Dose Administration: It is the optimal delivery of the dose into laboratory animals by a wide variety of routes.
Oral Administration in Rat or Mouse
Calculate the maximum volume to be administered to the animal. Recommended maximum dose volume is 10 ml/kg body weight (b.wt.) and in case of aqueous solution 20 ml/kg b.wt. may be used. Select the oral gavage needle from details mentioned in table.
Animal will be restrained firmly by gripping a fold of skin from the scruff of neck down the back, straighten the head and when the neck extended position is vertical it creates a straight line through the neck and esophagus. Prior to performing the oral dosing, distance (outside of the restrained animal) will be measured from the oral cavity to the end of the Xiphoid process with the gavage needle, this will indicate how far the gavage needle will be inserted into the esophagus. The gavage needle will slide down the esophagus from the back of the tongue if require rotates slightly as it passes the epiglottis. Once the gavage needle is into the premeasured distance, dose will be slowly administered to minimize the dose coming back from the esophagus.
If the animal gasps for air, then it may be an indication that the gavage needle is inserted into the larynx or trachea. If so, withdraw the gavage needle and re-insert. Do not force the gavage needle down the esophagus, this may cause tears in the esophagus and injury to the animal.
Table 1: Gavage Needle Size for Use in Mouse and Rat
Parenteral Administration
Subcutaneous Administration (SC) in Rat, Mouse, or Rabbit
Recommended maximum dose volumes are 10, 5 and 1 mL/kg b. wt. for mouse, rat, and rabbit, respectively. The preferred syringe needle size is 24 – 27 Gauge. Firmly restrain the animal, swab the injection site with 70% alcohol and lift the skin (loose skin between the shoulders, abdomen, or flank) to form a tent (Image 3). Insert needle (bevel up) at the base of the tent (close to the body), parallel to the animal’s body to avoid puncturing the underlying structures. Gradually inject the dose. A bubble (bleb) may be felt at the injection site, after dose administration gently removes the needle. Wipe the injected site with 70% alcohol.
Intraperitoneal Administration (IP) in Rat or Mouse
The preferred syringe needle size is 23 – 27 Gauge. Recommended maximum dose volumes are 20 and 10 ml/kg body weight for mouse and rat respectively. Manually restrain the animal and hold it in supine position with its posterior end elevated, or the head can be titled lower than the body. Swab the injection site with 70% alcohol. The needle shall be pushed in the lower quadrant of the abdomen. Proper insertion yields no blood on aspiration, if blood is seen in the tip of the syringe then carefully remove the needle and try in different site, again ensure by aspiration. Gradually inject the dose.
Intravenous Administration (IV) in Rat and Mouse
26-30G syringe needle will be used for IV dosing. • Recommended maximum dose volume is 5 mL/kg body weight for rat and mouse. Suitable size restrainer will be used to ensure that the animal is comfortable. Animal will be held by its tail and gently pull into the restrainer. Tail vein will be dilated with warm water vein in case the vein is not prominently visible. Tail will be disinfected with 70% alcohol. Once the lateral tail vein is visualized, a needle will be inserted slowly into the vein. Slight back pressure will be applied to pull blood in the syringe to confirm proper placement in the vein before injecting. If blood is not seen in the syringe, then carefully remove the needle, press the punctured site gently with 70% alcohol swab to stop bleeding and try again by slightly moving towards the proximal end of the tail. The dose will be administered gradually.
Intramuscular Administration (IM)
The hind leg quadriceps muscles shall be selected as the site for intramuscular administration of dose. Recommended maximum dose volumes are 0.05 mL/Site, 2 sites/day for mouse, 0.2 mL/Site, 2 sites/day for rat & guinea pig and 0.5 ml/kg (Max 1 ml limit) for rabbit. Clean the area with 70% alcohol. Gently insert the needle into the muscle at a 90-degree angle, being careful not to hit any bones or nerves. Aspirate the syringe, blood in the syringe indicates improper placement, if so, reposition the needle. Gradually administered the dose. Remove the needle & apply gentle pressure to the injection site with a cotton ball or gauze to stop any bleeding. Again, wipe the area with 70% alcohol.
Dermal (Topical) Application
For Rat
Approximately before the 24 hours of the test, fur should be removed by closely clipping (at least 10% body surface area) the dorsal area of the trunk. Apply the dose uniformly over the clipped area. If direct application is not possible, the dose should first be applied to the gauze patch, which is then applied to the skin. Hold the dose in contact with the skin with a porous gauze dressing and non-irritating tape in a suitable manner and ensure that the animal cannot ingest and inhale the dose.
For Rabbit
Approximately 24 hours before the dosing, fur shall be removed by closely clipping the dorsal area of the trunk of the animals. Apply the dose to a clipped area (approximately 6 cm X 6 cm) of skin and cover with a gauze patch, which will be held in place with non-irritating tape. If direct application is not possible, the dose should first be applied to the gauze patch, which is then applied to the skin. The patch shall be loosely held in contact with the skin by means of a suitable semi-occlusive dressing for the duration of the exposure period and ensure that the animals cannot ingest or inhale the dose.
For Guinea Pig
Approximately 24 hours before the application of the test patch system both the flank (right and left) should be cleared of hair (closely clipped). The test patch system fully loaded with test item should be applied to the test area and held in contact with the skin by an occlusive patch for exposure period. 2X4 cm size filter paper and 4X4 cm size cotton pad shall be used for Guinea Pig Maximization Test and Buehler Method respectively.
