Apparatus and Equipment

Precautions

References

1. Ehrenwerth, J., Eisenkraft, J. B., & Berry, J. M. (2013). Anesthesia equipment: Principles and applications. Philadelphia, PA: Elsevier Saunders.
2. Fish, R. E. (2008). Anesthesia and analgesia in laboratory animals. Amsterdam: Elsevier.
3. Križmarić, M. Functions of anesthesia reservoir bag in a breathing system. Retrieved from http://vestnik.szd.si/index.php/ZdravVest/article/view/2204/21564.67

Open Breathing Systems

T-Piece System

Bain Coaxial Breathing System

Magill Breathing System

Closed Breathing System

References

Rat Kit

Introduction

A brain microdialysis surgical kit is used for intracerebral microdialysis. Intracerebral microdialysis is a surgical technique primarily used for in vivo sampling of neurotransmitters. The method is widely used in neuroscience for sampling, assessment, and quantification of neuropeptides, hormones, drugs, and other molecules in the brain, periphery, and interstitial fluid. The technique is also applied for the investigation of the pharmacological effects of potent drugs on amino acid and monoamine neurotransmitters.

There are two basic types of brain microdialysis experiments namely conventional microdialysis and no-net-flux microdialysis. In the conventional microdialysis, neurotransmitter-free artificial cerebrospinal fluid (aCSF) is perfused through the cannula, and the neurotransmitter is collected in the dialysate. And, in the no-net-flux or zero-net-flux method, aCSF with several different concentrations of neurotransmitter is pushed through the probe and the amount of the analyte of interest increased or decreased in the probe is measured (Vladimir, Alexis, Agustin, & Shippenberg, 2009). Both methods are applied for brain microdialysis in rodents.

Microdialysis is prominently used for sampling of molecules from several organ systems including the blood, muscles, liver, eyes, etc. The history of the use of microdialysis as a sampling technique dates back to 1960s when it began with the push-pull method, which used a semi-permeable membrane to sample electrolytes and amino acids from the neuronal extracellular fluid. The development of the dialysis bag further developed the technique for sampling. The quantification, characterization, and sampling of neuropeptides and neurotransmitters in awake-freely-moving laboratory animals are the most widely used applications of brain microdialysis.

The microdialysis sampling technique is based on the law of diffusion. The law of diffusion explains the passive movement of molecules across the concentration gradient developed between the membrane and the interstitial fluid. The microdialysis procedure is an interchange between the dialysis membrane with “the probe,” the perfused liquid, the target tissue or organ, and the interstitial or extracellular fluid. An equilibrating fluid is perfused in the membrane tissue fluid to outside the membrane (Darvesh et al., 2011). The molecules of interest such as electrolytes, neuropeptides, amino acids, hormones, neurotransmitters, or neuromodulators, etc. are present in the dialysate outflow after equilibration.

Surgical microscope

Surgical tools and supplies

Preoperative Set-Up and Anesthesia Induction

Prior to the surgical procedure, examine the animals physically. Monitor the animals for nutritional status, fur quality (thinning, dirty), and behavior (limbs and trunk movement, abnormal gait, rigid walking, and a flat abdomen). Also, inspect the natural orifices for discharge from the nose, increased salivation, and filths around the anus and genitals, and observe the condition of the eyes. Monitor the breathing pattern of the subjects because non-manifesting subclinical pulmonary diseases may lead to severe respiratory failure following general anesthesia with subsequent death of the animal.

Anesthetize the animals with halothane (1.5-2% in a 50:50 O2/NzO mixture). After anesthesia induction, asses the depth of the anesthesia using the toe pinch test. Observe and assess the subjects for physiological parameters throughout the experimental procedures to ensure that the anesthesia is effective.

Brain Microdialysis Procedure

1. Place the subject in the stereotaxic frame and set the incisor bar at 3.3 mm.
2. Implant a semi-permeable membrane-containing probe or guide cannula in the ventral hippocampus.
3. Allow the animals to recover for 24 hours.
4. Pump the perfusion fluid into the probe via a perfusion pump slowly at an optimal rate (generally 1.8 – 2.2 μl/min) and collect the dialysate with the help of the collection device after equilibration.
5. Quantify and characterize the analyte as per experimental needs.

Post-Operative Care and Pain Management

Recover the animals on a flat paper bedding (sterile paper towels, etc.). For a speedy recovery, keep the animals warm. Place the recovery cage half-on a heating pad so that animals can choose their preferred temperature as they recover from the anesthesia. The animals which underwent surgery must have regained the ability to move in the cage freely. Monitor the animals post-operatively for unexpected signs of illness and pain. To regain weight quickly, provide the animals with proper analgesia and food.

Keep the animals under observation for five to seven days after surgery. Animals should be bright, alert, and active after the recovery. The animals should generally be interacting with the cage mates, eating and drinking. Depression, anorexia, or sluggishness indicate abnormal behavior. Food/fluid intake is essential to recovery. Provide animals with easier access to food and water. Inflammation, redness, swelling, discharge (purulent or serious), pain, anxiety or the opening of the incision (dehiscence) are the signs of inflammation. Treat the symptoms with proper ointment.

Applications

Comparison of the effects of intra-cerebrally administered MPP + (1-methyl-4-phenylpyridinium) in mouse and rat: microdialysis of dopamine and metabolites (Rollema et al., 1989)

Intracerebral microdialysis was used to measure the basal output of dihydroxyphenylacetic acid (DOPAC), Dopamine (DA), homovanillic acid (HVA) and 5-hydroxyindole-acetic acid (5-HIAA) from rat and mouse striatum in vivo. DOPAC/HVA ratios in the output dialysates from the mouse and rat striatum were 1:2 respectively. It was observed that the extracellular dopamine levels were 3 times lower than the level in the tissue concentrations. Similar effects of the intra-cerebrally administered dopaminergic neurotoxin l-methyl-4-phenylpyridinium (MPP +) were seen in both the subjects. The metabolites output accompanied the immediate and massive release of dopamine. At 5-12 h after MPP + administration the basal dopamine release was not detectable, and the subsequent MPP ~ perfusion did not potentiate the dopamine release. The brain microdialysis enabled the researchers to compare and analyze the effects of the test molecules in rodents.

Assessment of changes in 5-HT release in the ventral hippocampus in rats (Wright, Upton, & Marsden., 1992)

The study was conducted to combine in vivo microdialysis with behavior on the elevated X-maze in Sprague-Dawley rats to determine the levels of 5-HT release in the ventral hippocampus. Brain microdialysis was used to sample the 5-HT and 5-HIAA. The subjects were exposed to elevated X-maze for 20 minutes. An increase in the extracellular 5-HT in the ventral hippocampus was observed. However, there was no change in the level of extracellular 5-HIAA. The release of 5-HT in the extracellular was increased when the rats were exposed to either the closed or the open arms of the elevated X-maze, however, the 5- HT levels were not significantly increased when the subjects were restricted to the open arms as compared to the closed arms restriction. Diazepam (2.5 mg kg- 1 IP) treatment significantly reduced the level of extracellular 5-HT in the ventral hippocampus and induced anxiety over 5 min and 20 min X-maze exposure. The results suggested that the 5-HTIA receptor partial agonist ipsapirone (1 mgkg -1 IP) inhibits the release of extracellular 5-HT in the ventral hippocampus but does not has any effect on the animal’s behavior. The novel anxiolytic F2692 (10mgkg -1 IP) was found as antagonistic to the increase in extracellular 5-HT in the ventral hippocampus and induced anxiety over the 5 min but not in the 20 min period on the X-maze. The brain microdialysis enabled the measurement and characterization of the anxiolytic compounds on extracellular 5- HT levels and helped to delineate their behavioral profile on the X-maze.

Evaluation of the basal levels of neurotransmitters in the brain extracellular fluid (Boschi, Launay, Rips, & Scherrmann, 1995)

Brain microdialysis has been the feasible technique to sample the neurotransmitters from small brain areas of OF1 (iffa Credo mice) mice. In the study, dorsal hippocampus and nucleus accumbens were the brain regions of interest. Using the brain microdialysis, biogenic amine metabolites were quantified in the dialysate samples and were measured then by high-performance liquid chromatography (HPLC) accompanying the electrochemical detection (ED). DOPAC (3,4-Dihydroxyphenylacetic acid), MHPG (3-Methoxy-4-Hydroxyphenylglycol), and HVA (homovanillic acid) in the dorsal hippocampus were obtained soon after the probe was inserted, whereas 5-HIAA (5-Hydroxytryptamine indole acetic acid) concentration declined gradually. After 80 minutes the levels of DOPAC, HVA, and 5HIAA were stabilized in the nucleus accumbens. The compounds collected from the nucleus accumbens could be used for the assessment of drug-induced interactions. The brain microdialysis allowed the measurement of the basal levels of neurotransmitters and could allow the correlation of biochemical changes and pharmacological effects. The method can also facilitate further pharmacokinetic and biochemical research in rodents.

Assessment of the effects of aripiprazole on dopaminergic and serotonergic systems in rodents (Bortolozzi et al., 2007)

In the study, the in vivo effects of aripiprazole on serotonergic and dopaminergic systems were evaluated using the brain microdialysis. The research was conducted using male albino Wistar rats and C57BL/6 mice. It was observed that the 5-HT (5-hydroxytryptamine output was reduced in the medial prefrontal cortex (mPFC) of the hippocampus and the dorsal raphe nucleus in the rat following aripiprazole systemic administration. Also, the extracellular levels of 5-HT were reduced in the mPFC of wild-type (WT) but not in the 5-HT1A (−/−) knockout (KO) mice after the aripiprazole administration. The results suggested that aripiprazole has a reversal effect on extracellular 5-HT output potentiated by the 5-HT2A/2C receptor agonist DOI local application in mPFC. However, dopamine output in mPFC of WT was increased because of the aripiprazole. Whereas, the dopamine level was not increased in the 5-HT1A KO mice. In contrast to these, the haloperidol increases the activation of dopamine neurons in the ventral tegmental region of the brain. The dopaminergic activity was moderately reduced following the aripiprazole administration. The study indicates that the aripiprazole regulates the in vivo 5-HT and DA release in the medial prefrontal cortex by activating the 5-HT1A receptors. It was also concluded that the aripiprazole serves as a partial agonist at dopamine D2 autoreceptors, an action that is contrasting to the effects of haloperidol. The brain microdialysis technique is a feasible and reliable method to delineate the in vivo effects of atypical antipsychotic drugs.

Precautions

Do not exceed pre-anesthetic fasting beyond 2 hours because of the high metabolic rate of the rodents as extended food deprivation can disturb the balance leading to the metabolic acidosis, and hypoglycemia. Handle the animals gently to avoid stress which may lead to cardiac arrest following tachyarrhythmia during the general anesthesia. Also, the strain and breed of the laboratory animal must be selected as per the experimental requirements. During the surgical procedures, take care not to damage the surrounding tissues and muscles. Use clean home cages for animals to avoid contaminating the surgery area.

Summary

• In vivo brain microdialysis is a quantification and sampling technique used to measure the levels of the neurotransmitters in the brain extracellular fluid.
• Brain microdialysis has been widely used to evaluate the physiological and pharmacological effects of potential drugs in the extra-neuronal fluid.
• Rodent brain in vivo microdialysis has also been employed to assess the region-specific neurochemical changes induced by psychotic drugs and pharmacological compounds in the freely moving and awake rodents.
• The technique has enabled the researchers to measure the changes in concentrations of neurotransmitters and their respective metabolites following the drug administration.
• In vivo microdialysis helps to characterize the neuropharmacological agents such as the drugs of abuse, antidepressants, and antipsychotic agents.
• In vivo brain microdialysis has been considered as the backbone of neurological disorders treatment.

References

1. Bortolozzi, A., Díaz-Mataix, L., Toth, M., Celada, P., & Artigas, F. (2007). In vivo actions of aripiprazole on serotonergic and dopaminergic systems in rodent brain. Psychopharmacology (Berl), 191(3), 745-58.
2. Boschi, G., Launay, N., Rips, R., & Scherrmann, J. M. (1995). Brain microdialysis in the mouse. J Pharmacol Toxicol Methods, 33(1), 29-33.
3. Darvesh, A. S., Carroll, R. T., Geldenhuys, W. J., Gudelsky, G. A., Klein, J., Meshul, C. K., & Schyf, C. J. (2011). In vivo brain microdialysis: advances in neuropsychopharmacology and drug discovery. Expert Opin Drug Discov. 2011, 6(2), 109-127.
4. Rollema, H., Alexander, G. M., Grothusen, J. R., Matos, F. F., & Castagnoli, N. (1989). Comparison of the effects of intracerebrally administered MPP+ (1-methyl-4-phenylpyridinium) in three species: microdialysis of dopamine and metabolites in mouse, rat and monkey striatum. Neurosci Lett., 106(3), 275-81.
5. Vladimir, I. C., Alexis, C. T., Agustin, Z., & Shippenberg, T. S. (2009). Overview of Brain Microdialysis. Curr Protoc Neurosci.
6. Wright, I. K., Upton, N., & Marsden., C. A. (1992). Effect of established and putative anxiolytics on extracellular 5-HT and 5-HIAA in the ventral hippocampus of rats during behaviour on the elevated X-maze. Psychopharmacology (Berl)., 109(3), 338-46.

Specifications

Introduction

Apparatus

Pre-operative Set-up and Anesthesia Induction

Stereotaxic Surgery Protocol

1. Hold the needle at its distal third with the help of a needle holder, and pierce one edge of the wound.
2. Push the distal portion of the needle through while maintaining its curvature.
3. Release the needle and grasp once again with the needle holder between the wound edges. Repeat the procedure for the opposite edge of the wound.
4. Position the needle holder between the two ends of the thread and wrap it once by the needle strand and then lift the short tail strand.
5. Drag the long strand towards the wound and establish a simple single overhand throw. Repeat the procedure and make a second overhand throw in reverse order. Knot both the threads.

Post Operative Care and Handling

Applications

Precautions

• Take note of the entire life experience of the animal to consider other factors that could affect the surgery such as transportation, individual history, and housing and feeding.
• Before starting the surgical procedure, consider the conditions such as operant conditioning, food/drink restriction, irradiation, environmental distress, disease conditions, and exposure to predator scents.
• Identify the pain and distress symptoms pre-and post-operatively to ensure the animal's health and safety.
• If found treat the pain with the proper administration of analgesic.
• In the case of survival surgery, it is essential to observe the subjects post-operatively closely. Follow-up observation must include monitoring of animal behavior and evaluation of pain and distress.
• Do not perform stereotaxic surgeries in paralyzed animals without anesthesia. Always combine neuro-muscular paralytics with anesthetic agents in stereotaxic neurosurgery.
• Before starting the surgical procedure, ensure the adequate depth of the anesthesia using a toe pinch test or other anesthesia assessment tests.
• Carry out the whole surgical procedure under aseptic conditions.

Summary

• Stereotactic neurosurgery is the most widely used surgical technique for precisely locating the brain regions relative to an external frame of reference.
• Stereotactic neurosurgery has provided precise localization of targets of interest to assess the anatomic, metabolic, and functional aspects of the brain regions.
• Stereotaxic surgery is applied for the treatment of motor disorders, interstitial radiotherapies, assessment of tumoral pathologies, hydrocephaly treatment, neuronal stem cell transplantation, and evaluation of cerebral arteriovenous malformations.
• Neurotoxic surgery is used to assess the in vitro efficacy of chemotherapeutic loaded microsphere in the rat brain.
• The stereotaxic transplantation of neuronal stem cells into the spinal cords of mice, affected by MHV, improved the re-myelination and the pathological condition.
• Stereotaxic neurosurgery should not be performed in paralyzed animals without anesthesia.
• Make sure that the entire surgical procedure is performed under aseptic conditions.

References

Rat Kit

Small Animal Kit

Introduction

Pre-operative Set-up and Euthanasia

Cervical Dislocation

1. Hold the head with one hand and place the thumb and the index finger on either side of the neck at the base of the skull.
2. Pull the base of the tail with the other hand quickly.
3. Just press the thumb tip firmly into the neck behind the skull immediately.

Carbon Dioxide Inhalation

1. Place the cage containing the animal in a chamber with a flow of 100% CO2 gas initiated.
2. Monitor the flow of CO2 gas using a gas flow meter to reach 20-30% of chamber volume per minute (a higher flow rate may result in animal stress due to pain before the loss of consciousness, whereas a lower rate would be too slow).

Overdose of Inhalant Anesthetic Agents

Dissection Protocols

1. Place the animal with the ventral side up on a clean dissection board and pin it. Avoid holding the animal in one hand to keep your hands clean and to cut the organs accurately.
2. Using soaked cotton, clean the ventrum to avoid the introduction of animal hair into the body that may infect the organs.
3. Hold the skin with dissecting forceps and raise it above the abdomen.
4. Make an incision through the body wall muscles by putting the scissor just anterior to the genital opening and continue cutting on one side of the midline, ventral to the thoracic cavity.

5. Keep the fur away from the body by pulling the skin from both sides of the thoracic cavity.
6. Isolate the larger parts of the lungs and store them in the cryogenic tubes quickly.
7. Clean the forceps and scissors using a solution containing bleach, water, and ethanol.
8. Find the spleen by moving the stomach to the left (the spleen has a triangular shape).
9. Pull the spleen out by holding it with the forceps and get rid of the white tissue with the help of sharp pointy scissors.
10. Clean the forceps and scissors with the cleaning solution.
11. Displace the bowels to find the kidneys beneath the intestines.
12. Separate the kidneys and place them in two cryogenic tubes for storage (if the kidneys are too big, cut them into pieces).
13. Clean the dissecting instruments with the cleaning solution.
14. Dissect the diaphragm and keep it in a cryopreservative tube.
15. Place the cryogenic tubes containing the dissected organs in a liquid nitrogen tank (Do not open the tank again and again rather put the tubes at one time to prevent nitrogen evaporation from the tank).
16. Wash the forceps and scissors with the solution containing bleach, water, and ethanol).
17. Identify the sex of the animal by observing the genital organs and counting the embryos or measuring the size of the testes.
18. Tag the leg of the animal. Using fine point forceps, perforate the skin and introduce the string.
19. Place the animal into an ethanol-filled jar.
20. Clean and wash the dissection board and the operative area.

Dissection of Mouse EDL and Soleus Muscles (Bröllochs, 2017)

1. Fix the leg of the animal with pins in a flexed position to the dissecting dish and cover the leg with Ringer solution.
2. Remove the skin of the leg and get rid of the excessive fascia surrounding the muscles.
3. With the help of the forceps, hold the Achilles tendon. Use the bodkin side of the scissors to rub the gap open between the Achilles tendon and the other tendons. Cut the Achilles tendon as close as possible to the foot when the gap is visible.
4. As the gap starts to open, support that gap to open up fully.
5. The Soleus muscle will be visible as the gap opens up to the knee. The Soleus muscle is dark pink, identify it by its color. Carefully cut the tendon near the knee.
6. Hold the Soleus muscle from its upper tendon gently with the forceps and pull it down slowly and carefully while (if needed) removing the surrounding fascia that is still holding it. Cut the lower tendon after freeing the whole muscle from the fascia.
7. Isolate the Soleus muscle from the leg and remove the leftover fascia. For more comfortable handling, one should move the muscle to a new dissecting dish, fix it with pins, infuse it with Ringer solution, and store it at 4 °C.
8. To dissect the EDL muscle, place and pin the leg into a stretched position first and remove the fascia.
9. Rub the lower leg’s tendons. The 4 EDL tendon ends are next to the V-looking Tibialis tendon. Carefully lift the tendons to observe the EDLs. The toes of the paw stretch while lifting. Make an incision at the ends of the tendon as close as possible to the foot.
10. A pocket will open up as the ends are cut. Remove the surrounding fascia. Isolate the EDL muscles from the pocket to visualize the upper tendon. Dissect the upper tendon then.
11. Isolate the EDL muscle from the leg and remove all the fascia and the fat.
12. Place both the dissected muscles in a clean dissecting dish. Lightly stretch the muscles and pin them. Cover the dissected muscles with a 3.7% PFA.
13. Store the dish at 4°C overnight after covering it with the parafilm.

