterile saline and sterile fill solution.

  1. Place mouse in a restrainer or sedate the mouse with a gas or injectable anesthetic.
    Important: Always clamp the port with rubberized or smooth hemostats to prevent unintended blood flow and port damage when changing syringes and flushing the catheter.
  2. Clamp port and remove the pin from the catheter.
  3. Insert an empty syringe assembly (SA) and release hemostats.
  4. Gently withdrawal fill solution to the point of seeing blood in the needle hub; clamp port.
  5. Attach a second SA and administer test article or withdrawal sample; clamp port.
  6. Release hemostats and slowly flush catheter with sterile saline ~100µl; clamp port.
  7. Attach microsyringe and refill with 10µl lock solution – avoid overfilling the lock solution!; clamp port and replace pin.
  8. If blood fails to flow in step 3, remove the empty SA and replace with a SA containing saline. Gently flush the catheter with 10µl of saline and repeat as outlined above.

Housing:
Individually house animals to prevent cage mates from chewing on one another’s catheters.

Note:
  • Using needles larger than 23G will stretch the port and make future sampling difficult. Additionally, employing needles with bevels or rough edges will damage the port – decreasing the longevity of the catheter.
Catheter:
Catheter material consists of sterile 3.5 Fr polyurethane (PU) tubing. The catheter is sealed with a sterile stainless steel pin. 22 gauge blunted needles are required to access the port. Fill volume of the catheter is 50µl.

Lock Solution:
Heparinized Glycerol (500 IU/ml): 10.0 mL stock heparin (1000 IU/mL) + 10.0 mL 99% Glycerol solution (Sigma).

Catheter Maintenance:
To maintain animals over longer periods of time, catheters need to be flushed twice a week (once every 3-4 days). Catheters can be flushed by following the sampling procedure below minus the withdrawal of the whole blood sample.

Sampling:
JVC’s are reliable for obtaining blood samples over the course of hours or even day(s). 200-300 gram rats can tolerate 1000 to 1500µl of total blood withdrawal in a 24 hour period; sample size can vary, but recommendations are in the range of 200-300µl. Sample size and frequency should be minimized to essential time points to maintain the health of the animal. For blood withdrawal, gather the following materials: Syringe assemblies (1cc syringe attached to a 22G blunted needle), sterile saline and sterile fill solution.

  1. Place animal in a restrainer (small open topped boxes the size of a pipette container work well).
    Important: Always clamp the port with rubberized or smooth hemostats to prevent unintended blood flow and port damage when changing syringes and flushing the catheter.
  2. Clamp port and remove the pin from the catheter and set aside.
  3. Insert an empty syringe assembly (SA) into the port and release hemostats.
  4. Gently withdraw fill solution and blood; clamp port.
  5. Attach a second SA, release hemostats and withdraw sample (syringe may contain anticoagulant); clamp port.
  6. Release hemostats and slowly flush catheter with sterile saline ~125µl (or greater to match blood withdrawal); clamp port.
  7. Add 50µl of sterile lock solution, clamp port and replace pin.
  8. If blood fails to flow in step 4, remove the empty SA and replace with a SA containing sterile saline and gently flush the catheter and repeat as outlined above.

Housing:
Individually house animals to prevent cage mates from chewing on one another’s catheters.

Staple Removal:
Staples should be removed 7-10 days post-operatively; do not remove staple around catheter port.

Notes:
  • During animal manipulation (dosing / weighing) it is important not to place undue stress on the catheter.
  • Using needles larger than 22 gauge will stretch the port and make sampling difficult. Additionally, sampling by means of needles with bevels or rough edges will damage the port, again making sampling difficult.
Cannula:
Guide cannulas consist of a length of 22-gauge (for rats) and 26-gauge (for mice) stainless steel hypodermic tubing encased in plastic (Plastics One, Roanoke, VA). Injector cannula consist of a 28-gauge stainless steel hypodermic tubing for rats and a 33-guage stainless steel tubing for mice.

Standard Coordinates:
    ICVC:
  • Rats (>150g) AP= -0.8 mm, ML = +1.2 mm (left side) and DV= -4.8 mm
  • Mice (>16g) AP= -0.2 mm, ML = +1.0 mm (left side) and DV= -2.3 mm
  • TVC:
  • Rats (>150g) AP= -1.6 mm, ML = 0.0 mm and DV= -8.0 mm
Compound Administration:
Injector Cannulas will be provided with shipment. Injector cannulas are required to administer test articles via the guide cannula. Single injections should not exceed a rate of 1µl/min and a total volume of 5µl. Additional injections may be given after a washout period. Injector cannulas are typically attached to micro-syringes via a length of polyethylene tubing (PE 50). Injection is achieved by unscrewing and removing the dummy cannula and inserting an injector cannula until it locks in position. Connector assemblies from Plastics One can also be used for administration via infusion pumps.