Intradermal Injection for Guinea Pig
Recommended dose volume of injection is 0.1 mL per injection site. The recommended needle size is 26 Gauge. Approximately 24 hr before exposure, the test site should be clipped. Restrain the guinea pig by the handler’s thumb from the neck to under the jaw of the guinea pig and supported by the handler’s other hand on the hind limb of the animal. Insert the needle (bevel up) just under the surface (clipped area of scapular region) of the skin and inject the dose. Distinct bleb should form.
Intranasal Instillation
Maximum Recommended dose volume is 100 µL/animal for rat and 50 µL/animal for mice. Anaesthetize the animal by isoflurane, and gently hold the animal in one hand and put the tip of the micropipette with the other hand near the nostril and slowly release the test item through the micropipette. So, animal will slowly take the drugs without any struggle. Hold the animal in vertical procedure for a few seconds to complete instillation of drugs. After dosing keep the animals in their respective cage.
Usage of Anaesthesia in Laboratory animals
Intrathecal Injection in rat
The skin dorsal to the lumbar and sacral spine of the animals shall be clipped free of fur. Animals shall be anesthetized with 5% isoflurane, and 2% isoflurane shall be maintained during the surgical procedure. The area of surgery shall be wiped with 70% Ethanol before injection. The injections shall be performed by holding the rat securely in one hand by the pelvic girdle and inserting a 26-G needle, into the tissues between the dorsal aspects of L3 and L4, perpendicular to the vertebral column. Syringe shall be inserted into the spinal subarachnoid space through the site in a cephalad direction at an angle of about 45° to the sagittal plane. Successful IT location of the needle shall be confirmed by the presence of at least one of the following signs: twitch of the tail and/or presence of cerebrospinal fluid (CSF) in the needle hub. If none of these signs was seen, or if blood is visible in the needle hub, the needle shall be withdrawn, and the operation shall be repeated with another needle. These reflexes are used as an indicator of successful puncture. No other specific behaviour or sign of distress or pain shall be observed at this time.
Intrauterine Administration
All the animals shall be fasted for 16 hours (if required), water shall be provided. Prior to surgery animals shall be anesthetized by using a combination of Ketamine and Xylazine injection, care shall be taken for asepsis using 70% ethanol as antiseptic and disinfectant. The fur shall be clipped with a clipper at the selected surgery area. Both sides of flanks near the hump of the rat shall be shaved by hair clipper.
Prior to surgery ensure that all surgical items shall be autoclaved for maintaining sterility. The area of surgery shall be wiped with 70% Ethanol before making incision. Both sides of flanks shall be sterilized with a Betadine dipped gauze sponge and after area shall be cleaned with Deionized (DI)/Milli Q water. The skin layer shall be cut with Adson and 12 cm blunt scissors along the abdominal line. Then a nick cut shall be made on the muscle layer with 12 cm straight sharp scissors to form a small hole. Forceps shall be used to find the uterus tube surrounded by a variable amount of fat. The fallopian tube will be pulled out through the muscle incision. The required amount of test item will be injected slowly into the uterus. After dosing the fallopian tube shall be placed in its normal place. Bleeding is usually slight and inconsequential and soon stops on its own. Stitch the open area with the help of suture.
After surgery and prior to recovery from anesthesia, warmed sterile fluids (saline or lactated ringers’ solution) shall be given by subcutaneous route.
Post Operative care:
After dosing the rat shall be placed back into the cage and need to keep warm by room heaters (2 to 3 hours). Then, the animals shall be housed individually provided with clean and dry bedding sets made of 100% sterilized cotton fabric for extra comfort and warmth for a period of one week to avoid hypothermia and to prevent possible contamination. Animals shall be monitored for the continued need for analgesics, and observations will be made at least twice daily in the first three days postoperative.
Analgesic shall be used based on the half-life. Buprenorphine 0.01 - 0.05 mg/kg or Meloxicam 0.3 – 1.0 mg/kg body weight by S.C route shall be given for at least three days.
Subretinal Administration or injection
Injection shall be performed in a sterilized room. Anesthesia shall be given. Check the depth of anesthesia by simply testing the animal response to gentle pressure on the hind paws.
Dilate the pupil with an eye drop of diluents (ex: bromide or phenylephrine 2.5% eye drops) and Proparacaine 0.5% applied for topical anesthesia. Fill the required volume of test item or vehicle in the microliter syringe (preferably Hamilton syringe). Open the eyelid and protrude the eye to expose the equator for convenient injection. Maintain the protruded eye position until finishing the injection, (if not, displacement of the needle can occur during the injection). To hold the eyeball firmly, place the fingers outside the orbital rim.
Puncture a small hole at slight posterior to the limbus using a sterile gauge needle (needle size between 24- 28 G ½) for the further subretinal injection. Make the hole inferior for the right eye, and superior for the left eye for convenience.
Place the blunt needle of the microliter syringe through the pre-punctured hole and approach the needle into the subretinal space until the point when mild resistance is felt (needle size between 28 - 36 gauge). Inject the dose volume (maximum 3µL volume or as specified in study plan) gently into the subretinal space without tremor to avoid unwanted tissue damage and withdraw the needle gently.
Observe the formation of subretinal bleb after the injection under an operating microscope to make sure there is no retinal bleeding. Gently close the eyelid to cover the injection site for self-sealing. Return the animal to the home cage and keep it alive until the evaluation.
Post-operative care:
After subretinal injection, the animals shall be placed back into the fresh cage and need to be kept warm by room heater (1 to 3 hours). Eyes shall be kept wet until animal recovery from anesthesia by using hydroxypropyl methylcellulose eye drops to avoid the dry. The animals shall be housed individually in respective cages and provide fresh sterilized bedding material to prevent possible contamination. Animals shall be monitored.
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