Gross Postmortem Location and Examination of Heart, Lungs, Liver, Kidneys, and Spleen Method (Parkinson et al., 2011)

1. Examine the animal for skin and coat abnormalities, emaciation, or dehydration.
2. Keep a log of artificial manipulations, implants, or surgical scarring, if any.
3. With the help of the dissecting microscope examine all the external orifices (ears, eyes, nose, anus, genital openings, and oral cavity) for secretion or bleeding.
4. Place the euthanized animal in dorsal recumbency on a clean dissection board.
5. With the help of dissection scissors, make an incision through the skin from the length of the ventrum extending from the anus to the chin, reflecting the skin and incising the abdominal wall, opening the abdominal viscera, salivary and preputial/clitoral glands, and cervical and axillary lymph nodes. Examine the thoracic viscera by making 2 incisions laterally up on each side of the ribcage, then make a cut across, at the top of the sternum, to open a wide space enough to observe all the lobes of the lung thoroughly.
6. Observe the musculoskeletal structure.
7. Assess the condition of all the organs for abnormalities. Locate and identify the heart and the lungs in the thoracic cavity. Observe the liver, kidneys, and spleen in the abdominal cavity. Check the organs for any color changes, size differences, and absence or dislocation. Record the consistency of surfaces, any additional tissue (e.g., masses), fluid pockets, or the presence of fluid in the abdominal/thoracic cavities.
8. Examine the gastrointestinal tract for contents, lack of contents, thickened walls, masses, blood clots, and hemorrhage. Using a sharp blade, cut the kidneys (left-longitudinal section, right-cross section, on the midline, but off-center) to assess parenchyma for any abnormality. Check the presence of any large lymph nodes on the mesentery.
9. Observe the urogenital system and check for any blockage, fluid pockets, hemorrhage, or other abnormalities.

Postmortem Collection of Heart, Liver, Kidneys, and Spleen for Histopathology

1. Collect a container filled with a suitable amount of 10% neutral buffered formalin (NBF), and make sure that the container is appropriately sized and labeled.
2. Adjust the amount of 10% NBF to obtain a 20:1 ratio of fixative to tissue.
3. Lay the animal’s carcass in dorsal recumbency on a clean dissection board and expose the tissues of interest.
4. Isolate the tissues from the animals’ carcass with the help of dissecting forceps and scissors.
5. Scrap the tissues to remove fat and unnecessary connective tissue carefully. The blood should be clean; use normal saline to rinse.

6. Place the dissected tissues in the container containing 10% NBF.

Postmortem Collection and Perfusion of Lung Tissue

1. Fill a container with 10% NBF and adjust the volume of 10% NBF to obtain a 20:1 ratio of fixative to tissue.
2. Lay the animal to be dissected in dorsal recumbency on a clean dissection board.
3. Expose and display the trachea, heart, and lungs.
4. Remove the skin and muscle overlying the ventral thoracic and cervical regions using the dissecting forceps and scissors.
5. Trim the ribcage to expose the heart and lungs near the clavicle to open space, ample enough to observe all the lobes of the lung thoroughly.
6. Remove the neck muscles surrounding the sternum and ribs and ranging to the jaw, including those overlying the trachea.
7. Make two cuts by inserting the scissors under the anterior edge of the rib cage, one cut on either side, to remove the section of bone overlying the trachea.
8. With the forceps hold the trachea near the jaw and cut the trachea entirely by placing the scissors above the forceps
9. Pull out the trachea with the help of the forceps and cut the ventral tissue connections with scissors until all the thoracic tissues are removed from the body.
10. Place the lungs flat on the shelf.
11. Stitch the trachea with suturing material loosely.
12. Fill a syringe with a fixative and attach a needle, small enough to enter the trachea (For mice, a 1ml or 3ml syringe with a 26 gauge needle works well. For rats, a 5ml syringe with an 18 gauge syringe is accurate).
13. Hold the trachea with the forceps and insert the needle into the trachea. Fill the lungs with fixative slowly. Keep filling until the lungs are fully inflated. Do not over-or underinflate. The amount of fixative needed depends on the age, strain, and health of the animal.
14. Fluid seeping and foaming from the tissues indicate over-inflation.
15. Flat lungs indicate under-inflation.
16. Withdraw the needle from the trachea.
17. Tighten the stitches on the trachea to prevent backflow of fixative out of the lungs.
18. Keep the inflated lungs into the fixative with a 20:1 fixative to tissue ratio.

Postmortem Collection of Brain

1. Bring an appropriately sized, labeled container(s) and fill in the required amount of 10% NBF and adjust the amount of 10% NBF to obtain a 20:1 ratio of fixative to tissue.
2. Place the animal carcass in ventral recumbency on a clean dissection board.
3. Remove the skin and the muscles surrounding the calvaria with the help of dissecting scissors.
4. Dislocate and remove the head from the animal’s body.
5. Insert the bottom blade in the foramen magnum (the opening where the skull opens into the spinal canal) with the help of small scissors, keep the scissor tips pointed upwards, and begin cutting through the midline of the calvaria.
6. Reflect both halves of the calvaria and expose the brain.
7. Place the exposed brain encase in the skull into a fixative.
8. Invert the skull gently so that the tissues fell from the skull because of the gravity.
9. Move the forceps under the brain starting at the olfactory lobes and along the outer edge of the brain, moving under the cerebrum and towards the cerebellum carefully. Gently pinch the connective tissue or nerves, with the help of the forceps, inhibiting the brain from falling from the skull.
10. Place and keep the brain into the fixative using an approximate 20:1 ratio of tissue to fixative.

Postmortem Collection of Respiratory Aspirate

1. Place the animal to be dissected in dorsal recumbency on a clean dissection board.
2. For bronchial aspirate collection in rats, get access to the respiratory tract through the trachea. For nasal aspirate in rats, reach the respiratory tract either through the trachea or the nasopharyngeal meatus. For bronchial or nasal aspirates in mice, approach the respiratory tract from the nasopharyngeal meatus.
3. Tracheal Access (recommended for rats):

• Expose the subcutaneous tissues by moving the skin away from the cervical area.
• Separate the salivary glands and cervical musculature to expose and visualize the trachea.
• With the help of sterile dissecting instruments, cut the trachea to reach its lumen.

4. Nasopharyngeal Meatus Access:

• Cut the temporomandibular (jaw) joint and move the mandible away from the maxilla to expose the nasopharyngeal meatus.

• Draw approximately 1ml of sampling fluid into a sterile pipette.

5. Bronchial Aspirate:

• Inject the sampling fluid slowly into the bronchi and the lung by caudally inserting the pipette into the trachea. Pipette out the sampling fluid from the bronchi and the lungs and remove the pipette from the trachea.

5. Nasal Aspirate:

• Cranially direct and insert the pipette into the nasopharyngeal meatus (mice) or tracheal lumen (rats), and pour the sampling fluid into the nasal cavity.
• Access the nasal cavity by contacting the nasal palate with the pipette tip, or by observation of fluid forced into the cavity, seen as menisci forming at the nasal orifice (nares) or as fluid visible through the translucent oral palate.
• Draw the sampling fluid from the nasal cavity into the pipette and remove the pipette from the meatus.
• Transfer the sample to an appropriate media or container for testing.

Dissection of Mouse Parotid Glands

1. Euthanize the mice using the cervical dislocation method.
2. Cut and open the skin of the chest and head/neck region from the epigastrium toward the limbs.
3. Open the thorax by making a trapezoid cut.
4. Make a small incision in the apex of the left ventricle, insert the perfusion cannula into the incision and gently push into the aortic arch.
5. Perfuse [4% paraformaldehyde (PFA) in PBS with RNase-free water at pH 7.4] for 2–3 min.
6. Lift the parotid glands with a pair of forceps to locate the borders of the organ and cut out, including a part of the main excretory duct.
7. Separate the parotid glands from the surrounding tissues.

Mouse Dorsal Root Ganglia (DRG) Isolation Protocol

Spinal Column Isolation

1. After submerging the fur with 70 % ethanol, make a small incision in the dorsal skin at the level of the hips, and remove the pelt from the head to hind limbs.
2. Remove the head by cutting at the base of the skull (C1–2 level) and cut the arms beneath the shoulders to aid the removal of the skin.
3. Make an incision through the abdominal wall muscles and continue laterally to the spinal column in both directions.
4. Before the viscera detachment, incise the ribs closer to the spinal column from both sides.
5. Cut the femurs, and remove the spinal column by cutting transversely at the level of the femurs.

Spinal Cord Exposure

1. Cut the muscles, fat, and other soft tissues from the spinal column with the help of curved scissors. The T13 level DRG pair present at the caudal ribs is used as a landmark.
2. Remove the spinal nerves projected from the column.
3. Once cleared of soft tissues, cut the column into three pieces, with one cut at the level of the last rib to orientate the dissection.
4. Place the column segments dorsally with the side facing up.
5. Use thick forceps to secure the spinal column dorsal side up, before cutting it into two halves along the midline.
6. Pin the Hemi-segments of the spinal column in Petri dishes with the medial side up, using two insect pins through intervertebral discs, before rinsing with ice-cold PBS.

DRG Extraction and Cleaning

1. Peel the spinal cord from the pinned column in a rostral to caudal direction.
2. Identify the transparent meningioma sheets of tissue covering the DRG, and carefully remove it, making the DRG easier to see.
3. Dissect out the individual ganglia by grasping and lifting with forceps, and find the distally projecting axon bundles on the lateral side of the DRG. Do not damage the DRG with the forceps.
4. Pinout the DRG via their axons, and remove any residual meninges, before cutting the axons close to the DRG.

Applications

Precautions

• Keep a note of the preservative solution.
• Rinse the preserved animals under running water immediately upon removal from the preservative solution.
• Work in a well-ventilated area.
• It is recommended that contact lenses should not be worn while dissecting animals that are in preservative solution. As the fumes from the solution can penetrate between the eye and contact lens are irritating the eyes. Wear prescription glasses instead, with safety glasses over them.
• Be aware of issues such as allergies and chemical sensitivities from handling freshly euthanized or recently defrosted or preserved animals.
• Be aware of possible microbial aerosols and unpleasant odors released from freshly euthanized or recently defrosted animals if the stomach/intestines are accidentally cut.
• If using frozen animals, defrost overnight in a refrigerator beforehand and dissect within 24 hours.
• Observe good hygiene practices throughout the procedures by keeping the hands away from the mouth, nose, eyes, and face during and after the dissection and wash the hands immediately after handling the dissection material.

Summary

• In biological research, dissection is the process of cutting and disassembling the body parts of the laboratory animals to study their anatomical structures.
• Rodents are the most preferred species to be used as animal models for biomedical research due to their anatomical, physiological, and genetic similarities to humans.
• The rodents are euthanized before the dissection procedures. Different methods of euthanasia are applied, enabling a rapid death with the reduced pain for the animal and the safety of the field workers should be selected.
• The dissection procedure helps the researchers to perform histological analysis as well as anatomical investigations on rodents.

References

1. Bröllochs, A. (2017, August 11). Dissection of mouse EDL and Soleus muscles. Retrieved from Protocols.io: https://www.protocols.io/view/dissection-of-mouse-edl-and-soleus-muscles-jcrciv6?step=13
2. Parkinson, C. M., O'Brien, A., Albers, T. M., Simon, M. A., Clifford, C. B., & Pritchett-Corning, K. R. (2011). Diagnostic Necropsy and Selected Tissue and Sample Collection in Rats and Mice. J Vis Exp, 54, 2966.
3. Sleigh, J. N., Weir, G. A., & Schiavo, G. (2016). A simple, step-by-step dissection protocol for the rapid isolation of mouse dorsal root ganglia. BMC Research Notes, 9(82).
4. Watermann, C., Valerius, K. P., Wagner, S., Wittekindt, C., Klussmann, J. P., Vogt, E. B., & Karnati, S. (2016). Step-by-step protocol to perfuse and dissect the mouse parotid gland and isolation of high-quality RNA from murine and human parotid tissue. BioTechniques, 60, 200-203.

Introduction

Preoperative Set-up and Anesthesia Induction

Rodent Ophthalmic Surgery Protocols

• Measure the thickness of the Inner Plexiform (IPL), Inner Nuclear (INL), Outer Plexiform (OPL), and Outer Nuclear (ONL) layers, and the total thickness by taking the mean value of five different measurements at three points per section.

• Anesthetize the animal by giving an intraperitoneal injection of a mixture of ketamine hydrochloride (75 mg/ml) and medetomidine hydrochloride (1 mg/kg) in saline.
• Assess the depth of the anesthesia by toe pinching.
• Disinfect the eyelids by applying 70% ethanol using a cotton tip.
• Place the animal on a flat, dry, and smooth surface.
• Sterilize the forceps with the help of a curved, serrated tip (preferred tip size: 0.5 x 0.4 mm).
• Press the corner of the eye (canthus) gently with the help of forceps until the eyeball is displaced out from the socket and the optic nerve is accessible.
• Approach the eye from behind with the forceps. Push and grasp the optic nerve firmly, with the beginning of the curve and not the very tip of the forceps. The grip will help to lift the globe from the socket and to clamp the optic nerve completely.
• Move the forceps with the least resistance in a circular motion and the mouse swings along the surface according to the direction of the hand movement.
• Perform this action with gradually increasing speed until the optic nerve is constricted in two (usually between 7 to 15 circular movements, approximately a half to one full turn per second). The detached eyeball is removed.

• Anesthetize the animals by intraperitoneal administration of ketamine hydrochloride 100 mg/kg and xylazine hydrochloride 10 mg/kg.
• Carefully make an incision close to the temporal lid margin to expose the ILG.
• Excise the ILG from the orbit using ophthalmic forceps with the aid of a stereoscopic microscope.
• Make an additional incision anterior to the ear, and then expose and excise the ELG.
• Suture the surgical incisions using 6-0 ophthalmic nylon thread and apply the ointment containing Ofloxacin.

• Induce general anesthesia in the animal by IP injections of ketamine (86.98 mg/kg) and xylazine (13.04 mg/kg).
• Apply Puralube ointment on the eye not undergoing surgery to prevent dryness.

• After achieving full anesthesia, obtain adequate mydriasis by administering 1% tropicamide and 2.5% phenylephrine hydrochloride eye drops.
• Place the head of the donor animal horizontally on a board placed on robust, movable support. With a strip of tape, fix the head of the animal across the neck to ensure the horizontal eye position throughout the entire procedure.
• With the help of a 2 mm diameter trephine, whose tip is dyed with methyl blue, sketch the central cornea graft site.
• Using a sharp blade, infiltrate the cornea and inject sodium hyaluronate into the anterior chamber to deepen it reducing the chance of damage to the donor endothelium and the underlying lens.
• Excise the donor graft with vannas scissors and place it into a dish containing Hanks’ balanced salt solution until use.
• After the donor graft has been removed, euthanize the donor animal via CO2 inhalation.

• Repeat the first two steps mentioned above for the recipient animal.
• With the help of a 1.5 mm diameter trephine, outline the recipient graft site.
• Using a sharp blade, infiltrate the cornea and inject sodium hyaluronate into the anterior chamber to deepen it to reduce the chance of damage to the underlying lens.
• Remove the outlined central corneal button from the recipient using vannas scissors and discard.

• Set the donor cornea over the graft bed in the recipient's cornea. Make sure that adequate sodium hyaluronate is present under the donor cornea to protect the donor endothelial cells from damage by direct contact with the lens.
• Using super-fine tipped micro-forceps, place the first bite of 11-0 nylon suture into the donor side, through the donor with 90% depth of full thickness to recipient’s side, then tie off.
• Once the cornea is anchored in place, perform mid-cardinal interrupted sutures such that the cornea has 8 to 10 total sutures and the donor cornea is securely lined up with and attached to the recipient corneal graft bed.

• Penetrate the anterior chamber by injecting HBSS or air bubble into the anterior chamber and assess the strength and integrity of the corneal graft for leakage with a cellulose sponge gently.

• Anesthetize the animals as mentioned above and remove the lid suture after 48 hours or prescribed time.
• Anesthetize the animals on day 7 postoperatively. Carefully remove the sutures while securing the cornea. Return the animal to its cage after suture removal and consciousness regain.

• Anesthetize the animals by intraperitoneal injection of xylazine (2 % solution) and ketamine (30 mg/kg).
• Excise the sternocleidomastoid muscle via an anterior incision on the neck and then expose the external jugular vein and two main branches of the common carotid artery.
• Excise the anterior and posterior bellies of the digastric muscle, omohyoid muscle, and greater horn of the hyoid to get better exposure of the external carotid artery and its branches.
• Ligate and cut the internal carotid artery and all branches of the external carotid artery except the superficial temporal artery.
• Prepare the flap containing orbital tissues, external ear and a part of the facial skin.
• Cut the optic nerve at its most possible proximal end to the eyeball and include it in the flap.
• Raise the composite tissue flap based on the common carotid artery and external jugular vein.
• Expose and anastomose the external jugular vein and the common carotid artery via an anterior incision on the neck.
• Cut the great auricular nerve with fine dissection scissors, and use the proximal stump of the nerve for coaptation.
• Anastomose the vein of the flap to the external jugular vein of the recipient animal in end-to-end fashion using No. 10-0 interrupted monofilament nylon sutures.
• Anastomose the artery of the flap to the common carotid artery of the recipient animal in end-to-side fashion using the same suture material.
• Obtain the perfusion of the graft, and then coapt the optic nerve of the flap to the distal stump of the great auricular nerve of the recipient animal in end-to-end fashion with four epineural sutures.
• Finally, perform the complete tarsorrhaphy on the eye of the allograft using 8/0 monofilament nylon sutures.

Mode of Operation

Post-operative Care and Pain Management

Applications

Precautions

Precautions

Summary

• Ophthalmic research involves the improvement of treatments of ocular diseases and conditions.
• Rodent models have significant advantages over large animal models such as low maintenance and cost, shorter breeding cycles, and faster regeneration.
• Athymic, transgenic, and knock-out rodent model availability makes them ideal for various research requirements.
• The precorneal tear film glands in rodents may prolapse in pathological conditions.
• The orbital vascular plexus, present in rodents and lagomorphs, differs substantially between species and the understanding of its anatomy is essential in orbital surgery and enucleation.
• Rodent strain, breed, and allergic history among other factors may influence the ophthalmic investigations.
• Anesthesia induction can be done using inhalants or injectable agents. The depth of anesthesia should be verified before beginning surgical procedures.
• During surgical procedures, care must be taken not to damage surrounding tissues or muscles.
• A recovery area should be set-up and fluids should be replaced by subcutaneous or intraperitoneal injection of warm sterile saline.
• Infections may influence the results of the investigation, hence, euthanizing the subject is recommended should it occur.
• Appropriate pain management techniques should be followed.

References

Rat Kit

Introduction

Pre-operative Set-up and Anesthesia Induction

• Anesthetize the surgical subjects with pentobarbital 70 mg/kg in a solution of 15 mg/ml (Use a face mask or an anesthetic chamber to anesthetize the animals). Do not administer analgesics pre-operatively.
• Shave the chest with a hair clipper or a razor.
• Place the animal on the surgical stage in surgical position for the subsequent intubation, wire 3-0 silk behind the front incisors, pull taut and fix with tape.
• Intubate the animal with a 20-ga catheter, attached to the extender.
• Connect the intubation tube to the ventilator and start ventilating the animal.
• Perform the surgical procedure at room temperature (except for the ischemia-reperfusion).
• Prepare the surgical field.

Cardiovascular Surgery Protocols

• Open the chest with a lateral incision at the 4th intercostal space on the left side of the sternum and retract the chest.
• Remove the pericardial sac and pass an 8-0 silk suture in the myocardium at the anterior surface of the heart for one second and then remove it.
• Close the chest wall by approximating the third and fourth ribs with one or two interrupted sutures, return the muscles to their original position, and the skin closed with retract4-0 prolene suture.
• Gently disconnect the animal from the ventilator.

• Locally anesthetize the animal by injecting 0.1 ml of 0.2% lidocaine subcutaneously at the surgical site.
• Transversely incise the skin 5-mm with scissors 1–2 mm higher than the level of the ‘‘armpit’’ (with paw extended at 90^o) 2 mm away from the left sternal border.
• Separate the two layers of thoracic muscles.
• Separate the intercostal muscles in the 2d intercostal space.
• Interpolate the chest retractor to facilitate the view.
• Pull the thymus and surrounding fat behind the left arm of the retractor. Gently pull the pericardial sac and attach it to both arms of the retractor.

1. Bluntly dissect the ascending portion of the aorta on its lateral side from the pulmonary trunk with Foerster curved forceps.
2. Place Foerster curved forceps from the medial side under the ascending aorta, hold the 7-0 silk suture on the opposite side and pass it underneath the aorta.
3. Tie a loose double knot to create a loop 7–10mm in diameter.
4. Position a needle of proper size into the circuit.
5. Tie the loop around the aorta and needle, and bind with the second knot; immediately remove the needle to provide a lumen with a stenotic aorta. Make another knot to secure the tie.
6. Remove the chest retractor and re-inflate the lungs.
7. Close the chest wound layer-by-layer.

1. Bluntly depart the thymus, and pericardial sac with slightly curved forceps, then separate the aortic arch from the surrounding tissues and vessels.
2. With the help of a blunted needle, create a way underneath the aortic arch.
3. With a ‘‘wire and snare’’ device, deliver a 7-0 silk suture underneath the aortic arch between the innominate and left carotid arteries.
4. Tie a loose double knot to create a loop 7–10mm in diameter.
5. Position a needle of proper size into the circuit.
6. Tie the circle around the aorta and needle, and secure with the second knot; remove the needle immediately to provide a lumen with a stenotic aorta. Make another knot to bind the tie.
7. Remove the chest retractor and re-inflate the lungs.