Housing:
Animals are individually housed in solid caging to minimize chances cannulas will be dislodged in wire bars of caging and lids and to prevent cagemates from chewing on one another’s cannulas. With appropriate care, cannulas are expected to last at least three weeks.

Notes:
  • Mice should be lightly sedated prior to cannula manipulation.
  • Checking cannula placement after completion of studies is recommended.
Materials:
Our standard Microdialysis materials are purchased from Bioanalytical Systems, Inc (BASi). MD-2250 (rubber O-ring fastener) and MD-2251 (metal Omega-ring fastener) are for rats, 10mm in length and sold as guide/stylet sets. Specify MD-2255 for mice (5mm length).

Compound Administration:
Appropriate microdialysis probes are available for purchase from BASi according to needs.

Housing:
Animals must be single housed to prevent cagemates from dislodging / damaging headmounts.
Housing:
Animals can be group housed.

Notes:
  • Sham animals are identified with a right ear punch unless specified otherwise.
Housing:
Animals can be group housed.

Staple Removal:
Staples should be removed 7-10 days post-operatively.

Notes:
  • Sham animals are identified with a right ear punch unless specified otherwise.
Catheters:
Catheter material consists of sterile 3.5 Fr polyurethane (PU) tubing with a 2 Fr PU tip. The catheter is sealed with a sterile stainless steel pin. 22 gauge blunted needles are required to access the port. Fill volume of the catheter is 90µl.

Lock Solution:
Heparinized Glycerol (500 IU/ml): 10.0 mL stock heparin (1000 IU/mL) + 10.0 mL 99% Glycerol solution (Sigma).

Catheter Maintenance:
To maintain animals over longer periods of time, catheters need to be flushed twice a week (once every 3-4 days). Flush catheters by following the sampling procedure below, minus the withdrawal of the whole blood sample.

Sampling:
Blood sample size can vary, but recommendations are in the range of 100-200µl. Sample size and frequency should be minimized to essential time points to maintain the health of the animal. For blood withdrawal, gather the following materials: Syringe assemblies (1cc syringe attached to a 22G blunted needle), sterile saline and sterile fill solution.

  1. Place animal in a restrainer (small open topped boxes the size of a pipette container work well).
    Important: Always clamp the port with rubberized or smooth hemostats to prevent unintended blood flow and port damage when changing syringes and flushing the catheter.
  2. Clamp port and remove the pin from the catheter and set aside. Insert an empty syringe assembly (SA) into the port and release hemostats.
  3. Gently withdraw fill solution and blood; clamp port
  4. Attach a second SA, release clamps and withdraw sample (syringe may contain anticoagulant); clamp port.
  5. Release hemostats and slowly flush catheter with sterile saline ~200µl (or greater to match blood withdrawal); clamp port.
  6. Add 90µl of sterile lock solution, clamp port and replace pin.
  7. If blood fails to flow in step 4, remove the empty SA and replace with a SA containing sterile saline and gently flush the catheter. Continue as outlined above.

Housing:
Individually house animals to prevent cage mates from chewing on one another’s catheters.

Staple Removal:
Staples should be removed 7-10 days post-operatively; do not remove staple around the catheter port.

Notes:
  • During animal manipulation (dosing / weighing), it is important not to place undue stress on the catheter.
  • Using needles larger than 22 gauge will stretch the port and make sampling difficult. Additionally, sampling by means of needles with bevels or rough edges will damage the port, again making sampling difficult.
Housing:
Animals can be group housed.

Staple Removal:
Staples should be removed 7-10 days post-operatively.

Note:
  • Sham animals are identified with a right ear punch unless specified otherwise.
Housing:
Animals can be group housed.

Staple Removal:
Staples should be removed 7-10 days post-operatively.

Note:
  • Sham animals are identified with a right ear punch unless otherwise requested.
Housing:
Animals can be group housed.

Staple Removal:
Staples should be removed 7-10 days post-operatively.

Note:
  • Sham animals are identified with a right ear punch unless otherwise requested.
Drinking Supplements:
Thyro-parathyroidectomized animals are maintained and shipped with 2% calcium lactate solution, ad lib.

Housing:
Animals can be group housed.

Staple Removal:
Staples should be removed 7-10 days post-operatively.

Note:
  • Sham animals are identified with a right ear punch and are maintained on normal drinking water unless specified otherwise.
Housing:
Animals can be group housed.

Staple Removal:
Staples should be removed 7-10 days post-operatively.

Note:
  • Sham animals are identified with a right ear punch unless specified otherwise.