1. Approach the great vessels via the second intercostal space.
2. To supplement the animal with local anesthesia inject 0.1 ml of 0.2% lidocaine subcutaneously at the surgical site.
3. Make a transverse 5-mm incision of the skin with scissors 1–2 mm higher than the level of the ‘‘armpit’’ (with paw extended at 90^o) 2 mm away from the left sternal border.
4. Separate the two layers of thoracic muscle.
5. Isolate the intercostal muscles in the 2d intercostal space.
6. Interpolate the chest retractor to visualize the organs.
7. Pull the thymus and surrounding fat behind the left arm of the retractor. Gently draw the pericardial sac and attach it to both the arms of the retractor.
8. After moving the pericardium, observe the pulmonary trunk (partially covered by the left atrium).
9. With the help of Foerster curved forceps, bluntly dissect the pulmonary trunk from the aorta (on the left) and left atrium (on the right).
10. With a blunted needle, create a passage under the pulmonary trunk.
11. With the help of a ‘‘wire and snare’’ device, move the 7-0 silk suture underneath the pulmonary trunk.
12. Tie a loose double knot to create a loop 7–10mm in diameter.
13. Position an appropriately sized needle into the loop.
14. Tie the loop around the aorta and needle and secure with the second knot; immediately remove the needle to provide a lumen with a stenotic pulmonary artery. Make another knot to secure the tie.
15. As the heart rate significantly slows down, stimulate the mouse.
16. Remove the chest retractor and re-inflate the lungs.
17. Close the chest wound layer-by-layer.

1. For local anesthesia, inject 0.1 ml of 0.2% lidocaine subcutaneously at the surgical site.
2. Make an oblique 8-mm incision of the skin (parallel to ribs) 2 mm far from the left sternal border toward the left armpit (1–2 mm below it).
3. Separate the two layers of the thoracic muscle.
4. Separate the intercostal muscles in the fourth intercostal space.
5. Insert the chest retractor to facilitate the view.
6. Gently pull apart the pericardial sac and attach it to both arms of the retractor.
7. Locate the left anterior descending (LAD) coronary artery.
8. Pass the tapered needle with 7-0 silk suture underneath the LAD.
9. Tie the suture with one double and then with one single knot.
10. Remove the chest retractor and close the wound layer-by-layer.

1. Hold the artery and place the suture (same as step 1-8 above).
2. To create ischemia, tie the temporary suture around the LAD. Cover the incision with wet gauze to prevent drying.
3. Release the tie (do not remove the suture) at the end of ischemia and observe the reperfusion.
4. If the procedure is survival, leave the suture underneath the LAD and close the chest wound layer-by-layer.
5. Retie the suture to demarcate the area at risk by infusion of blue dye.

• Intubate the animals intratracheally by transesophageal illumination using a fiber-optic light.
• Connect the tubing to the ventilator.
• Expose the pectoral muscles by making an incision on the chest to view the ribs.
• For ischemia/reperfusion studies, place a 1 cm piece of PE tubing over the heart to tie the ligature to so that occlusion/reperfusion can be customized.
• For intramyocardial injections, use a syringe with a sterile 30gauge beveled needle.
• Complete the myocardial manipulations, close the rib cage, the pectoral muscles, and the skin sequentially.
• Apply 0.25% Bupivacaine in sterile saline to muscle layer before the closure of the skin.
• Give a subcutaneous injection of saline and place the mice in a warming chamber until they are sternally recumbent.
• Return the animals to the vivarium and housed under standard conditions until the time of tissue collection.
• Anesthetize the mice, arrest the heart in diastole with KCl or BDM, rinse with saline, and immerse in fixative.
• Subsequently, perform routine procedures for processing, embedding, sectioning, and histological staining.

• Administer the first dose of the analgesic (0.1 mg/kg) intraperitoneally at the completion of the surgery.
• Move the animal to another ventilator in the recovery area with 100% oxygen loosely connected to its inflow.
• Provide heat by either a 60-W lamp or a heating pad at a low setting.
• Once the mouse attempts to breathe spontaneously (generally after 45–60 min), disconnect the intubation tube from the ventilator.
• Keep the intubation tube in the trachea for another 10–15 min until the mouse resumes the regular breathing pattern.
• Provide supplementary oxygen (the mouse is placed next to the source of oxygen).
• Extubate the mouse and return it to a clean, warm cage.

Applications

Precautions

Strengths and Limitations of Rodent Models for Cardiac Surgery

Summary

• Cardiovascular surgery focuses on the heart and blood vessels. Cardiac surgery is mainly performed to treat the complexities of ischemic heart diseases, congenital heart diseases, valvular heart diseases, endocarditis, rheumatic heart diseases, and atherosclerosis.
• Mouse models have gained popularity because of their small size, rapid gestation age (21 days), relatively low husbandry costs, convenience in housing and handling, and fewer compound requirements for pharmacological studies.
• With the advancement of genetic modifications to the whole organism or the myocardium and an array of biological and synthetic materials, there is an excellent potential for any combination of these to mitigate the extent of the acute ischemic injury and impede the onset of heart failure pursuant to myocardial remodeling.
• Do not administer pre-surgical analgesia (buprenorphine) since narcotic analgesics are known to depress the respiratory center and may interfere with the survival after the open-chest surgery.
• The significant challenges that arise during cardiac surgery are the extremely thin and delicate walls of the pulmonary trunk and the incapacity of the right ventricle to undergo stress while the pulmonary artery is being manipulated. Be careful during the operative procedures.

References

• Hashmi, S., & Al-Salam, S. (2015). Galectin-1: a biomarker of surgical stress in a murine model of cardiac surgery. Int J Clin Exp Pathol, 8, 7157-7164.Tarnavski, O. (2009). Mouse surgical models in cardiovascular research. Methods Mol Biol, 573, 115-37.Tarnavski, O., McMullen, J. R., Schinke, M., Nie, Q., Kong, S., & Izumo, S. (2004). Mouse cardiac surgery: comprehensive techniques for the generation of mouse models of human diseases and their application for genomic studies. Physiol Genomics, 16, 349-360.Virag, J. A., & Lust, R. M. (2011). Coronary Artery Ligation and Intramyocardial Injection in a Murine Model of Infarction. J Vis Exp, 52, 2581.

Mouse Kit

Rat Kit

Introduction

Preoperative Set-Up and Anesthesia Induction

Microsurgery Protocols

Preparation of the Vessel

• Place a strip of silicon gloves behind the dissected vessel to avoid contamination of the anastomotic area.
• Gently place a vascular clamp on the proximal and the distal part of the artery.

• Cut the vessels by perpendicularly placing the scissors on the vessel axis. Irrigate the lumen with a solution.
• Remove the adventitia from a few millimeters of distance from the edges. If the distal end is not used for anastomosis, ligate the distal end of the vessel.
• Dilate the lumen of the dissected vessel with tweezers to avoid spasms and widen the diameter to a maximum of 1.5 times the original width.
• Use an isotonic solution to rinse the wounds. To prevent vasospasms wash the vessel lumen with the solution.

End-to-End Anastomosis

Continuous Suture

• The upper pole is defined as “12 o’clock” and the lower pole as “6 o’clock”.
• Place the first suture at the center of the posterior wall (12 o’clock).
• Place the second suture on the anterior wall (6 o’clock).
• Fix one end of each stitch temporarily in gentle tension to keep the vessel lumen flattened and expanded.
• Begin anastomosis from 12 o’clock from the posterior wall of the vessel towards 6 o’clock (To avoid the accidental stitching of the dorsal wall place the plastic cannula with the appropriate diameter into the lumen during the suture).
• Knot the first thread of the suture to one or both strands at 6 o’clock. Complete the continuous sutures by suturing the anterior wall from 6 o’clock to 12 o’clock, where the thread at 6 o’clock is tied to one of the threads at 12 o’clock.

Interrupted Suture

• Place the additional sutures between the main stitches on the anterior vessel wall.
• Then rotate the clamps by 180o to allow the suturing of the posterior wall.
• Place the posterior and anterior sutures. Rotate the vessel 90°s by traction of the two stay sutures, to present half of the vessel anteriorly.
• Place the third, fourth, and fifth sutures. After traction on the two stay sutures rotate the vessel 90°s back to its original position.
• In a further 90° rotation, present the opposite unstitched half of the vessel anteriorly.
• After completing the suturing process, reposition the vessel to its original position.

End-to-Side Anastomosis

Continuous Suture

• Knot the corner at the heel of the anastomosis leaving the short end longer so that it can be used for tying later.
• Stitch the apex of anastomosis at a 180° angle from the first corner knot by taking the second thread.
• Make a continuous suture of the posterior wall by using the thread from the heel stitch.
• Gently tighten the suture and tie the knot with the shorter end of the “apex stitch.”
• Continue suturing the anterior wall by using the longer end of the “apex stitch.”
• Tie the final knot with the shorter end of the “heel stitch” by tightening the threads.

Interrupted Suture

• Knot the corner from inside to outside by suturing into the graft vessel.
• Place a stitch from inside towards out in the recipient’s vessel by using the second needle of the same thread.
• Tie the first knot by gently tightening both vessels.
• Stitch the second corner at a 180° angle from the first one and knot it. Using a backhand technique, place other stitches on the posterior wall.
• Place the sutures by starting from the corner towards the center of the anastomosis. Tie the knots consecutively after placing the stitches on the posterior wall.
• Expose the anterior wall by turning the graft over.
• Examine the interior of the anastomosis to make sure that the sutures are placed correctly.
• Complete the suturing of the anterior wall in the same fashion as the posterior wall.

Cuff Technique for Vascular Anastomosis

• Pull the donor vessel through the circular cuff which is held beside the extension handle of the cuff.
• Fold the vessel over the cuff. Bind the folded vessel to the cuff with a nylon tie.
• Insert the cuffed donor vessel into the recipient's vessel and knot it with another nylon tie.

Organ Transplantation

Kidney Graft Procurement Protocol

• Make a midline incision opening the abdominal cavity of the animal.
• Remove the retroperitoneal adipose tissue present around the renal artery and vein.
• Ligate the suprarenal vein and dissect it.
• Separate the renal vessels from the retroperitoneal tissues.
• Isolate the ureters from the retroperitoneal tissue and cut them at the distal end.
• Separate the kidney from the surrounding tissues.
• Use a solution to irrigate the inferior cava vein.
• Place the holding clip on the aorta above the stump of the renal artery.
• Cut the renal vessels nearby the aorta and the caudal caval vein.
• Remove the excess tissue from the distal part of the ureter and renal vessels so that the ends can be prepared for anastomosis and perfuse the kidney with the saline solution through the renal artery.

Kidney Graft Transplantation Protocol

• Open the abdominal cavity of the animal by making a midline incision.
• Clean the capillary bleeding from the incision edges.
• Perform left nephrectomy of the recipient to ligate the renal vessels and ureter and to dissect the renal vessels nearby the kidney hilus.
• Wash the renal vessel stumps with a cold solution.
• Begin the arterial anastomosis by placing two fixating stitches at a distance of 40% in diameter from each other on the recipient's renal artery.
• Place four stitches equidistantly on the anterior side of the arterial anastomosis. Rotate the artery by 180° to complete the stitching of the posterior side of the anastomosis.
• Put five stitches equidistantly on the posterior side of the anastomosis.
• Place two fixating sutures on both corners of the recipient renal vein.
• Perform continuous sutures for the venous anastomosis of the dorsal vein.
• Start the stitching from the upper polus thread towards the distal polus.
• Perform the anastomosis on the anterior vein wall in the same manner.
• Congeal the sutures and take away the fixating stitches. Remove the clips and restore the blood flow to the graft (first from the vein after the artery).
• Stop the bleeding by pressing with warm sterile gauze.
• Get rid of the adipose tissue from the recipient and donor ureters.
• Make two opposite stitches on the ureter-ureteral anastomosis.
• Complete the anastomosis by putting two stitches equidistantly between the recipient and donor ureters.
• Flush the abdominal cavity with warm saline and close it in two layers.

Microsurgical Implantation of the Rodent's Heart Protocol

Pre-surgery Preparations

• Check the donor's physical condition, weigh the donor animal, and calculate the dose of anesthesia.
• Anesthetize the donor using a nose cone.
• Shave the complete incision line and some surrounding skin.
• Treat the skin using alcohol.
• Cover the donor's eyes with a proper ointment and put it onto the heating pad with the backside down.
• Cover the animal with a sterile cloth to provide a sterile field around the incision. Give a single bolus dose of solution for intra-operative pain relief.

Recipient's Preparation

• Open the abdominal cavity by making a midline incision.
• Clean the blood from the incision edges of the vessels and insert abdominal retractors.
• Withdraw the bowels to the left side of the animal and wrap them in wet gauze.
• Uncover the infrarenal portion of the great vessels, 4 cm by length, by dissecting the retroperitoneal fascia.
• Dissect around great vessels and underneath them using two forceps (one straight, one curved).
• Ligate all branches with silk. Separate the remaining tissues surrounding the great vessels and put vascular micro-clamps on the inferior vena cava and the abdominal aorta.
• Put the bowels back into the abdomen and flush them with warm saline during the donor harvesting operation.

Heart Graft Procurement Protocol

• Expose the great vessels by making a midline incision in the abdominal cavity.
• Collect 5 ml of blood from the aorta in the abdomen.
• Close the puncture site immediately with a hemostat to avoid excessive bleeding.
• Inject heparin diluted in cold saline in the inferior vena cava via a needle and congeal the puncture site with the hemostat.
• Expose the aortic arch by dissecting the thymus and performing the midline sternotomy.
• With the help of fine forceps, clean the innominate artery and place a long silk suture around to ease the innominate artery catheterization with a blunt cannula and seal it by ligation.
• Open the innominate artery, insert the blunt cannula heading to the aortic root, and tighten the strap.
• Clip the rest of the aortic arch with a hemostat and administer cold cardioplegic solution slowly via the aortic cannula to arrest the heart and control the coronary vessels washout.
• Meanwhile, incise the superior and inferior vena cava to avoid heart-filling and distention. Apply ice to the heart as soon as possible.
• After heart arrest, take the cannula out from the brachiocephalic trunk.
• Ligate the right and left lungs close to the hilus with a silk suture and cut both lungs away. Do blunt dissection, ligation, and cut off the inferior vena cava, left superior vena cava, and right superior vena cava.
• Clean the aorta and pulmonary artery for blood and excessive tissues. Split the aorta just before the brachiocephalic trunk, the pulmonary artery, and just below its bifurcation, in one cut (straight microsurgical scissors).
• Use the ice-cold cardioplegic solution to keep the heart until re-implantation.

Heart Transplantation Method

• Carefully place the bowels on the recipient's left side and drape them in wet gauze carefully.
• Place vascular micro-clamps on the inferior vena cava and the abdominal aorta on prepared spots.
• With the help of a needle, make a small hole in the center of the clipped aortic segment, then perform a midline aortotomy approximately 4 mm long (to match with donor aorta circumference) and immediately flush the lumen.
• Similarly cut the inferior vena cava; however, the vein incision should be slightly longer (6 mm) to prevent obstruction.
• Place the donor's heart on the right side of the abdomen and correctly orient it so that the graft aorta leads to the abdominal aorta and the pulmonary artery to the inferior vena cava wrap the graft into gauze soaked in ice-cold saline and add some ice if needed.
• Perform the suturing with nylon stitches with a round-bodied needle.
• Place two anchor stitches at the heel and the toe. Stitch continuously from one to the opposite anchor stitch with fine steps.
• After reaching the opposite anchor stitch, knot the continuous sutures to prevent purse stringing the anastomosis.
• After completing one side of the anastomosis, flip the heart to the opposite side of the abdomen and finish the other half of the anastomosis.
• Proceed with the anastomosis of the pulmonary artery and inferior vena cava similar to the aortic anastomosis, but without flipping the heart.
• Begin with the posterior wall and then finish the anterior part of the anastomosis.
• Remove the gauze and ice, and check both anastomoses for any gaps, which can be repaired with a single stitch.
• First, remove the distal clamps and look for any excessive bleeding.
• Later, remove both proximal clamps and hold the graft aorta closed with fine forceps, and let the potential air exit via needle holes.
• Place a dry gauze around the anastomosis. Put a soaked gauze over the heart, and the transplanted heart should start beating within 5s.
• Remove all the instruments and gauze from the abdominal cavity and check for excessive bleeding.
• In case of no bleeding, replace the bowels carefully in their anatomical position, wash them with warm saline, and close the abdominal cavity in anatomical layers.

Heterotopic Abdominal Heart and Lung Transplantation Protocol

Recipient Preparation

Heart and Lung Grafts Procurement

• Steps 1-6 are the same as for the heart transplantation procedure. And the rest of the procedure is discussed below:
• Ligate the inferior and pos-caval lobe of the right lung with a silk suture, so two upper lobes remain untouched.
• Ligate and cut the left lung close to the hilus later. Bluntly dissect, ligate, and cut off the inferior vena cava, left superior vena cava, and right superior vena cava.
• Clean the aorta and cut just before the brachiocephalic trunk.
• Use the ice-cold cardioplegic solution to keep the heart until re-implantation.

Heart and Lung Transplantation Protocol

• Carefully withdraw the bowels to the recipient's left side and cover them in wet gauze.
• Put vascular micro-clips to the abdominal aorta in prepared spots.
• Make a small hole in the center of the clamped aortic segment, then with the help of straight micro-scissors, perform a midline incision of the aorta approximately 4 mm long and immediately flush the lumen with the heparin-saline solution.
• Put the donor heart on the right side of the abdomen and orient it in the direction that the graft aorta leads to the abdominal aorta and cover the graft with gauze soaked in ice-cold saline.
• Perform the stitching with a nylon suture with a round-bodied needle.
• Place two anchor stitches at the heel and the toe.
• After completing one side of the anastomosis, flip the heart to the opposite side of the abdomen and finish the anastomosis.
• Remove the cold gauze and ice, and check both anastomoses for any gaps, which can be repaired with a single stitch. First, remove the distal clamps and look for severe bleeding. Later, remove both the proximal clamps and meanwhile, hold the graft aorta close with fine forceps, letting the potential air exit via needle holes.
• Place dry gauze around the anastomose to support hemostasis. Put gauze soaked in warm saline over the heart, and the transplanted heart should start beating within 5s.
• Take away all the instruments and gauze from the abdominal cavity and once again examine for potential bleeding.
• In the case of proper graft functioning with no bleeding observed, put back the bowels carefully in their anatomical position, wash them with warm saline, and close the abdominal cavity in anatomical layers.

Orthotopic Lung Transplantation Procedure

Triple Axis Stabilizer

Operating Procedure

• Make a skin incision approximately 1 cm below the inferior margin of the scapula and cut the subcutaneous tissue and muscles exposing the lateral chest wall. For hemostasis, use bipolar cautery. For accurate visibility of the hilar structures, open the thoracic cavity in the fourth interspace reaching from the sternum anteriorly to the thoracic vertebrae posteriorly.
• Apply a common wound retractor in the 4th interspace between the 4th and 5th rib to enable access to the left hemithorax.
• Separate the left inferior pulmonary ligament using bipolar cautery and withdraw the left native lung outside the thoracic cavity. Use ordinary Q-tips to anchor the retracted lung.
• Start the dissection with the mobilization of the left phrenic nerve using a non-touch technique and subsequent preparation of the right wall of the left inferior segmental vein and left central pulmonary vein from distal to central.
• Begin with the right wall of the pulmonary vein and dissect the anterior aspect of the vein. If the correct layer of tissue is detected, it is possible to dissect the anterior bronchial wall at the same time moving from right to left.
• Cut the left pulmonary artery and mobilize the vessel as far distally as possible. Observe the fibrous tissue connection between the distal pulmonary artery and the left main bronchus.
• Cutting off this fibrous fixation leads to extravascular length simplifying the subsequent process of cuffing.
• To stop perfusion of the left native lung, ligate the distal pulmonary artery and cut distal to the ligature. To keep minimum blood loss, the pulmonary artery must be clamped before the pulmonary vein as early ligation of the vein may cause pooling of blood in the native lung.
• Dissect the left and posterior venous walls and partially mobilize the main pulmonary vein.
• To gain extra vessel length, which is of importance for venous anastomosis, mobilize the superior and inferior segmental veins to release the pulmonary vein from its surrounding tissue wholly.
• Double-ligate the superior segmental vein as close to the venous trunk as possible using a 7-0 silk suture and cut in-between close to the central ligature to not hamper venous cuffing. The inferior segmental vein is used later for anastomosis and is ligated as distal as possible and cut. At this late point of hilus dissection, focus on small veins, e.g., the inferior segmental vein on its posterior aspect proximal to the ligature, as they can be torn apart as the main pulmonary vein is withdrawn after dissection, leading to major retrograde bleedings from the left atrium.
• All vascular structures have now been dissected and cut. The left main bronchus constitutes the remaining connection to the native lung.
• Release the trachea thoroughly from surrounding tissue and obstruct vessels on the outer bronchial wall with bipolar cautery. Ensuring hemostasis is essential.
• Fix the left main bronchus using a microvascular aneurysm clip, cut the bronchus distally, and remove the native left lung.
• Stabilize the aneurysm clip with the triple-axis precision movement clip holder to limit the movement of the heart and the contralateral lung without touching the heart at all. Introduce the donor lung into the recipient's thoracic cavity and cover the allograft with wet and cooled gauze throughout the process of implantation.
• Remove the fixators of the trachea and the right main bronchus, respectively.
• Shorten the donor’s left main bronchus to an appropriate length and get rid of the remnants of the trachea and right main bronchus.
• To facilitate suturing of the bronchial anastomosis, tapering the donor bronchial stump with a slanting cut from the membranous to the cartilaginous part is recommended.
• Begin the bronchial anastomosis with two interrupted stabilization sutures at 3 o’clock approximating the membranous part of the recipient and donor bronchial wall, respectively.
• Complete the interrupted suture technique clockwise and anastomose the anterior half of the membranous part as well as the cartilaginous part of the bronchial wall using approximately 8 single sutures. Then flip the lung over the heart to allow a better vision of the posterior bronchial wall and finish the airway anastomosis with approximately 8 interrupted sutures starting posterior to the initial stabilization sutures and continuing anti-clockwise.
• Check the airway for patency before the last stitch and remove the intrabronchial fluid.
• Once bronchial continuity has been restored, remove both the stabilization system and the aneurysm clip and re-inflate the lung. From this moment on, mechanically ventilate the transplanted lung; however, it does not take part in the process of oxygenation, as the recipient circulation is not yet reconnected to the donor’s lung.
• Examine the bronchial anastomosis for air leakage with 0.9% sodium chloride at body temperature.
• Flex the lung back with its coastal surface placed on the rat’s back. Place a wet gauze on the lung to re-warm the allograft for the subsequent reperfusion at body temperature.
• Start the vascular anastomoses by reconnecting the pulmonary artery using the cuff technique.
• After the initial process of vessel ligation, keep one thread long so that it can now be “orchestrated” through an intravenous catheter together with the pulmonary artery.
• Similar to bronchial clamping and stabilization, clip the pulmonary artery by using a microvascular aneurysm clamp and stabilize both clip and vessel with the help of the triple-axis stabilizer.
• Incise the ligature and apply heparin topically on the cut edge.
• Turn the blood vessel wall over the cuff and fix it using a 7-0 silk suture ligature.
• Pull the corresponding donor pulmonary artery over this complex of the cuff, evert the recipient’s vessel, and secure the arterial anastomosis with a ligature (7-0 silk suture).
• The continuity between the recipient and donor pulmonary artery is now restored but can only be checked for patency and leakage following the completion of the venous anastomosis by opening the clamps.
• Employ the same techniques as for arterial anastomosis but use a 16G tube as a cuff instead to reconnect the vein.
• To prevent dehydration of the allograft replace the wet gauze at regular intervals. To initiate reperfusion of the transplanted lung remove the venous and arterial aneurysm clips.
• Administer 25 mg of cortisone to all the recipient animals immediately after opening the clamps to prevent hyperacute rejection.
• Reperfuse the allografted lungs for 10–15 min.
• To stop minor bleeding use bipolar cautery or an absorbable fibrillar hemostat. As soon as good reperfusion is obtained, close the thoracic incision with 4-0 Prolene and use 4-0 Vicryl for subcutaneous tissue reattachment and skin closure, respectively.
• Drain the thorax with an 18G intravenous catheter.

Orthotopic Lung Transplantation Method

Animal Preparation

Anesthesia

Intubation

Lung Graft Procurement

• Set the donor animal for median sternotomy.
• Shave the anterior thorax and abdomen and wash the incision sites with a 75 % alcohol solution.
• Make a median skin incision reaching from the jugular notch to the pubic symphysis and extend it laterally from a mid-abdominal level to a T-shaped incision followed by the cutting of subcutaneous tissue and anterior abdominal musculature.
• Cut the peritoneum from the xiphoid to the pubic symphysis and, following the original T-shaped skin incision, continue the peritoneal incision laterally to both sides. To allow direct visualization of the significant ascending and descending retroperitoneal vessels, evert the abdominal viscera extra-peritoneally.
• Separate the abdominal aorta and inferior vena cava and depart both the inferior margin of the liver and the tributary renal vessels.
• Intubate the infra hepatic inferior vena cava using a 27G needle.
• Following anticoagulation, detach the thoracic diaphragm from its coastal attachments and the tripartite of thoracic organs until entirely visible.
• Discontinue the artificial ventilation intermittently to permit separation of the thymus and pericardial tissue from the posterior sternal wall as well as dichotomization of the inferior pulmonary ligaments.
• Perform a median sternotomy using traditional scissors. Apply needle holders on each side of the transected sternum spreading apart the chest walls.
• Excise the thymus to observe the large central thoracic vessels accurately and to allow the detachment of the thoracic aorta ascending from the pulmonary trunk.
• Cut the left atrial auricle to allow drainage of the perfusion solution later.
• Ischemia starts and the donor rat begins to exsanguinate as the left atrial auricle is cut.
• Fill the ice in the thoracic cavity to potentiate cardiac arrest.
• Put wet gauze on the abdominal viscera to avoid dislocation of the intestines into the thoracic cavity.
• Insert a 21G needle into the pulmonary trunk via the subvalvular pulmonary and myocardial valve and infuse 20 ml of an anterograde cold preservation fluid of low potassium dextran glucose with 20μl/20 ml of sodium bicarbonate in the lungs. Stretch the original median incision cranially to the level of the larynx when the perfusion of the lungs is homogenous.
• Depart the infrahyoid muscles medially and ligate the trachea with the lungs fully inflated at 100 % of total lung capacity.
• Cut the trachea proximal to the ligature while keeping the lungs entirely inflated.
• Excise the cardiopulmonary block by cutting the supra-aortic trunks consisting of a brachiocephalic trunk, right subclavian artery, right common carotid artery, and by transecting the thoracic aorta, inferior vena cava, superior vena cava, and pulmonary ligaments.
• Put the explanted cardiopulmonary block in a Petri dish with crushed ice and cooled wet gauze.

Lung Dissection

• Clear away the remnants of the left inferior pulmonary ligament spanning between the left pulmonary vein and the right inferior pulmonary margin.
• Owing to its most anterior position, dissect the left pulmonary vein first.
• Ligate and transect the right inferior pulmonary and vein draining the right postcaval lobe to gain additional vessel length.
• By incising the left central pulmonary vein medial to the ligated venous branch and closing the left atrium, create a long donor left pulmonary vein.
• Congregate the artery carefully and dissect the artery from the pulmonary trunk to the hilus of the left lung.
• To facilitate the process of implantation and to achieve a maximum vessel length, transect the ligament and cut the artery as close to its origins as possible.
• Rinse both vessels to prevent local thrombus formation. As the left pulmonary artery crosses the left main bronchus anteriorly, make the left main bronchus the last structure to dissect.
• Get rid of the peribronchial tissue, which mainly consists of fat. Otherwise, the vision of the bronchial lumen is challenging to gain during anastomosis, and suturing of the airway during anastomosis is impeded.
• Undertake efforts to keep the lungs entirely inflated for as long as possible as very high positive end-expiratory pressure (PEEP) is necessary to detach the atelectasis of a fully collapsed transplanted allografted lung.
• Dissect, ligate, and cut both trachea and right main bronchus distally.

Post-operative Care and Pain Management

Applications

Lymphaticovenous Anastomosis Model in Rat (Yazici & Siemionow, 2015)

Fallopian Tube Anastomosis (2015)

Microcirculatory Models in Plastic Surgery Research (Kusza, Siemionow, & Cyran, 2015)

Transplantation of Small Intestine in Rodents Using Microsurgery (Kudla & Balaz, 2015)

The Microsurgical Groin Skin Flap Microsurgery using the Rat Model (Gurunluoglu & Siemionow, 2015)

Precautions

Summary

• Microsurgery is a surgical technique that combines magnification with advanced diploscopes, specialized precision tools, and various operating procedures.
• Significant purposes of microsurgery are to transplant tissue from one part of the rodent's body to another and to reattach the amputated parts.
• Shorter breeding cycles, faster regeneration, lower costs, and easy handling make rodents ideal for microsurgery research.
• Extended periods of food deprivation can lead to disturbances in balance, metabolic acidosis, and hypoglycemia.
• As small rodents possess high metabolic activity, do not exceed pre-anesthetic fasting beyond 2 hours.
• A complete physical examination of the animal should be performed before the surgical procedure for a smooth duration of general anesthesia.
• The most frequently used anastomotic techniques are interrupted, continuous, and sleeve techniques.
• Microsurgery and free tissue transfer offer plastic and reconstructive surgeons a variety of options. Microsurgical techniques are serving as the backbone of surgical lymphedema treatment.
• During the surgery, take care not to damage the surrounding tissues and muscles.

References

1. Balaz, P., & Kriz, J. (2015). Basic Techniques for Microsurgery Experiment. In P. Girman, P. Balaz, & J. Kriz, Rat Experimental Transplantation Surgery: A practical guide (pp. 49-65). New York: Springer.
2. Gurunluoglu, R., & Siemionow, M. Z. (2015). The Microsurgical Groin Skin Flap Rat Model. In Plastic and reconstructive surgery (pp. 53-62). Chicago: Springer.
3. Kudla, M., & Balaz, P. (2015). Small Intestine Transplantation. In P. Girman, J. Kriz, & P. Balaz, Rat Experimental Transplantation Surgery (pp. 199-213). New York: Springer.
4. Kusza, K., Siemionow, M. Z., & Cyran, M. (2015). Application of Microcirculatory Models in Plastic Surgery Research. In M. Z. Seemionow, Plastic and reconstructive surgery (pp. 71-81). Chicago: Springer.
5. Kwiecien, G. J. (2015). Fallopian Tube Anastomosis. In M. Z. Seimionow, Plastic and reconstructive surgery (pp. 39-43). Chicago: Springer.
6. Yazici, I., & Siemionow, M. Z. (2015). Lymphaticovenous Anastomosis Model in Rat. In Plastic and reconstructive surgery (pp. 33-38). Chicago: Springer.
7. Yu, H., Sagi, A., Ferder, M., & Strauch, B. (1986). A simplified technique for end-to-end microanastomosis. J Reconstr Microsurg, 2, 191–194.

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The novel weight-drop TBI model employs the use of a glancing blow to the head of a freely moving rodent which transmits acceleration, deceleration, and rotational forces onto the brain. Recently, an increasing amount of scientific evidence has come to light indicating that childhood embodies a major risk period for mild traumatic brain injuries resulting from sports-related accidents, motor vehicle injuries, and concussions from falls. However, there has been a significant lack of progress in the area of developing a reliable animal model of mTBI. While the availability of techniques used for inducing moderate and severe traumatic brain injury (TBI) is widespread, very few of these methods have been extended to generate mild, closed head injuries in rodents.

Due to the fact that mild traumatic brain injury (mTBI) is thrice as more likely to occur than moderate and severe brain injury collectively, there is a growing need to develop a dependable model of mTBI to assist the research related to physiopathology, neurobiological outcomes and behavioral consequences, and therapeutic procedures.

When specifically carried out on juvenile rodents, the weight-drop mTBI model generates clinically relevant behavioral outcomes that are depictive of post-concussion symptoms.  The simple and basic nature of this method provides researchers with a dependable model of mTBI that can be utilized in an extensive range of behavioral, molecular, and genetic studies.

Apparatus and Equipment

Weight Drop mTBI apparatus generally consists of two main parts: the U-shaped Plexiglas box and the collection sponge.

The typical weight drop mTBI equipment consists of a U-shaped Plexiglas stand (38 x 27 x 27cm³), a plastic guide tube (1" in diameter, 150cm length, holes at 50cm/100cm), brass weights (1" diameter), a clamp stand, a fishing line, and a collection sponge (38 x 18 x 25 cm³). The lightly anesthetized rodent is placed supine on a scored tin foil with its head directly in the pathway of the falling weight.

Protocol

All animals must be housed with freely accessible food and water along with properly maintained optimal room temperature. 30 days Post-natal, juvenile rodents can be used for taking part in the experiment.

The overall goal of the weight-drop mTBI model is to generate mild traumatic brain injury in juvenile rodents. This is accomplished by first preparing the injury induction platform. Attach a metal loop to the top end of the desired weight allowing the fishing line to be fastened to the weight. A tin foil is scored with a razor-sharp blade making certain that the scored tin foil supports the bodyweight of the rodent. The foil is then placed on the U-shaped stage made of Plexiglas and securely taped to it. The U-shaped plastic stage is placed in the accurate position under the guide tube made of clear plastic. The plastic guide tube is held in place with a clamp stand and positioned above the scored tin foil. The fishing line is then attached through the metal loop to the weight ensuring that the bottom of the weight hangs freely over the scored tin foil. The fishing line is then connected to the clamp stand, and the weight is drawn up through the plastic guide tube with the fishing line and secured in its position with an Allen key pin.

Next, a rodent is taken and lightly anesthetized until it is non-responsive to a paw or tail pinch. The anesthetized rodent is placed face down on the scored tin foil. Once in place, the Allen key pin is pulled allowing the weight to drop and generate a glancing blow to the head. The animal swiftly undergoes a 180° rotation and lands on the collection sponge. The animal is then placed in a supine position and the time it takes to right itself is recorded.

Like any technical method, certain stages of the protocol are predominantly important to generate reliable outcomes. Primarily, the tin foil should be scored properly to produce the desired effect. If the tin foil is not effectively scored, the force exerted by the weight for the duration of the glancing blow will not be sufficient to thrust the juvenile rodent through the tin foil onto the collection sponge. Hence, the desired rotational acceleration and deceleration will not occur. Secondly, during the induction of the mTBI injury, the level of anesthetic that each rodent is exposed to should remain consistent. An important benefit of this technique over countless other TBI methods is the low level and duration of anesthesiology. However, the juvenile rodent must be non-responsive to a toe or tail pinch to make certain that they do not wake up on the stage before the injury is produced.

Applications

Reliable animal models for inducing mild traumatic brain injury can make use of the modified weight drop mTBI device to induce an injury that strongly embodies the pathophysiology and symptoms related to concussions and repetitive mTBI in the adolescent populations. The implications of this technique extend to the therapy of pediatric concussion and mild traumatic brain injury because the model induces clinically relevant symptomology in a heterogeneous fashion.

The present definition of mTBI states that the injury must be the outcome of acceleration and deceleration forces affiliated with blunt trauma. Therefore, the modified weight drop model described above is the ideal method to use in experiments aiming to study mTBI and concussion-like injuries as it makes use of a glancing impact to transmit quick rotational acceleration and deceleration forces to the brain of the unrestrained animal. These forces imitate the biomechanical forces specifically linked to sports-related injuries and motor vehicle accidents (Mychasiuk et al., 2014).

Additionally, this model can be easily modified to observe repetitive mTBI, an incidence that is rising as a severe medical and socioeconomic issue. Evidence indicates that rodents may possibly be exposed to up to 10 distinctive mTBIs with minimum mortality rates. Furthermore, the technique is inexpensive and can be executed quickly, allowing for a highly thorough examination of numerous therapeutic compounds and treatment controls.

Strengths and Limitations

The main advantage of the weight drop mTBI model over others is that the animals are only briefly anesthetized. While the majority of the present TBI techniques impose such intense injuries, it is frequently hard to induce another injury and almost impossible to study repetitive TBI without widespread damage to the entire cortex. However, in the weight drop mTBI technique, a closed head injury is inflicted with no overt damage to the brain which allows the researcher to repeat the injury in the same animal several times.

Another advantage of the weight drop mTBI model is that there's a rotational component to the injury that generates a more clinically relevant concussive-like symptomology. On the basis of the biomechanical pathophysiology of the induced injury and the behavioral outcomes studied, the modified weight-drop mTBI technique has emerged as a dependable model for the examination of pediatric TBI and concussion. Although preliminary studies of this new model have detected some fundamental molecular and structural transformations, further studies will be required to determine how the brain acts in response to a TBI with this injury etiology.

Summary

• The weight-drop TBI apparatus is usually comprised of a U-shaped Plexiglas box and a collection sponge and makes use of a plastic guide tube, brass weights, clamp stand, and fishing line.
• The weight drop mTBI model is a technique that generates a glancing blow to the head of a freely moving rodent.
• The blunt force, in turn, transmits acceleration, deceleration, and rotational forces onto the brain that generates more clinically relevant symptoms.
• The closed head injury allows the researcher to study the outcomes of repetitive mTBI in the same animal.

References

Mychasiuk, R., Farran, A., Angoa-Perez, M., Briggs, D., Kuhn, D., Esser, M. J. A Novel Model of Mild Traumatic Brain Injury for Juvenile Rats. J. Vis. Exp. (94), e51820, doi:10.3791/51820 (2014)

Apparatus And Equipment

Types Of Endotracheal Tubes

Mode Of Operation

Endotracheal Intubation Methodology

Cat, Dog, and Sheep

Pig

Rat

Birds

Rabbit

Intranasal Intubation

Precautions

References

1. Fish, R. E. (2008). Anesthesia and analgesia in laboratory animals. Amsterdam: Elsevier.
2. Flecknel, P. (2009). Laboratory Animal Anaesthesia. Elsevier.

• R-LS-Y  Laser Goggles against green and yellow light.
• Transmittance T<1%, Applicable wavelength range: 528 nm-561 nm
• R-LS-G Laser Goggles against blue and green light.
• Transmittance T<1%, Applicable wavelength range: 200 nm-550 nm

Introduction

Optogenetic laser goggles are specially designed safety glasses that protect human eyes from the hazards caused by the laser. The brightness, beam collimation, and coherence properties of lasers can be detrimental to human eyes. Therefore, the ANSI Z136 1 Safe Use of Lasers standard in the US and the international standard IEC 60825 necessitate laser safety glasses while working in laser-equipped labs. 

The widespread use of lasers generally in medicine, industry, and laboratory research, can cause accidental injuries. Out of the four laser classes, the first two are harmless; however, class 3 and 4 lasers are pernicious to the human eye. The visible and near-infrared radiations are concentrated on the retina of the eye. Therefore, the retina is the tissue most vulnerable to injury caused by accidental laser exposure (Barkana and Belkin, 2000).  Powerful laser exposures (with mid-infrared or ultraviolet light) can also result in cataracts and corneal burns. Symptoms of a laser burn include post-exposure headaches, watery eyes, the appearance of floaters in vision, and the permanent loss of vision in extreme cases (Löfgren et al., 2013). Therefore, the use of Optogenetic Laser Goggles is a prerequisite while working with Optogenetics Laser to prevent the eyes from reversible and permanent laser-induced ocular injuries. 

Apparatus and Equipment

The durable, high-quality acrylic laser safety glasses are lightweight and comfortable to wear. They are specially designed to provide efficient protection against accidental laser exposure. Conduct Science's optogenetic laser safety goggles are available in two variants. One is the R-LS-Y Laser Goggles that protect against green and yellow light. They shield the eyes from rays with a wavelength between 528nm and 561nm. The other variant is R-LS-G Optogenetic Laser Goggles which provide protection against blue and green light and have a wavelength coverage of 200nm to 500nm. Both R-LS-Y and R-LS-G have less than 1% transmittance.

Applications

Optogenetic laser goggles protect the experimenter from accidental laser-induced injuries while working in high-power laser-equipped laboratories. These safety goggles are specially designed to shield the human eye against radiation exposure such as corneal burns, retinal edema and hemorrhage, loss of central vision, cataracts, choroidal infarction, and focal retinal detachment.

Strengths and Limitations

Optogenetic laser safety goggles have numerous advantages, such as ease of wearing, simpler process, and high visible light transmittance. High-power laser beams are attenuated to a level where they can be rendered harmless to the human eye. However, there are a few disadvantages of the product as well. For instance, people with visual impairments need correcting glasses of certain dioptre power.  One can fix the glasses with certain dioptre strengths to make it a little easier. Moreover, some goggles might not protect the eyes from the radiation reaching from the sides. Therefore, while working with high-power lasers, the experimenter must use closed design goggles. 

Summary

• High power lasers (of classes 3 and 4) can be harmful to human eyes.
• Optogenetic Laser Safety goggles must be worn while working in laser-equipped laboratories.
• Accidental laser exposure can cause retinal injuries, corneal burns, and cataracts. 
• Laser safety goggles are easy to wear and have high visible light attenuation. They need to have a closed design to shield against radiations coming to the eyes from the sides.
• People having problems with vision need correcting glasses with particular dioptre strength.

White Series

Black Series

• Low insertion loss
• Good Repeatability
• High return loss
• Stable temperature
• Good mutual insertion performance

Documentation

Introduction and Principle

1. The laser's output wavelength must be in accordance with the sensitivity of the appropriate opsin (light-sensitive proteins).
2. The incident light must activate the opsin.
3. The laser power should be adjustable to compensate for all the losses in the fiber optic delivery system.
4. The power must be stable, and the laser power variation should be less than 2%.
5. The experimenter should also consider the laser's modulation frequency based on opsins' kinetic properties.

1. Laser Diodes: These are economical and available in blue and red wavelengths only. They are modulated directly and have high accuracy and speed. Their stability remains the same at zero power as well as high powers.
2. DPSS (Diode-pumped Solid-state) Laser: It is available in many wavelengths and has a limited modulation capacity. The power stability is also limited.

Apparatus and Equipment

Applications

Strengths and Limitations

Summary

• Laser light, often coupled with optical fibers, is used in optogenetics research.
• The optogenetics laser covers the entire visible light spectrum, including red, green, blue, and yellow.
• The output wavelength of the laser must be in accordance with the sensitivity of the appropriate opsin.
• The laser power must be stable, adjustable, and compensate for all losses that occur during delivery.
• There are two types of optogenetics laser, laser diodes, and DPSS.
• Optogenetics laser has a wide range of applications in the field of neurobiology.

References

1. Mahmoudi, P., Veladi, H., & Pakdel, F. G. (2017). Optogenetics, tools and applications in neurobiology. Journal of medical signals and sensors, 7(2), 71.
2. Arrigoni, M. (2016). Lasers for Optogenetics and Multimodal Microscopy: Next ultrafast laser generation opens up a diversifying and dynamic range of non‐linear imaging applications. Optik & Photonik, 11(2), 27-30.

Specifications

Evaporation Tank Details

Features

• Multi-channel design can meet the needs of multiple animals(1-5 small animals) simultaneously
• Accurate oxygen flow meter, Flow control range: 0-4L/min available
• High accuracy vaporizer (Each one has an independent test report)
• Gas toggle switch, button switching gas path, more than 100,000 times of service life.
• Quick oxygenation switches to remove residual anesthetic mixture in the tube or anesthesia induction chamber.
• Oxygen or air can be chosen as a gas supply.
• Easy Fill, Key Fill for adding drugs is optional.
• Can be upgraded into a portable anesthesia machine to save space and facilitate movement
• The gas flow of the induction chamber can be adjusted independently, ranging from 0 to 2.0 L/min.
• Toggle switch, quick switching of the gas circuit, 100 thousand times of life

Protocol

1.  Accommodates animals ranging in size from neonate mice to rabbits.
2.  Six outputs supply anesthesia to the induction chamber and up to five masks.
3.  Incorporates two flowmeters: one controls the vaporizer (0-4000cc/min) and the other one controls the induction chamber (0-2500cc/min).
4. Allows investigator flexibility in designing protocols without calculations.
5.  Each output (mask) is individually controlled with a simple ON/OFF toggle switch.
6.  Push-button oxygen flush purges induction chamber before removing animals.
7. Carry handle facilitates safe, easy transport of the unit.
8. Available in both tabletop and wall mount, and optional post mount with 5-leg spider base (ask for  more information)
9. The built-in temperature compensator ensures the stable concentration of anesthetic gas at different temperatures and flow rates, the flow range is 0.2-10L/min;

Additional Features & Information

Our RWD-R550 Multi-output Anesthesia Machine (size 13x6.75x12.8inch) is our most advanced system and offers highly efficient, a precise gas flow that is easy to use.

1. Mixed anesthetic gas output pressure range △ P ≤ 2.5kPa;
2. Applicable temperature range: 10-35 ℃;
3. Good tightness, internal 50kPa pressure can be maintained, zero leakage, safe and reliable;
4. Internal capacity is not less than 120ml, and the range of isoflurane vaporizer concentration is 0-5% (Sevoflurane: 0-8%);
5. Anti-accident open lock structure and closed state security protection structure to ensure safety

Note: manifold with 4LPM Flowmeter Vaporizers, Inlet/Outlet End Caps, are included in the RWD-R550 series anesthesia machine

Summary

• Multi-output Design: Can anesthetize 1-5 animals (rats, mice, cats, rabbits) simultaneously.
• Independent Control: Each channel has its own control, allowing for efficient and flexible research.
• Accurate Vaporizer: High precision with independent calibration for reliable output.
• Gas Flow: Adjustable flow for both induction chamber and masks (0-4L/min, 0-2L/min for induction).
• Durable: Over 100,000 toggle switch operations, with options for oxygen flush and portable upgrades.
• Safety: Stable anesthetic concentration with temperature compensator and anti-accident features.

1. RWD-RM300 Veterinary Monitor  (It can be used to test the parameters like SpO2, ECG, RESP, NIBP,TEMP and HR)
2. RWD-R620 Anesthesia Machine
3. RWD-R409 Veterinary Ventilator

Features:

1. The built-in temperature compensator ensures the stable concentration of anesthetic gas at different temperatures and flow rates, the flow range is 0.2-10L/min;
2. Mixed anesthetic gas output pressure range △ P ≤ 2.5kPa;
3. Applicable temperature range: 10-35 ℃;
4. Good tightness, internal 50kPa pressure can be maintained, zero leakage, safe and reliable;
5. Internal capacity is not less than 120ml, and the range of isoflurane vaporizer concentration is 0-5% (Sevoflurane: 0-8%);
6. Anti-accident open lock structure and closed state security protection structure to ensure safety

Anesthesia Machine Pole Mount

Compact Small Animal Anesthesia Device

5ch Multi-output Small Animal Anesthesia Machine

Anesthesia Machine: Enhanced Small Animal Anesthesia Device

1. Automatic flow rate and temperature compensation.
2. Flow rate 0.2-10,000cc/min and temperature 10-35℃
3. Safe and reliable tanks with 50kPa pressure tolerance.
4. 120cc Liquid capacity
5. Output precise and adjustable concentration
6. Integrated safety lock to prevent unintentional engagement.
7. A large sight glass for at-a-glance liquid level monitoring.
8. Use with Isoflurane 0-5% output or sevoflurane 0-8% output,
9. Pour-fill and easy attachment-fill styles are available for each vaporizer.

Introduction

Apparatus and Equipment

Mode of Operation

Precautions

References

1. Chakravarti S, Basu S (2013). Modern anaesthesia vapourisers. Indian J Anaesth. 57(5):464-71. DOI: 10.4103/0019-5049.120142.
2. Fish, R. E. (2008). Anesthesia and Analgesia in Laboratory Animals. Amsterdam: Elsevier.
3. Flecknel, P. (2009). Laboratory Animal Anaesthesia. Elsevier.

1. Pour-Fill dosing method (IP): When the Pour-Fill evaporation tank needs to be dosed, open the cap of the anesthetic bottle and pour the anesthetic directly into the dosing port of the evaporation tank at an appropriate flow rate.
2. Easy-Fill dosing method (IE): Easy-Fill evaporation canister needs to use the dosing adapter. When using it, you need to install the dosing adapter to the anesthetic bottle first, align the adapter to the evaporation can’s dosing port, and press the anesthetic bottle At the end, the sealed dosing can be realized.
3. Key-Fill dosing method (IK): Key-Fill evaporation tank requires a key filling adapter, which can be inserted into the evaporation tank to ensure slow dosing and minimize the waste of anesthetics. This method is a completely enclosed dosing method.

1. Cagemount: conventional loading inlet and outlet ports, fixed by screws.
2. Selectatec: Suspended component installation, evaporator knob adjustment, ready to take away.

Introduction

Mechanical ventilation plays a crucial role in the survivability of patients and animals with poor respiratory functions. Ventilation in these situations can be either support or completely replace spontaneous breathing. Inadequate gas exchange indicates the need for mechanical ventilation. This can be indicated by severe hypoxemia despite oxygen therapy (Pa_O2 <60 mm Hg), severe hypoventilation (defined as P_CO2 >60 mm Hg), excessive work of breathing, or severe circulatory shock. Hypoxemia can result from inadequate inspired oxygen, the elevation of the partial pressure of carbon dioxide (PCO2) (hypoventilation), or venous admixture.

Mechanical ventilation also is a common component of anesthesia practices due to the respiratory system depression caused by most anesthetic agents. Further, to make the handling of animals easier, neuromuscular blocking (NBM) agents are often used during anesthesia and surgical procedures. These agents induce paralysis and hence require the use of mechanical ventilators to ensure adequate respiratory functions. Though specific animal mechanical ventilators are available, mechanical ventilators no longer used for humans can also be repurposed for use in animal-based studies. Mechanical ventilators are selected based on the size and weight of the animal.

The most common design of large animal ventilators is the bag-in-the-box design system. These systems rely on compressed gas to power a bag or bellow to empty and push a volume of gas into the lungs in a large animal circle system. They provide an isolated interface between the breathing system gases and the gases that power intermittent positive pressure ventilation (IPPV). The bag-in-the-box systems, however, do not offer positive end-expiratory pressure (PEEP) or are limited to a value of 10 cmH₂O. Further, they tend to be rather noisy due to their significant dependence on compressed gas to operate. The latest large animal ventilators make use of electrical motors and arrangements for the movement of the gas in the system to overcome the shortcomings of the bag-in-the-box systems. Compared to their older counterparts, the latest mechanical ventilators’ microprocessor capabilities offer a direct and wide range of control of the ventilation parameters. Apart from the achievability of desired patterns, the latest systems also allow a variety of ventilation modes including a wide range of PEEP and do not require disconnection or switching of breathing bags during spontaneous breathing.

Apparatus and Equipment

Conduct Science offers a mechanical ventilator for large animals weighing up to 100 kgs. The ventilator can be used for large mammals including rabbits, dogs, monkeys, pigs, sheep. (For small animal ventilators click here and here)

The 5.8 kg device has a dimension of 22.5 x 2.8 x 3.2 cm. The ventilator comes with two interchangeable bellows of capacity 0 to 300 ml and 300 to 1500 ml. The mechanical ventilator is also equipped with dual LED and sound alarm modifiable trigger settings for monitoring proximal airway pressure in real-time. It also has provisions for multiple gas supplies; O₂, nitrogen, and clean, dry air. Apart from these features, the mechanical ventilator has the following technical specifications and features,

Mode Of Operation

As opposed to manual ventilation, mechanical ventilators allow the precise control of the duration of inspiration and expiration, the volume of gas delivered to the lungs, and the pressure reached in the airway during inspiration. Ventilators achieve controlled ventilation by application of intermittent positive pressure to the airway, which can be accomplished by directly delivering gas to the anesthetic breathing system or indirectly by compressing a rebreathing bag or bellows.

The first step of ventilating an animal involves the calculation of the required tidal volume (approximately 7 to 10 ml/kg body weight) and selection of the respiratory rate. Generally, the rate is selected slightly lower than the normal resting rate of the animal when it’s conscious. In case the ventilator does not have direct settings for tidal volume and respiratory rate then they are most likely to have settings for inspiratory time and inspiratory flow rate.

Further, some ventilators may only provide controls for inspiratory time and inspiratory: expiratory (I: E) ratio. The ratio is usually set to 1:2, however, a ratio of 1:3 and 1:4 can also be used without causing significant cardiac depression while maintaining inflation pressures below 20 cm water. Apart from these variables, maximum inspiratory pressure should also be set on the ventilator at less than 15 cm water for small animals and not more than 25 cm water for large animals, under most circumstances.

Protocol

Mechanical ventilation in animals is often used in researches that require long-term use of anesthesia or when neuromuscular blocking agents are used during surgeries. Controlling ventilation relies on the use of an endotracheal tube or the placement of a tracheal cannula if the animal is not intended to recover. These methods prove successful in comparison to using a face mask, which risks the inflation of the stomach, and laryngeal masks. Once the tube or cannula is placed, it is then connected to the ventilator to allow mechanical ventilation.

Prior to beginning the procedures of mechanical ventilation, it should be ensured that all the equipment and instruments used are thoroughly cleaned. The subject must be adequately and appropriately anesthetized before and during the entirety of the procedure to ensure humane treatment.

Endotracheal Intubation

Endotracheal intubation is assisted by a laryngoscope. The type of endotracheal tube and laryngoscope (Miller laryngoscope and Macintosh laryngoscope) used are dependent on the size and weight of the subject. An appropriate length for an endotracheal tube is usually the approximate distance from the external nares to just anterior to the thoracic inlet. The tubes should be lubricated with a small quantity of lidocaine gel to allow easy passage. Further, cough and swallowing reflexes should be adequately numbed. It is recommended that the animal is administered with oxygen for about 2 minutes before being intubated to slow the advent of hypoxia should the larynx be obstructed. The detailed methodology of endotracheal intubation of different animals can be found here.

The procedure of insertion of the endotracheal tube is performed by resting the animal on its back (generally), with the neck and head flexed. The laryngoscope is then advanced over the tongue towards the pharynx. Care must be taken to extend the tongue and avoid damaging the surface of the teeth of the animal. Following successful laryngoscopy, the endotracheal tube is advanced into the trachea.

Performing Laryngoscopy Using MacIntosh Blades

1. Place the subject on its back.
2. Hold the laryngoscope in your left hand.
3. Extend the subject’s tongue forward and to the left.
4. Slowly insert the blade into the right side of the subject’s mouth.
5. Advance the blade inward and midline towards the base of the tongue.
6. Place the tip of the blade in front of the epiglottis in the vallecula.
7. Apply pressure caudally and upward with the handle placed at an angle of 45 degrees.
8. Lift the handle until vocal cords are visualized.
9. Perform intubation under direct visualization of vocal cords.
10. Withdraw the blade while firmly holding the endotracheal tube in place.

Tracheostomy

1. Adequately anesthetize the subject and place it in the supine position with its limbs restrained.
2. Extend the neck. Remove the hair below the mandible and ensure local asepsis of the area.
3. Make an incision midline below the neck.
4. Separate the salivary glands and fix the muscles surrounding the trachea with sutures.
5. Once the trachea is visualized, make an incision between the fourth and the fifth tracheal ring and insert the cannula.

Applications

Evaluation of effects of mechanical ventilation on coagulation and fibrinolysis in ARDS model

Wang and Shen (2015) evaluated the effects of ventilation using different tidal volumes on coagulation and fibrinolysis in rabbits with acute respiratory distress syndrome (ARDS). ARDS was induced using a sequential injection of 0.1 mL/kg oleic acid (OA) and 500 μg/kg lipopolysaccharide (LPS) via an auricular vein.  Healthy adult male rabbits were divided into 5 groups of which one group served as a sham and another as a model group. Animals in the other groups were either ventilated on low V_T (6 mL/kg), routine V_T (10 mL/kg), or high V_T (15 mL/kg). The experiment was concluded after 6 hours of LPS injection. Data collected based on the blood samples showed that mechanical ventilation with low V_T improved coagulability and fibrinolytic status, while ventilation with routine V_T had little effect on coagulability and fibrinolytic status. However high V_T ventilation significantly deteriorated the coagulability and fibrinolytic function in ARDS.

Investigation of the effect of airway humidification during mechanical ventilation

Jiang et al. experiment aimed to establish a relationship between airway humidification and mechanical ventilation-induced lung inflammatory responses. For their investigation male Japanese white rabbits were divided into control animals, dry gas group (no humidification), and experimental animals. The control animals were sacrificed soon after anesthesia while the dry gas and experimental group were on mechanical ventilation for 8 hours. The experimental group was further divided into groups that underwent ventilation with humidification at 30, 35, 40, and 45°C. Results of the investigation showed that the dry gas group showed increased tumor necrosis alpha factor levels in bronchoalveolar lavage fluid (BALF) compared to the controls. At a humidification temperature of 40°C, it was observed that tumor necrosis factor-alpha and interleukin-8 levels in the BALF reached baseline levels.  Further, at 40°C humidification pathological injury was reduced in comparison to other groups based on the histological score. It was concluded that appropriate humidification reduced inflammatory responses, reduced damage to cilia, and reduced water loss in the airway.

Assessment of the process of de-recruitment in normal lungs ventilated for 16 hours

Tucci et al. evaluated lung dysfunction and inflammation in anesthetized sheep ventilated at 8 mL/kg V_T and PEEP at zero. Based on the emission scans taken at baseline and after 16 hours, it was observed that the gas fraction decreased in dorsal (0.31±0.13 to 0.14±0.12, P<0.01), but not in ventral regions (0.61±0.03 to 0.63±0.07, P=nonsignificant), with time constants of 1.5 – 44.6 hours. Further, shunt increased in dorsal regions (0.34±0.23 to 0.63±0.35, P<0.01), though no change in the vertical distribution of relative perfusion was observed. After 16 hours, an increase in the average pulmonary net F-fluorodeoxyglucose uptake rate (3.4±1.4 to 4.1±1.5 10(-3)/min) was observed in the regions of interest along the ventral-dorsal direction. Also, in the corresponding average regions of interest, the F-fluorodeoxyglucose phosphorylation rate increased from 2.0±0.2 to 2.5±0.2 10(-2)/min (P<0.01).

Assessment of effects of mechanical ventilation on abdominal edema and inflammation in sepsis

Lattuada et al. speculated that mechanical ventilation led to an increase in abdominal edema and inflammation in sepsis. This speculation was tested in the porcine endotoxemia model. Anesthetized pigs were divided into healthy controls and endotoxemia groups. Healthy control pigs either breathed spontaneously with continuous positive airway pressure (CPAP) of 5 cm H₂O or were mechanically ventilated with PEEP of 5 cm H₂O for 5 hours. The endotoxin group received intravenous endotoxin at a dose of 15 μg/kg/h for 2.5 hours during mechanical ventilation (MV) with PEEP of 5 cm H₂O (MV + PEEP5). Following this, they were divided into groups that were maintained on MV + PEEP5, MV with PEEP of 15 cm H₂O, or spontaneous breathing (SB) with CPAP of 5 cm H₂O for another 2.5 hours. Isotope technique was used to estimate abdominal edema formation and inflammatory markers were measured in liver, intestine, lung, and plasma. The data obtained from the investigation suggested that mechanical ventilation with positive end-expiratory pressure increased abdominal edema and inflammation in the intestine and liver by increasing systemic capillary leakage and impeding abdominal lymph drainage.

Precautions

Animals should be handled appropriately, and humane treatment must be ensured throughout the procedures to avoid undue injury or stress to the animal. All equipment and apparatus must be thoroughly cleaned before use to avoid chances of infections and other complications. Animals should be anesthetized appropriately, and measures to maintain the appropriate depth of anesthesia should be followed. It is important to monitor the anesthesia and vital signs of the animal throughout the procedure. Further, the tubes leading to the animal containing inspired air and the tubes returning from the animal containing expired air should be kept as short as possible to minimize dead space. If the animal is expected to recover from the procedure, an appropriate recovery area should be maintained, and a recovery routine should be followed.

Strengths and Limitations

Strengths

Modern mechanical ventilators, unlike their older versions, rely on electricity to operate. In addition to power backups and other modes of power, modern ventilators also allow the option of manual ventilation. Further, as opposed to the older versions, modern ventilators allow more control over ventilation parameters. In clinical practices, a single device for mechanical ventilation makes the technical features of the ventilation procedure simple. The device mitigates the need for a separate piece of equipment and the hassle of shifting the subject during the experiment as seen with manual devices. The programmability of mechanical ventilators also helps guarantee the standardization of protocol and repetition of the procedure between subjects; This capability helps reduce ambiguity in research and promotes the accuracy of results. The diversity in sizes of the mechanical ventilators permits researchers to research animals of different weights and sizes without any restrictions.

Limitations

Classical large animal ventilators do not generally offer positive end-expiratory pressure or have PEEP restricted at 10cmH₂O. Further, these systems are dependent on a substantial amount of compressed gas in addition to being noisy. As with any equipment, the mechanical ventilator must be monitored to avoid any issues that can arise during assisted ventilation. It is important that breathing motion and lung volume are monitored throughout the various stages of the breathing cycle. Prolonged ventilation may affect the survivability of the subject. Additionally, extrapolating the research results to other animals of different ages or species has its limitations.

Summary

1. Mechanical ventilation is used to support or replace spontaneous breathing. They often are part of the anesthetization process, especially when neuromuscular blocking agents are used.
2. Bag-in-the-box design is a commonly used system of mechanical ventilation.
3. Mechanical ventilators can potentially cause ventilator-induced lung injury, leading to the rapid type of disuse atrophy in respiratory-related muscles, barotrauma, and impairment of mucociliary motility in the airways.
4. The use of mechanical ventilators has been extended to the research of ventilator-induced lung injuries and for understanding the relationship between ventilator settings and certain biological responses.
5. Mechanical ventilation is performed either via endotracheal intubation or by cannulating the trachea (tracheotomy).
6. It is important that the animal is sufficiently anesthetized before ventilation and is placed in an appropriate position to prevent injury.
7. Good anesthesia practice should involve appropriate anesthesia management and monitoring techniques, and recovery care.
8. Neuromuscular blocking agents only produce paralysis. Thus efforts to monitor the depth of anesthesia is critical for the humane treatment of the animals.
9. Anesthetization of pregnant animals should be done with extreme caution to prevent undesirable effects.

References

1. Hopper K, Powell LL (2013). Basics of mechanical ventilation for dogs and cats. Vet Clin North Am Small Anim Pract. 43(4):955-69. DOI: 10.1016/j.cvsm.2013.03.009.
2. Jiang M, Song JJ, Guo XL, Tang YL, Li HB (2015). Airway Humidification Reduces the Inflammatory Response During Mechanical Ventilation. Respir Care. 60(12):1720-8. doi: 10.4187/respcare.03640.
3. Lattuada M, Bergquist M, Maripuu E, Hedenstierna G (2013). Mechanical ventilation worsens abdominal edema and inflammation in porcine endotoxemia. Crit Care. 17(3):R126. DOI: 10.1186/cc12801.
4. Tucci MR, Costa EL, Wellman TJ, Musch G, Winkler T, Harris RS, Venegas JG, Amato MB, Melo MF (2013). Regional lung derecruitment and inflammation during 16 hours of mechanical ventilation in supine healthy sheep. Anesthesiology. 119(1):156-65. DOI: 10.1097/ALN.0b013e31829083b8.
5. Wang X, Shen F (2015). [Effects of mechanical ventilation with different tidal volumes on coagulation/fibrinolysis in rabbits with acute respiratory distress syndrome].Zhonghua Wei Zhong Bing Ji Jiu Yi Xue. 27(7):585-90. DOI: 10.3760/cma.j.issn.2095-4352.2015.07.009. Chinese.

Holder secures the cordless microdrill to stereotaxic instruments. controls the depth of drilling by the manipulator. Easy and accurate operation protects brain tissue from damage caused by excessive drilling. Holds the microdrill with diameter about 19.5mm

Principle

Mode of Operation

Precautions

References

Performing Laryngoscopy Using Miller Blades

• Place the subject on its back.
• Hold the laryngoscope in your dominant hand.
• Extend the subject's tongue forward and to the left.
• Slowly insert the blade into the right side of the subject’s mouth.
• Advance the blade inward and midline towards the base of the tongue.
• Place the tip of the blade under the epiglottis.
• Apply pressure caudally and upward with the handle placed at an angle of 45 degrees.
• Lift the handle until vocal cords are visualized.
• Perform intubation under direct visualization of vocal cords.
• Withdraw the blade while firmly holding the endotracheal tube in place.

Precautions

References

Includes

The Y-maze is often preferred to the T-maze because gradual turns decrease learning time as compared to the sharp turns of the T-maze. It is also a smaller maze* to allow fewer degrees of freedom of movement, focusing the animal on the task at hand. The Y Maze can also be baited with food for rewarded alternation. Food wells are standard 1cm deep.

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Modifications

Backlights

$295

Available in fluroscent white, blue, red, IR

Stand

Mouse: 32cm, $150

Rat: 45cm, $200

Escape Tubes

Mouse: 4cm diameter, $350

Rat: 8cm diameter, $400

3x tubes

Cue Lights

3x, push for on/off

$250

Food wells

1, 2, or 3 arms

$100

Water Sealing

$300

Doors

To fit.

$100

Goal Boxes

To fit at the end of each arm.

$250

Introduction

The Y-Maze is a modified version of a T-Maze. Unlike the T-Maze that has sharp turn angles, the Y-Maze provides a much more natural turn of the arms (120 degrees in the Y-maze compared to 90 degrees in the T-Maze). The natural tendency of rodents to explore new environments serves as the basis of the Y-Maze task. This exploratory task involves many parts of the subject’s brain including the hippocampus, septum, basal forebrain and pre-frontal cortex. Many types of research involving neurodegenerative and neuropsychiatric diseases focus on the hippocampus as lesions or damage to it is believed to affect cognition and spatial learning. Thus Y-Mazes are part of the widely used behavioral tasks in the assessment of spatial learning and memory.

The Y-Maze, just like the T-Maze, offers only two choices to the subjects. In a baited Y-Maze task, once the subject has retrieved the reward from one of the arms, it is expected to visit the next arm on the next trial. This simple behavioral task tests cognitive function based on the ability of the subject to remember spatial locations requiring hippocampal-dependent reference memory. Administration of certain drugs or in disease model can affect this ability.

Spence and Lippitt (1946) used a simple Y-Maze as an experimental test for the sign-gestalt theory of trial and learning that was suggested by Tolman. The Y-Maze goal arms were baited with water in one arm, and food rewards in the other. Subjects that were either thirst or food deprived were used to put to the test Tolman’s idea of learning. Dember and Fowler (1959) studied alternation behavior using Forced Alternation and Spontaneous Alternation tasks and suggested their findings in the T-Maze can also be replicated on a Y-Maze. Since then the Y-Maze has been used in studying various aspects of spatial learning, memory, and cognition.

The Y-Maze is usually a symmetrical capital ‘Y’ shaped maze with the arms spaced at an angle of 120 degrees. This construction makes the Y-Maze ideal for the Continuous Alternation protocol. Modifications such as guillotine doors and escape tunnels can easily be added to the maze. The Y-Maze can also be adapted for investigation of spatial learning specifics in different animals and disease or intervention models.

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History

Origin

In 1946, Spence and Lippitt made use of a simple Y-Maze that had one arm baited with water and the other arm baited with food. Their experiment was an investigation into the ongoing debate among psychologists regarding Thorndike’s law of effect and Tolman’s theory of learning. Spence and Lippitt’s experiment attempted to translate the theoretical representation of simple trial-and-error learning as suggested by Tolman. In their experiment, rats were divided into two groups and given 12 days’ experience of thirst motivation in a simple Y-Maze. The maze had one path that led to water and the other to an empty box for one group and food in the case of the other group. Their experiment’s result was in disagreement with the Tolman theory as both the groups chose the arm baited with water.

In the late 1950s, Dember and Fowler researched the alternation behavior in animals. Their series of published papers described spontaneous vs. free alternation behaviors and the influence of rewards and reinforcement on these tasks (Dember and Fowler 1959, Fowler et al.,1959a, Fowler et al.,1959b). Since then, the Y-Maze and T-Maze have been developed as relatively simple and have been widely used for assessing spatial memory.

In 1961, Hellyer and Straughan tested the hypothesis that alternate behavior is learned by first pre-training food and water deprived rats on a rewarded straight runway or table top and then testing them on a baited Y-Maze. The resulting analysis showed that animals trained on the table top alternated significantly more than those trained on the straight runway.  The data also revealed that the animals trained on the table top exhibit a relatively high percentage of alternation at the outset but alternate progressively less with trials. The researchers, based on the results, concluded that the effects of preliminary training were temporary.

Developments

The Y-Maze also serves as a simple tool in understanding the role of different parts of the brain in spatial learning. In 1986, Cazala tested the effects of electrical stimulation of certain brain areas of mice and subjected them to a spatial discrimination task in a Y-Maze. Each subject received one electrode implanted in the following structures: the medial hypothalamus, the medial lemniscus, the lateral tegmentum, the reticular formation, the dorsal part of the mesencephalic central gray area and the lateral hypothalamus. In the Y-Maze trials, one-half of the animals had to learn to self-stimulate by interrupting the photocell beam situated in the right arm, whereas the other half had to go to the left arm. The results showed that the animals were able to discriminate between the reinforced arm and the non-reinforced arm of the Y-maze to self-administer the stimulation.

The effects of chronic stress on spatial memory and cognition were investigated by Conard et al. in a Y-Maze that had extra-maze visual cues only. The results of the test revealed that chronic stress impaired spatial memory in the subjects. Interestingly, however, stressed females showed an improved performance in the latter minutes of the maze in contrast to the stressed males that continued to visit the arms at random. Further investigation of chronic stress by Conard and Wright (2005) showed that chronically stressed rats performed well in the intra-maze cue version of the Y-Maze. The subjects spent a similar amount of time exploring both the arms in the traditional Y-Maze while in the cued Y-Maze the rats spent more time exploring the novel arm. This behavior supports the hypothesis that chronic stress leaves novelty-seeking behavior intact while impairing spatial memory.

A 2009 paper by Hoeffer et al. tested the hypothesis that FKBP12-deficient mice showed increased preservation or decreased inhibition by using a water Y-Maze. In the trial, the subjects were trained to locate an escape platform in the goal arms. When the subject was successful in finding the escape platform, the subjects were given a reversal test where the escape platform was reversed to the alternate arm and their ability to learn the new location was measured. The results of the test revealed that FKBP12-deficient mice have impaired performance in learning and memory assays that require inhibition of previously reinforced responses.

Recent Developments

The role of cerebellar cortex in conditioned goal-directed behavior was studied by Burguière and colleagues (2010) using a water Y-Maze. They tested heterozygous transgenic L7-PKCi mice in a stimulus-dependent water Y-maze conditioning task resulting in the conclusion that the mutant mice were able to acquire the stimulus-response association though they showed reduced optimization of motor performance. This result suggested that PKC-dependent process in cerebellar Purkinje cells is required for optimization of motor responses.

A 2012 study by Bai et al., showed the anticipatory activity in rat’s medial prefrontal cortex using a Y-Maze task. The study aimed to explore the encoding, by the neuronal ensemble, of a working memory event. A similar study was also conducted by Yang et al. using a two-choice spatial delayed alternation task in a Y maze.

Due to the ability to readily test functions of the hippocampus in learning and memory, the Y-Maze has been extensively used in testing the effects of drugs and toxins on spatial memory and age-related cognitive decline. The effects of melatonin on attentional, executive and working memory processes were tested by Liet et al. Spontaneous Alternation Y-maze task was employed to test the immediate working memory in rats administered with acute melatonin. Result analysis showed that acute administration of melatonin did not alter any of the processes in rats.

Yin et al. studied the effects of paradoxical sleep deprivation on the Y-Maze task performance. Using Forced Alternation, they evaluated the short-term spatial working memory of the subjects. Although initially, the mice showed no impairment, after 7 days of deprivation, working memory impairment was observed. Another study looked into the neuroprotective effect of hispidulin. Huang et al. used an aged mice model and tested the sevoflurane administered rats in a Y-Maze. The rats showed significant memory impairment which was remarkably reversed by pre-treatment with hispidulin.

Apparatus & Equipment

The apparatus is a 3-arm maze shaped like a capital ‘Y’ with each arm spaced at an angle of 120 degrees. The arm lengths range between 30 to 50 cm and are usually symmetrical, although asymmetrical mazes are also used. The pathway widths are approximately 10 cm. The maze is usually used as an enclosed maze with approximately 30 cm high walls and the entire maze is elevated to a height of 50 cm from the floor. The two-goal arms may contain a well with a food reward; alternatively, the goal arms may contain an exit tube that allows the animal to escape the maze (Deacon 2013). Generally, there are guillotine doors at the entrance of each arm that can be used to confine the animal to a specific arm or prevent the animal from entering one of the goal arms. The maze is usually painted a dark color to prevent the animals from feeling extra anxiety while performing the task. Intra-maze cues can also be used to help animals distinguish and remember goal arms.

Fully automated Y-Mazes are also available which detect the location of the subject within the maze and control the opening and closing of the arm doors as a trigger response. The automated maze is also capable of detecting food rewards.

The Y-Maze can also be adapted to include the benefits of water mazes and used for rodent models (Burguière et al.,2010, Deacon 2013). The Y-Maze has also seen adaptations as an aquatic model (Natt et al.,2017, Abreu et al.,2017) and for use with arthropods ( Jaworski et al.,2017, Simonnet et al.,2014).

To avoid shadows in the maze, the Y-Maze should be well lit from above. Proper lighting also ensures that the subject is able to see the food rewards. Tracking software such as Noldus Ethovision XT, mounted above the maze assist with live scoring, tracking and recording the subject and its movements within the maze. The apparatus must be cleaned thoroughly before and after each trial to limit influence from any residual stimuli from previous trials.

Training Protocol

The Y-Maze has been widely utilized in assessing spatial memory and learning in animals, usually in control versus disease model or intervention group. The task involves observation of the ability of the subject to remember the previously visited arm. The task takes advantage of the natural explorative nature of rodents and the idea that they tend to use an optimal search strategy to obtain food with minimal effort.

The performance in the task weakens or decreases for subjects with hippocampal lesions or damage as seen in neurodegenerative or neuropsychiatric diseases or as a result of natural aging.

Several protocols exist to be used with the Y-Maze, though the two most commonly used protocols are the Rewarded Alternation task and Spontaneous Alternation task. For the Rewarded alternation task, the experimenter decides which arm is the “correct choice” whereas the Spontaneous Alternation task uses the subject’s natural explorative drive and allows it to choose which arm to explore first. Spontaneous alternation task has been shown to be more successful in the assessment of subjects with hippocampal lesions as these animals tend to develop a side preference (Deacon and Rawlins 2006). The task is also used to quantify cognitive deficits in transgenic strains of mice and evaluate effects of drugs and toxins on cognition.

Another protocol used with the Y-Maze is the Delayed Alternation task. The Delayed Alternation task allows assessing spatial working memory by first allowing the subject to explore a baited arm of its choice and removing the subject once the choice is made and limiting it to the start box of the base arm using a door. This trial is followed by a formal trial, after a predetermined delay between the two trials, wherein the subject is reintroduced to the maze by opening the start arm door and is expected to choose the arm that it did not choose in the previous trial.

Pre-training for the Y-Maze

Rodents tend to be wary of eating anything new. Thus it is important to familiarize the subject with the food rewards used, prior to the testing, within a familiar environment (housing cage). Food rewards can be solid (such as sweetened breakfast cereal) or liquid (such as chocolate milk.) Generally, liquid rewards are used in case of drug testing as some drugs may make eating solid food an unpleasant experience for the subject. Subjects are usually maintained at about 85 to 95% of their free-feeding weight throughout the training and testing phases.

For the food-deprived task, the food-scavenging behavior is encouraged by depriving the subject of food the night prior to the testing. Subjects may be familiarized to the maze in pairs (usually cage-mates) to reduce the anxiety in a novel space during the initial phase of exploration. The acclimation is usually done individually, although, this may require more time than paired familiarization. The maze is scattered with food rewards, and the subject is placed in the start arm and allowed to explore the maze freely. The food rewards are replaced on consumption. The familiarization procedure is repeated four times with at least ten minutes between each exposure. The acclimation period usually lasts 1 to 2 days. Next, one of the arms is blocked, and the open goal arm is baited with a food reward. During this forced trial the subject is placed in the start arm and forced to visit the open arm. This trial is repeated by randomly varying the closed arm, for an equal number of trials for each arm, until the subject has been familiarized with the task.

Evaluation of Spatial Memory using Rewarded Alternation in the Y-Maze

The choice trial begins by baiting both the arms and allowing the subject to visit only one arm and consume the food reward. This procedure is done by pre-selecting the correct choice and blocking it with a door. Now the unvisited arm becomes the correct choice in the next trial. For the immediate trial both the arms are open and only the correct choice arm is baited. The subject is expected to visit the correct choice arm. If it makes the right choice, it is allowed to consume the food reward, and if it chooses the wrong arm, it is allowed to see that the food well is empty and then it is removed from the maze.

Each trial lasts approximately for 2 minutes. The procedure is repeated for each of the animals on a ten trial per day basis for up to twelve testing days. The correct choice arm is randomly varied throughout the testing sessions.

Spatial Learning using Spontaneous Alternation in the Y-Maze

The Spontaneous Alternation task is based on the novelty of the maze. Thus the task is performed without prior familiarization of the subjects to the maze. The subject is given a free choice, during the trial, to choose either one of the baited arms unlike in the Rewarded Alternation task where one of the arms is blocked. Once the subject has made its choice, it is confined to that goal arm by closing the respective door for 30 seconds. The subject is removed from the goal arm after it has consumed the food reward and the doors in the maze are opened. The subject is then once again placed in the start arm and is expected to alternate its choice from its previous selection. If the subject visits the unvisited arm, it is allowed to consume the food reward or else, in case it visits the already visited arm it is allowed to see that the food well is empty before it is removed from the maze.

Each trial lasts no longer than 2 minutes, and the procedure is repeated for each of the animals on a ten trial per day basis for up to twelve testing days.

Modifications

Similar to the T-maze, Y-Maze has also seen modifications and adaptations to meet the needs of various cognitive neuroscience research to allow studying different aspects of spatial learning and alternate behavior.

A simple modification to the maze can be done using discriminative stimuli such as patterns or objects to which the subject must respond to, in order to obtain a reward. For example, Wright and Conrad (2005) used large enamel-painted rocks as intra-maze cues. Such cues ease the task difficulty and assist the animals in remembering which goal arms they have visited. The subjects can be trained to only choose the arm with the specific rock or cue.

Other variants of the Y-Maze include the water Y-Maze that is a hybrid between the Morris Water Maze and the traditional Y-Maze. Deacon (2013) used a shallow paddling pool Y-Maze as a relatively non-aversive environment for testing mice. The maze included false escape tubes with only one true exit. Another water Y-Maze variant was used by Burguière et al.,2010) to test the role of the cerebellar cortex in conditioned goal-directed behavior. The maze arms were filled with water and used two different conditioned stimuli: a light or a sound. A wrong turn resulted in air puff which acted as an aversive stimulus.

Aquatic models of the Y-Maze also exist for research using fish. Aoki et al. used an automated aquatic Y-Maze that used computer controlled visual cues on an LCD screen placed under the Y-Maze tank. The zebrafish were trained to avoid one arm by pairing the floor color with an electric shock. Aquatic Y-Mazes have also been in studying shoals (Ward et al.,2011). Drosophila Y-Mazes also exist and allow evaluation of chemosensory responses and other behaviors of drosophila.

Multiple Y-Mazes (Botwinick et al.,1963, Ainge et al.,2007) provide a multi-choice complex maze for assessing memory performance and spatial learning. Elevated Y-Mazes are often employed for testing anxiety or stress related performance of animals. Fully automated Y-Mazes are also available which detect the location of the subject within the maze and control the opening and closing of the arm doors as a trigger response. The automated maze is also capable of detecting food rewards.

The Y-Maze can easily be adapted and modified to be used with birds, insects and other animals for assessment of spatial working memory and cognitive abilities. The ease of construction and the simplicity of the apparatus make it one of the popular behavioral assays.

Data Analysis

The data obtained from the Y-Maze is generally very straightforward and consists of the number of correct (the animal enters the opposite arm on the second run) vs. incorrect (the animal enters the same arm previously entered on the previous run) arm entries in each trial. The time it takes the animal to retrieve the food reward can also be recorded. As the animal learns that entering a new arm results in a food reward, the number of incorrect arm entries should decrease. The percentages of correct arm choices can be graphed and compared across a sham control group and a disease model/intervention group.

Using graphs to compare arm entries between different disease or treatment groups allows easy visualization of the effect on spatial memory and learning. Animals in the controls groups should show significant improvements in their correct arm choices. Animals as disease models of neurodegenerative disorders, for example, should show a much slower learning curve with more incorrect choices, even after several trials. Generally, animal cohorts of 10-30 animals are sufficient to obtain p-values of <0.05 using ANOVA (Wright & Conrad 2005, Conrad et al.,2003).

Traslational Research

Fernandez et al. tested the combined effect of leptin and pioglitazone in a mouse model of Alzheimer’s disease. They tested the short-term spatial recognition of the mice on a Y-Maze with black and white visual cue at the end of each arm. After 2-week of acute treatment with combined leptin and pioglitazone, the mice showed a reduction in spatial memory deficits thus suggesting that the combined dosage can be used as a potential treatment for Alzheimer’s disease.

To date, clinical trials have failed to find an effective therapy for victims of traumatic brain injury (TBI) who live with motor, cognitive, and psychiatric complaints.  Pre-clinical investigators are now encouraged to include male and female subjects in all translational research, which is of particular interest in the field of neurotrauma given that circulating female hormones (progesterone and estrogen) have been demonstrated to exert neuroprotective effects. Tucker et al., taking into consideration the sex of the subject, performed Spontaneous Alternation behavior test using visually cued Y-Maze to test the short-term memory of cortically impacted control mice. The results showed that the females’ performance was inferior to its male counterpart in the working memory task.

With advancing technologies, the application of Y-maze in human-based investigation is possible. Using virtual environments in combination with real-world elements, behaviors of humans in the Y-Maze task can be cost-effectively assessed in a safe environment (see also Virtual T-Maze).

Strengths and Limitations

The Y-Maze allows for easily reproducible results due to its simplicity as compared to other behavioral tasks that test spatial learning and memory. Symmetrical Y-Mazes allow for continuous alternation tasks and usually require minimal training and testing time. Unlike the Morris Water Maze Test, there is minimal stress placed on the subject in a conventional Y-Maze. However, the Y-Maze can be adapted for stress and anxiety related task (Water Y-Maze and Elevated Y-Maze). The Y-Maze is a highly adaptable and modifiable task that allows research into different aspects of spatial learning and cognition. The absence of significant stressors and familiarization with the maze prior to testing enable better observations of working memory in the animals as they perform in the maze.

Although simple to construct and easy to use, the Y-Maze has its own limitations. A simple Y-Maze is a single-point two-choice maze thus it has a higher rate of success possibility as the probability of the subject choosing the correct arm is 50% by default. The subjects may also use strategies other than spatial learning based on the spatial and non-spatial cues. Odor trails too may affect the quality of research if olfactory cues are not an intended part of the investigation.

For the task to deliver appropriate results, the subject’s exploratory drive must be maintained throughout. Extensive handling and overtraining of the subjects may place undue stress on them, affecting their performance on the maze. It is also important to maintain minimum variability in the amount of reward used during the training and testing lest the subject should suffer from ‘contrast effects’ wherein its motivation decreases due to receiving less than expected reward (Deacon & Rawlins 2006). In case of Y-Mazes that make use of guillotine doors, it is essential that doors are closed carefully and not dropped too close to the subject to avoid startling it as this experience may be stressful for the subject and cause it to avoid that arm in subsequent trials.

As with all mazes that measure aspects of learning and memory, it is important to remember that many different processes play into the behavior in the maze. In many cases, the Y-Maze is used in conjunction with other mazes to study disease models or transgenic animals and gain a fuller understanding of spatial learning and memory.

Summary

• Y-Maze is a modification of the T-Maze, in which the arms are placed at a natural angle (120 degrees) unlike the sharp 90-degree turn of the T-Maze arms.
• The natural turns make the Y-Maze ideal for Continuous Alternation task.
• The task exploits the innate explorative nature of rodents and subjects them to tasks that require them to alternate between the goal arms to retrieve food rewards.
• Commonly used protocols with the Y-Maze are Forced Alternation task and Spontaneous Alternation task.
• The T-Maze can be easily adapted to investigate the different aspects of spatial learning and for different subjects.
• By using guillotine doors, the Y-Maze can be easily adapted for Delayed Alternation protocol.
• The Y-Maze has been extensively used in the study of hippocampal functions, age-related cognitive decline and anxiety.
• The Y-Maze is also utilized in understanding the effects of drugs and toxins and in understanding underlying pathology of diseases on spatial learning and memory.
• Subjects of the diseased model show a much slower learning curve in comparison to the control group in Y-Maze tasks.

Features

Shape/Material

• Monofilament material
• Silicone-coated
• Smooth passage through tissues
• Biocompatible

Size Options

• Various sizes
• Different experimental needs
• Uniform diameter
• Uniform consistency

Visibility

• Enhanced visibility for
• certain imaging techniques
• Uniform diameter

Research

• Tailored
• For neurological research
• Stroke studies

1. Made of silicone, ideal to improve the reuse probability of suture occlusion
2. The use of nylon thread with better flexibility allows for a higher success rate of modeling
3. The unique technique of open-molded manufacturing provides uniform height and further ensures a stable MCAO model
4. A cylinder-like silicone head ensures that the blood flow decreases immediately after the suture occlusion enters the MCA

Introduction

Middle Cerebral Artery Occlusion (MCAO) is a commonly used focal ischemia model in rodents for stroke research. The endovascular suture or filament technique is popular in vivo model of MCAO that allows reproducible middle cerebral artery (MCA) infractions, does not require craniectomy, and allows reperfusion by the withdrawal of the occluding filament. First described by Koizumi et al. in 1986 and improved upon by Longa et al. in 1989, the method achieves the cessation of blood flow and subsequent brain infarction by forwarding a surgical filament via the internal carotid artery until the tip occludes the origin of MCA. However, the endovascular filament MCAO fails to mimic clinical situations.

On the other hand, the embolic MCAO first described by Kudo et al. is another popular model that closely mimics human ischemic stroke. Thromboembolic clot models and non-thromboembolic (microsphere or macro-sphere induced) stroke models are the two major types of embolic MCAO. Thromboembolic MCAO is achieved by injection of in vitro clots or via in situ clotting by the endovascular installation of thrombin. This model of MCAO closely reflects the pathophysiology of human cardioembolic stroke. The composition of the clot (thromboembolic model) and the size of the spheres (non-thromboembolic model) were used to have a direct effect on the quality of the MCAO and the infraction induced.

Protocol

Pre-surgery procedures

Endovascular filament MCAO

1. Make a midline neck incision between the manubrium and jaw under a dissecting microscope.
2. Bluntly divide the underlying submandibular gland, leaving the left gland in situ, and retract the right gland cranially and secure it along with the sternocleidomastoid muscle.
3. Carefully divide the exposed division of the omohyoid muscle covering the carotid sheath.
4. Separate and isolate the right common carotid artery (CCA) from the vagus nerve and its sheath.
5. Dissect the external carotid artery (ECA) near its bifurcation into lingual and maxillary arteries, cauterize and divide it with micro-scissors.
6. Prepare two loose collar sutures (7-0 silk) around the proximal internal carotid artery (ICA) just above the CCA bifurcation.
7. Temporarily close the CCA using a vessel clip and verify cortical perfusion values.
8. Next, using a vessel clip above the two loose collar sutures, temporarily close ICA to avoid retrograde flow during arteriotomy.
9. Perform arteriotomy in the reflected ECA close to the stump.
10. Introduce the heat blunted silicone coated 7-0 nylon suture via the arteriotomy in the ECA and advance till vessel clip in ICA. Gently tighten the two-loose collar suture around the proximal ICA to avoid reflux blood flow, without traumatizing the arterial wall and then withdrawing the vessel clip.
11. Advance the occluding suture via the internal carotid artery (ICA) towards the cranial base until a mild resistance is observed. Tighten the two collar sutures around the filament.
12. For the transient model of MCAO, gently withdraw the filament after the pre-defined duration. Cauterize the base of ECA and loosen the collar sutures at the proximal ICA to restore carotid flow.
13. Lay the submandibular gland and sternocleidomastoid over the operative field and close the incision using a tissue adhesive or an absorbable suture or a metal staple.

Embolic MCAO

1. Under a dissecting microscope make a 2 cm midline incision. While exposing the surgical field using retractors, dissect the right CCA, ECA, ICA, and pterygopalatine artery (PPA) away from the surrounding nerves and fascia.
2. Place a 5-0 silk suture under the CCA. Tie it into a slip knot and pull taut towards the body.
3. Dissect the ECA and its two branches, the occipital artery (OA) and the superior thyroid artery (STA). Using a veterinary electrosurgical unit, coagulate the two branches.
4. Place two 6-0 silk sutures under the ECA, one towards the head and the other towards the body. Tie the suture towards the head tightly and loosely knot the other suture.
5. Tie the PPA with a 6-0 suture. Using a slip knot with 6-0 suture or microvascular clip, tie/clip the ICA.
6. Between the two sutures of the ECA, cut a small incision in the ECA and insert the modified PE-%) the tube containing the blood clot. Advance the tubing to the bifurcation of the CCA. Tighten the loose suture of the ECA around the lumen, such that to allow mobility of the tube.
7. Free the stump by cutting off the ECA at the incision site. Position the stump below the bifurcation of ECA and the ICA. Open the ICA and advance the tube into it until the tip of the catheter reaches the origin of MCA.
8. Inject the clot through the modified PE-50 catheter along with 10 μl of saline over 10 seconds using a 100 μl Hamilton syringe.
9. Withdraw the catheter from the ECA 5 min later. Tie the ECA and reopen CCA. Suture the incision on the neck.

Apparatus and Equipment

Evaluation of intraarterial administration of norcantharidin

Investigation of the effect of OTULIN on ischemic stroke model

Evaluation of rapamycin in preventing cerebral stroke

Strenght and limitations

Summary

1. Endovascular filament and embolic MCAO are two popular focal ischemic stroke models.
2. Endovascular filament MCAO does not require craniotomy and allows for both permanent and transient occlusions of the MCA.
3. Endovascular filament MCAO does not closely mimic clinical situations. However, it is a relevant model in endovascular therapy.
4. Embolic MCAO has two prominent categories based on the occluding material; thromboembolic clot and non-thromboembolic models.
5. Embolic MCAO closely mimics the human ischemic stroke. However, there is high variability of the MCAO.
6. The thromboembolic clot can be sourced from spontaneously formed or thrombin-induced thrombotic material.
7. Microspheres enable microembolization of multiple vessels.
8. Macrospheres allow delaying of occlusion to induce a stroke.

References

1. Chiang T, Messing RO, Chou W-H. Mouse model of middle cerebral artery occlusion. J Vis Exp. 2011;955(48):9-10. doi:10.3791/2761.
2. Shimamura N, Matchett G, Tsubokawa T, Ohkuma H, Zhang J. Comparison of silicon-coated nylon suture to plain nylon suture in the rat middle cerebral artery occlusion model. J Neurosci Methods. 2006;156(1-2):161-165. doi:10.1016/j.jneumeth.2006.02.017.
3. Chen Y, Ito A, Takai K, Saito N. Blocking pterygopalatine arterial blood flow decreases infarct volume variability in a mouse model of intraluminal suture middle cerebral artery occlusion. J Neurosci Methods. 2008;174(1):18-24. doi:10.1016/j.jneumeth.2008.06.021.
4. Lee S, Hong Y, Park S, Lee S-R, Chang K-T, Hong Y. Comparison of surgical methods of transient middle cerebral artery occlusion between rats and mice. J Vet Med Sci. 2014;76(12):1555-1561. doi:10.1292/jvms.14-0258.

Modifications

Food Wells

Doors/Divider

Light Cues

Backlights

Stand

H Maze

Escape Tubes

Housing

Advancements in cognitive neuroscience research have led to the development of many variations of T-Maze to study particular aspects of spatial learning and alternation behavior.

One of the easiest ways to bring variation in the T-Maze is by using discriminative stimuli such as patterns or objects to which the subject must respond to, in order to obtain a reward. Another simple possible variation would be to train the subjects to select the white painted arm instead of the black painted arm that it would usually prefer instinctively. By training the subject to always choose the white painted arm, regardless of it being the left or the right arm, testing the reference memory of the subject is possible (Deacon and Rawlins 2006).

The use of guillotine doors can also serve as a critical feature in the maze. Using a guillotine door in the start arm to create a confined start area helps assist in Delayed Alternation task and also to prevent the rat from exploring the maze between choice trials (Wenk 2001). Further, the start arm can be modified by dividing it lengthwise with an opaque divider that blocks off one of the two pathways created from the division to investigate the reference memory of the subject (Wenk 2001).

Anxiety related tasks are often carried out using an elevated T-Maze. These tasks usually employ a T-Maze with no walls, forcing the subject into a state of anxiety thus placing more stress on them as they naturally tend to avoid open spaces. A variation of this model used a T-Maze that had the start arm enclosed with lateral walls while the goal arms remained un-walled (Graeff et al.,1998).

Water-based T-Maze models combine the advantages of Morris Water Maze and the T-Maze while minimizing the disadvantages associated with the individual mazes. The Water T-Maze uses a T-shaped tank filled with water with either one of the arms installed with an escape platform or ladder. Locchi et al. used a water escape T-Maze model to assess the sensitivity of spatial memory in response to pharmacological manipulations and suggested that the Water T-Maze served as a valid method of investigation. On the other hand, Guariglia et al. used a Water T-Maze as a simple and inexpensive task for assessing repetitive behaviors in mice which other conventional methods, such as the Morris Water Maze, could not conclusively demonstrate.

The multiple T-Maze makes use of many simple T-Mazes joined together to create a complex, multi-choice maze with identical choice points to determine place vs. response learning and cognitive maps. This complex maze was used by Tolman and Honzink (1930) to put into perspective their theory that rats actively process information rather than operating on a stimulus-response relationship.

Aquatic T-Mazes have also been used in research of effects of toxins on memory and learning related tasks and cognitive flexibility in fish (Zheng et al.,2017, Byrnes et al.,2017, Miletto et al.,2017). Vertical T-Mazes have also been used as an adapted version of the classical T-Maze in studies of arthropods (Stelinski and Tiwari 2013). This adaptation takes advantage of the natural tendency of many arthropods towards positive phototaxis and negative geotaxis.

Another version of the T-Maze is the continuous angled T-Maze that has often been used in Grid cell experiments. The goal arms of the continuous maze angle back to the start. This modification limits the interference of the experimenter in moving the subject from the goal arms to the start position. The Two Problem T-Maze is yet another adaptation of the T-Maze. This maze is usually used in the examination of the effects of prelimbic lesions in the rat prefrontal cortex to determine how it affects working memory through assessing the acquisition and retention of nonmatching to sample (NMTS) and matching to sample tasks (MTS).

Many species-based adaptations of the T-Maze exists to serve as a tool for assessment of spatial working memory and cognitive abilities. The ease of construction and the simplicity of the apparatus make it one of the popular behavioral assays.

Variants

Continuous Angled T Maze

Two Problem T Maze

Elevated T Maze

Y/T Maze

Light Dark T Maze

Split T Maze

Rodent Trident Maze

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Introduction

The T-Maze has been widely used in neuroscience for studies of spatial learning and memory. The task is based on the explorative nature of rodents to locate food quickly and efficiently. The basic model of the maze provides the subject with two options: left arm and right arm. The natural tendency of the rodent would be to explore the un-visited arm after retrieving food reward from one of the arms. Typically left-right discrimination and forced alternation tasks are used to assess reference and working memory using the T-Maze. Alternation task, both rewarded and spontaneous, has been shown to be effective in detecting hippocampal dysfunction and lesions.

In the late 1950s, Dember and Fowler were the first to study alternation behavior in animals and published papers for the same. It has since been widely used to measure spatial memory and learning. The T-Maze serves as a simple test to assess hippocampal learning and detection of cognitive dysfunction, and due to its simple construction and usability, it has been used extensively to study treatments and toxins that affect spatial memory and age-related cognitive decline (Sharma et al.,2010). While rewarded alternation requires food rewards and takes fewer trials, spontaneous alternation does not require food rationing and can be interleaved with other tests.

The basic T-Maze consists of just two goal arms and a base arm forming a ‘T.’ The simplest modification to the T-Maze can be done using guillotine doors that can be automated or manually operated. Other versions of the T-Maze include the Y-Maze wherein the arms form a ‘Y’, the continuous angled T-Maze that has the choice arms angled and return to the base arm, H Maze conversion of the maze to minimize experimenter grasping of the subjects in recurrent trials, and multiple T-Maze as used by Tolman and Honzik in 1930 to investigate latent learning. The maze can also be adapted for research in aquatic environments.

History

In 1946, Tolman utilized a combination T-Maze to investigate place leaning and reference learning in rats. He hypothesized that animal learning included purpose and cognition (Tolman et al.,1925). His experiment suggested that the learned behavior in the T-Maze was a disposition to orient or to go towards the location of the goal.

Kendler in 1947, conducted an investigation of latent learning in a simple T-Maze as an improvement on the experiment carried out by Spence and Lippitt (1946). The experiment also examined the adequacy of Tolman’s non-reinforcement learning theory (Tolman, 1932). Kendler’s experiment concluded that motivational satisfaction indeed played a major role in learning as observed in the choices made by the subjects in the trials, which were in contrast to the expected results deduced from Tolman’s theory.

The late 1950s saw Dember and Fowler investigate alternation behavior in animals in a two-choice situation. Their published papers (Dember and Fowler 1959, Fowler et al.,1959a, Fowler et al.,1959b) described spontaneous versus forced alternation behavior and the influence of rewards and reinforcements.

Since the research conducted by Dember and Fowler, the T-Maze has been developed as a relatively simple and widely used task in measuring spatial memory and its many variations.

In 1979, Thomas carried out experiments using a T-Maze to study the effect of small posterodorsal septum lesions on alternation behavior of rats. The trials compared the performance of spontaneous and rerun correction alternation in rats with small posterodorsal septal lesions and found that the lesions not just reduce alternation to the randomness of choice but, plausibly, turn the subject (on average) from alternators to preservators.

The effects of fornix fimbria lesions, medial septal lesions, and lateral septal lesions were tested with rats trained on a rewarded alternation task, run as a spatial working memory task on an elevated T-Maze by Rawlins and Olton in 1982. They concluded that rats have difficulty in using information about ‘places,’ and that control and lesion rats learn the tasks in the same way.

A 2002 study, by Reisal et al. on spatial memory dissociations used gene-targeted mice lacking the α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptor subunit GluR1 (GluR-A). In comparison to the Morris Water Maze task, the T-Maze alternation task was successful in detecting deletion of the GluR1.

Due to its ability to readily test the function of hippocampal learning, the T-Maze has been used extensively to study treatments that affect spatial memory and age-related cognitive decline (Sharma et al.,2010).

The T-Maze has also been adapted for research in aquatic life; Aquatic T-Maze. Zheng et al. analyzed the effect of polybrominated diphenyl ethers (PBDEs) on the learning and memory abilities of male zebrafish. Another study assessed T-Maze turn preferences in the Port Jackson sharks to examine laterality during exploration of a novel habitat (Byrnes et al.,2017). Miletto et al., 2017) used male and female guppy fish in the aquatic T-Maze model to investigate the difference in cognitive flexibility of the sexes in a discrimination reversal learning task and concluded that although the meta-analysis supported the hypothesis of females having greater reversal learning ability this difference could, however, be dependent on the task that they are subjected to.

Ichinose and Tanimoto (2016) further developed and adapted T-Maze for Drosophila into a multiplex T-Maze to study the dynamics of memory-guided choice behavior. This new system allowed them to demonstrate that serotonin neurons control the speed and the eventual result of the conditioned odor approach.

In 1946, Tolman utilized a combination T-Maze to investigate place leaning and reference learning in rats. He hypothesized that animal learning included purpose and cognition (Tolman et al.,1925). His experiment suggested that the learned behavior in the T-Maze was a disposition to orient or to go towards the location of the goal.

Kendler in 1947, conducted an investigation of latent learning in a simple T-Maze as an improvement on the experiment carried out by Spence and Lippitt (1946). The experiment also examined the adequacy of Tolman’s non-reinforcement learning theory (Tolman, 1932). Kendler’s experiment concluded that motivational satisfaction indeed played a major role in learning as observed in the choices made by the subjects in the trials, which were in contrast to the expected results deduced from Tolman’s theory.

The late 1950s saw Dember and Fowler investigate alternation behavior in animals in a two-choice situation. Their published papers (Dember and Fowler 1959, Fowler et al.,1959a, Fowler et al.,1959b) described spontaneous versus forced alternation behavior and the influence of rewards and reinforcements.

Since the research conducted by Dember and Fowler, the T-Maze has been developed as a relatively simple and widely used task in measuring spatial memory and its many variations.

In 1979, Thomas carried out experiments using a T-Maze to study the effect of small posterodorsal septum lesions on alternation behavior of rats. The trials compared the performance of spontaneous and rerun correction alternation in rats with small posterodorsal septal lesions and found that the lesions not just reduce alternation to the randomness of choice but, plausibly, turn the subject (on average) from alternators to preservators.

The effects of fornix fimbria lesions, medial septal lesions, and lateral septal lesions were tested with rats trained on a rewarded alternation task, run as a spatial working memory task on an elevated T-Maze by Rawlins and Olton in 1982. They concluded that rats have difficulty in using information about ‘places,’ and that control and lesion rats learn the tasks in the same way.

A 2002 study, by Reisal et al. on spatial memory dissociations used gene-targeted mice lacking the α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptor subunit GluR1 (GluR-A). In comparison to the Morris Water Maze task, the T-Maze alternation task was successful in detecting deletion of the GluR1.

Due to its ability to readily test the function of hippocampal learning, the T-Maze has been used extensively to study treatments that affect spatial memory and age-related cognitive decline (Sharma et al.,2010).

The T-Maze has also been adapted for research in aquatic life; Aquatic T-Maze. Zheng et al. analyzed the effect of polybrominated diphenyl ethers (PBDEs) on the learning and memory abilities of male zebrafish. Another study assessed T-Maze turn preferences in the Port Jackson sharks to examine laterality during exploration of a novel habitat (Byrnes et al.,2017). Miletto et al., 2017) used male and female guppy fish in the aquatic T-Maze model to investigate the difference in cognitive flexibility of the sexes in a discrimination reversal learning task and concluded that although the meta-analysis supported the hypothesis of females having greater reversal learning ability this difference could, however, be dependent on the task that they are subjected to.

Ichinose and Tanimoto (2016) further developed and adapted T-Maze for Drosophila into a multiplex T-Maze to study the dynamics of memory-guided choice behavior. This new system allowed them to demonstrate that serotonin neurons control the speed and the eventual result of the conditioned odor approach.

Apparatus & Equipment

The apparatus is a capital ‘T’ shaped maze with arm lengths ranging from 30 to 50 cm having a width of approximately 10 cm to accommodate mice, rats, and small primates. Generally, the goal arms of the maze are equipped with guillotine doors that can be automated or manually managed to confine the subject within the arm or to block off one of the arms. The goal arms may have food wells or have goal boxes attached to them to hold rewards or stimuli. The maze is usually used as an enclosed maze with wall heights of 30 cm, and the apparatus is typically raised to a height of at least 50 cm above the floor.

The apparatus is usually painted a dark color to avoid unnecessary anxiety in the subjects. The T-Maze construction is usually adapted depending on the subject used for example the transparent miniature T-Maze used in studies of memory and cognition in Drosophila (Lin et al., 2017, Ichinose & Tanimoto 2016) and fish tank model used for experiments involving fish (Zheng et al.,2017, Miletto et al.,2017).

Automated T-Maze can detect the location of the subject within the maze and control the opening and closing of the arm doors as a trigger response. The automated maze is also capable of detecting food rewards.

To avoid shadows in the maze, the T-Maze should be well lit from above. Proper lighting also ensures that the subject is able to see the food rewards. Tracking software and video camera, such as Noldus Ethovision XT, mounted above the maze assist with live scoring and tracking and recording the subject and its movements within the maze. The apparatus must be cleaned thoroughly before and after each trial to limit influence from any residual stimuli from previous trials.

Training Protocol

The T-Maze task involves observation of the ability of the subject to remember the previously visited arm. The task takes advantage of the natural explorative nature of rodents and the idea that they tend to use an optimal search strategy to obtain food with minimal effort.

The performance in the task weakens or decreases for subjects with hippocampal lesions or damage as seen in neurodegenerative or neuropsychiatric diseases or as a result of natural aging.

Several protocols exist to be used with the T-Maze, though the two most commonly used protocols are the Rewarded Alternation task and Spontaneous Alternation task. For the Rewarded alternation task, the experimenter decides which arm is the “correct choice” whereas the Spontaneous Alternation task uses the subject’s natural explorative drive and allows it to choose which arm to explore first. Spontaneous alternation task has been shown to be more successful in the assessment of subjects with hippocampal lesions as these animals tend to develop a side preference (Deacon and Rawlins 2006). The task is also used to quantify cognitive deficits in transgenic strains of mice and evaluate effects of treatments on cognition.

Another protocol used with the T-Maze is the Delayed Alternation task. The Delayed Alternation task allows assessing spatial working memory by first allowing the subject to explore a baited arm of its choice and removing the subject once the choice is made and limiting it to the start box of the base arm using a door. This trial is followed by a formal trial, after a predetermined delay between the two trials, wherein the subject is reintroduced to the maze by opening the start arm door and is expected to choose the arm that it did not choose in the previous trial.

The T-Maze can be modified to allow disassociation between working and reference memory. This protocol is accomplished by changing external cues and splitting the stem arm lengthwise using an opaque divider. The end of one of the paths created by the divider is blocked using a clear Plexiglas. If the subject chooses the unblocked path, the protocol is the same as that of an Alternation task. (Wenk 2001)

Rodents tend to be wary of eating anything new. Thus it is important to familiarize the subject with the food rewards used, before the testing, within a familiar environment (housing cage). Food rewards can be solid (such as sweetened breakfast cereal) or liquid (such as chocolate milk.) Generally, liquid rewards are used in case of testing as some treatments may make eating solid food an unpleasant experience for the subject. Subjects are usually maintained at about 85 to 95% of their free-feeding weight throughout the training and testing phases.

For the food-deprived task, the food-scavenging behavior is encouraged by depriving the subject of food the night prior to the testing. Subjects may be familiarized to the maze in pairs (usually cage-mates) to reduce the anxiety in a novel space during the initial phase of exploration. The acclimation is usually done individually, although, this may require more time than paired familiarization. The maze is scattered with food rewards, and the subject is placed in the start arm and allowed to explore the maze freely. The food rewards are replaced on consumption. The familiarization procedure is repeated four times with at least ten minutes between each exposure. The acclimation period usually lasts 1 to 2 days. Next, one of the arms is blocked, and the open goal arm is baited with a food reward. During this forced trial the subject is placed in the start arm and forced to visit the open arm. This trial is repeated by randomly varying the closed arm, for an equal number of trials for each arm, until the subject has been familiarized with the task.

The choice trial begins by baiting both the arms and allowing the subject to visit only one arm and consume the food reward. This procedure is done by pre-selecting the correct choice and blocking it with a door. Now the unvisited arm becomes the correct choice in the next trial. For the immediate trial both the arms are open and only the correct choice arm is baited. The subject is expected to visit the correct choice arm. If it makes the right choice, it is allowed to consume the food reward, and if it chooses the wrong arm, it is allowed to see that the food well is empty and then it is removed from the maze.

Each trial lasts approximately for 2 minutes. The procedure is repeated for each of the animals on a ten trial per day basis for up to twelve testing days. The “correct choice” arm is randomly varied throughout the testing sessions.

The Spontaneous Alternation task is based on the novelty of the maze. Thus the task is performed without prior familiarization of the subjects to the maze. The subject is given a free choice, during the trial, to choose either one of the baited arms unlike in the Rewarded Alternation task where one of the arms is blocked. Once the subject has made its choice, it is confined to that goal arm by closing the respective door for 30 seconds. The subject is removed from the goal arm after it has consumed the food reward and the doors in the maze are opened. The subject is then once again placed in the start arm and is expected to alternate its choice from its previous selection. If the subject visits the unvisited arm, it is allowed to consume the food reward or else, in case it visits the already visited arm it is allowed to see that the food well is empty before it is removed from the maze.

Each trial lasts no longer than 2 minutes, and the procedure is repeated for each of the animals on a ten trial per day basis for up to twelve testing days.

Rodents tend to be wary of eating anything new. Thus it is important to familiarize the subject with the food rewards used, before the testing, within a familiar environment (housing cage). Food rewards can be solid (such as sweetened breakfast cereal) or liquid (such as chocolate milk.) Generally, liquid rewards are used in case of testing as some treatments may make eating solid food an unpleasant experience for the subject. Subjects are usually maintained at about 85 to 95% of their free-feeding weight throughout the training and testing phases.

For the food-deprived task, the food-scavenging behavior is encouraged by depriving the subject of food the night prior to the testing. Subjects may be familiarized to the maze in pairs (usually cage-mates) to reduce the anxiety in a novel space during the initial phase of exploration. The acclimation is usually done individually, although, this may require more time than paired familiarization. The maze is scattered with food rewards, and the subject is placed in the start arm and allowed to explore the maze freely. The food rewards are replaced on consumption. The familiarization procedure is repeated four times with at least ten minutes between each exposure. The acclimation period usually lasts 1 to 2 days. Next, one of the arms is blocked, and the open goal arm is baited with a food reward. During this forced trial the subject is placed in the start arm and forced to visit the open arm. This trial is repeated by randomly varying the closed arm, for an equal number of trials for each arm, until the subject has been familiarized with the task.

The choice trial begins by baiting both the arms and allowing the subject to visit only one arm and consume the food reward. This procedure is done by pre-selecting the correct choice and blocking it with a door. Now the unvisited arm becomes the correct choice in the next trial. For the immediate trial both the arms are open and only the correct choice arm is baited. The subject is expected to visit the correct choice arm. If it makes the right choice, it is allowed to consume the food reward, and if it chooses the wrong arm, it is allowed to see that the food well is empty and then it is removed from the maze.

Each trial lasts approximately for 2 minutes. The procedure is repeated for each of the animals on a ten trial per day basis for up to twelve testing days. The “correct choice” arm is randomly varied throughout the testing sessions.

The Spontaneous Alternation task is based on the novelty of the maze. Thus the task is performed without prior familiarization of the subjects to the maze. The subject is given a free choice, during the trial, to choose either one of the baited arms unlike in the Rewarded Alternation task where one of the arms is blocked. Once the subject has made its choice, it is confined to that goal arm by closing the respective door for 30 seconds. The subject is removed from the goal arm after it has consumed the food reward and the doors in the maze are opened. The subject is then once again placed in the start arm and is expected to alternate its choice from its previous selection. If the subject visits the unvisited arm, it is allowed to consume the food reward or else, in case it visits the already visited arm it is allowed to see that the food well is empty before it is removed from the maze.

Each trial lasts no longer than 2 minutes, and the procedure is repeated for each of the animals on a ten trial per day basis for up to twelve testing days.

Data Analysis

The data obtained from the T-Maze is generally very straightforward and consists of the number of correct (the subject enters the opposite arm on the second run) vs. incorrect (the animal enters the same arm previously entered on the previous run) arm entries in each trial. Working memory and reference memory data can be obtained from the number of entries into the blocked side of the stem and number of re-entries into the un-baited arm (Wenk 2001). Data can also be collected for the latencies to explore and retrieval of the rewards, time spent in the goal arms and so on.

As the subject learns that entering into a new arm results in food reward the number of incorrect entries should decrease. Subjects with hippocampal lesions or damage are likely to make errors. The percentages of correct arm choices can be graphed and compared across a sham control group and a disease model/intervention group. Graphs allow easy visualization of effects on spatial memory and learning between different disease or treatment groups. Animals as disease models of neurodegenerative disorders, for example, should show a much slower learning curve with more incorrect choices, even after several trials. Generally, animal cohorts of 10-30 animals are sufficient to obtain p-values of <0.05 using ANOVA (Jang et al.,2013, Wang et al.,2014).

Translational Research

Transgenic mice have been used in animal models of human diseases to understand and to develop treatments that are effective. Vivithanaporn et al. used HIV-1 Vpr transgenic mice to investigate the neurotoxic actions of the HIV protease inhibitors, amprenavir (APV) and lopinavir (LPV).  The mice were divided into groups that were treated with fosAPV/ritonavir and LPV/ritonavir, and their spatial memory was tested in a simple T-maze under a food reward alteration protocol. The data obtained from the T-Maze showed that mice treated with APV took significantly more time to complete the maze in comparison to the control and the LPV group. The number of errors was comparatively higher in both APV and LPV group than in the control group. The researchers concluded based on the observed data that that protease inhibitor treatment impaired learning and memory.

A change in diagnostic practices has led to the improved identification of Autism Spectrum Disorder (ASD). ASD is characterized by alternation behaviors in social interactions, verbal and non-verbal communications and repetitive behaviors. Spontaneous Alternation task using T-Maze is used in the evaluation of repetitive behaviors that are usually seen in Autism and OCD models of a mouse. Chang et al. used the T-Maze along with the Marble Burying task in assessing the repetitive behavior symptom domain of ASD. Depending on the severity of the phenotypic behavior, the percent alternation varied from 60% to 10%.

Leqin et al. generated NRG2 knockout mice due to lack of investigation into the function of this closest Neuregulin-1 homolog. NRG2 knockout mice showed dopamine dysbalance as seen in schizophrenic brains. T-Maze Rewarded Alternation task was used to assess the working memory of NRG2 knockout mice. The results showed that with acute administration of Clozapine the performance of the subject improved in the rewarded task which was speculated to be either by indirectly augmenting D1R signaling through the increased dopamine levels, by directly blocking the excessive activity of D4Rs or by simultaneously activating of D1- and D2-type receptors to optimize temporal gating.

Corticotrophin-releasing factor (CRF) gene’s single nucleotide polymorphism has been associated with behavioral inhibition which is a childhood risk factor for panic disorder. Further studies have also shown that single nucleotide polymorphism in type 1 and type 2 receptors of CRF have correlations with major depression and panic disorders. Silva et al. used an elevated T-Maze to study the role of CRF receptors in the escape response of rats. The investigation revealed that ventromedial hypothalamus CRF type 2 receptor show a defensive response associated with panic disorder thus playing a crucial role in the understanding of the underlying neural mechanisms.

Virtual T-Maze allows direct evaluation of human behaviors. Using virtual reality tests can be performed cost-effectively and in a safe environment. Some virtual T-Mazes use a combination of real-world elements such as a T-Maze platform along with virtual elements.

Strengths and Limitations

In comparison to other tasks used in assessing spatial learning and memory, the T-Maze is a simple apparatus that is easy to construct and use. Simple modifications such as using guillotine doors on the arms allow the task to be adapted for Delayed Alternation task while joining together multiple T-Mazes creates a complicated version of the rather simple T-Maze. Generally, an enclosed T-Maze is used which places minimal stress on the subjects, although elevated and un-walled T-Mazes are also used especially in studies related to anxiety (Graeff et al.,1998). Researchers have also used a water T-Maze which combines the advantages of the simple T-Maze and the Morris Water Maze (Locchi et al.,2007, Guariglia et al.,2013).

The absence of significant stressors in the simple T-Maze coupled with familiarization with the maze prior to testing allow for better observations of working memory in the animals as they perform in the maze.

Despite the simplicity, the T-Maze has its own limitations. A simple T-Maze is a single-point two-choice maze thus it has a higher rate of success possibility as the probability of the subject choosing the correct arm is by default 50%. The subjects may also use strategies other than spatial learning based on the spatial and non-spatial cues. Odor trails too may affect the quality of research if olfactory cues are not an intended part of the investigation.

For the task to deliver appropriate results, the subject’s exploratory drive must be maintained throughout. Extensive handling and overtraining of the subjects may place undue stress on them, affecting their performance on the maze. It is also important to maintain minimum variability in the amount of reward used during the training and testing lest the subject should suffer from ‘contrast effects’ wherein its motivation decreases due to receiving less than expected reward (Deacon & Rawlins 2006). In case of T-Mazes that make use of guillotine doors, it is essential that doors are closed carefully and not dropped too close to the subject to avoid startling it as this experience may be stressful for the subject and cause it to avoid that arm in subsequent trials.

As with all mazes that measure aspects of learning and memory, it is important to remember that many different processes may come into play to form behavior. In many cases, the T-Maze is used in conjunction with other mazes to study disease models or transgenic animals and gain a fuller understanding of spatial learning and memory.

Summary

• The T-Maze is a simple single-point two-choice maze.
• The task exploits the innate explorative nature of rodents and subjects them to tasks that require them to alternate between the goal arms to retrieve food rewards.
• Commonly used protocols with the T-maze are Forced Alternation task and Spontaneous Alternation task.
• The T-Maze can be easily adapted to investigate the different aspects of spatial learning and for different subjects.
• By using guillotine doors, the T-Maze can be easily adapted for Delayed Alternation protocol.
• The T-Maze has been extensively used in the study of hippocampal functions, age-related cognitive decline, and anxiety.
• The T-Maze is also utilized in understanding the effects of treatments in understanding underlying pathology of diseases on spatial learning and memory.
• Subjects of the diseased model show a much slower learning curve in comparison to the control group in T-Maze tasks.

References

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• Removable Top.
• Black Nesting Chamber w clear nest holder included
• Matted finish

Specifications

Barnes Maze is available with the following characteristics:

• Includes 1 target box with the option to add more holes
• Easy to clean – removable escape box holder
• Entire top and dark escape box rotate for customization
• False floor modification available for added versatility
• Thick acrylic top eliminates visual cues
• Available in white, grey, clear, or blue

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Characteristics

• Six Stirrings
• Digital RPM Indicator
• Water Level Indicator
• Cell Holders: Diffusion Cells + Stirring Bars
• Stainless-steel body of cell apparatus

• Water Heater
• Speed Controler
• Six Transdermal Cups
• Constant Stirring of Solutions
• Digital Temperature Controller

Extra Nest box

Barnes Maze False Floor

Optogenetics Modification

Optogenetics Modification Type 2

$100

An extra nest box for easy switch and cleaning, minimizing delay between experiments

Add on Mouse: $650 / Rat: $850

Used to prevent falling into non-target holes. Rotates underneath the Barnes Maze.

$200

Optogenetics modification gives half holes and a step wise shortened target box to minimize teather interaction

$350
Gives modified target holes of 3/4 size as well as a gradient entrance to the target box. This box allows for more gradual entry for rodents with optogenetics tethers.

Target Floor Insert

Randomized Hole Pattern

Patterned Holes

Radial Arm Barnes MAze

$650

False floor blocks all the holes except one. The black rectangular insert is to block the target box, leaving no holes open.

For Mouse and Rat

Leads to a decrease in the serial strategy of goal hole choice.

Delayed Matching To Place (DMP) Barnes Maze

Combined System 

Combines the best elements of the Radial arm maze with the fear aversion motivation of the Barnes maze.

Cheeseboard Style

Juvenile Barnes Maze

Utilizes this ability to assess the working and reference memory of the rodents 

Assessed maturation of spatial learning and memory, emphasizing the emergence of spatial navigation skills by the end of the third postnatal week.

Introduction

Overview

Barnes Maze (BM) is a behavioral task often used in neuroscience for the study of spatial learning and memory. The task’s primary ability is to measure the capacity of the subject to learn the location of the target by using distal visual cues. The maze exploits the averseness that rodents feel towards open and brightly lit spaces to motivate them to find the target location. The Barnes Maze requires the use of hippocampal-dependent spatial reference memory to be able to locate the escape locations. This ability to remember the location of the target hole can be affected by the administration of certain drugs or disease models.

The Barnes Maze was designed by Carol Barnes in 1979 to evaluate spatial learning and memory. Initially intended for rats, the Barnes Maze has been increasingly adapted to be used with mice as well (Sunyer et al., 2007). The BM task draws similarities to the Morris Water Maze (MWM) and the Radial Arm Maze (RAM) task; however, unlike the aforementioned mazes, the Barnes Maze does not expose the subjects to strong aversive stimuli such as forced swimming and food/water deprivation and in comparison can be considered to be a low-stress alternative to these tasks (Harrison et al., 2006). The maze, since its conception, has been used not just for spatial learning and memory but also in testing and validating the effects of drugs and pharmacological compounds in models of diseases like Alzheimer’s (Harrison et al., 2006, Attar et al., 2013), and in understanding learning and memory deficits associated with mild traumatic brain injuries.

The Barnes Maze apparatus is a simple circular platform with circular holes serially placed along the edge of the platform. The task of the subject is to find the hole that serves as the target location. The target location leads to a small and dark recessed chamber beneath the platform and is not visible to the subject from the platform. Intra- and extra-maze cues are often used to assist the subject in finding the target hole.

Variants of the Barnes Maze include the Delayed Matching to Place Barnes Maze, Randomized Barnes, and  Radial Arm Barnes Maze.

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History

Origin

The BM was developed by Carol Barnes and described in her paper investigating memory deficits associated with senescence (Barnes 1979).

The effects of 3,4-diaminopyridine in the age-related improvement of short-term spatial memory was investigated using the BM, the results of which suggested that the 3,4-DAP selectively improved memory performance of the old subjects, and, within that age group, only improved performance on the short-term memory task (Barnes et al.,1989).

Markowska et al., 1989 utilized the BM for spatial memory and reversal tasks in their investigation to determine the correlations among different behavioral and neurobiological measures in aged rats.

Since its initial use, the BM has seen a slow but steady growth in use over the years in investigations related to neurodegenerative diseases such as Alzheimer’s and in understanding the effects of brain lesions.

Developments

Vorhees (1997) in his paper investigated the effects of prenatal exposure to neurotoxins. For his investigation, he used different behavioral assays to assess the different types of learning and memory, one of these tests being the BM task to assess spatial learning. These tasks were used to detect long-term CNS dysfunction after prenatal exposure.

Adult Lhx5-deficient mice were used in the investigation of learning impairments and motor dysfunctions by Paylor et al.,2001. The hippocampus plays a crucial role in memory and learning, and its absence or disorganized neuroanatomy as observed in the Lhx5 mutated mice reflects poor performance in the BM spatial learning task.

The potential of voluntary running in aiding cognitive brain and cognitive functions after Whole-brain irradiation was assessed by Wong-Goodrich et al.,2010. When the subjects were assessed in BM task after daily running following WBI, it was observed that running significantly prevented spatial memory retention decline observed months after irradiation. It was concluded based on their observations that exercise assisted in the recovery of hippocampal plasticity and could be used as a potential therapeutic intervention.

Recent Developments

Meyer et al.,2014 tested their hypotheses that neonatal leptin would prevent the development of Growth Restricted (GR) associated behavioral abnormalities. In the BM task, the baseline escape times were faster for GR mice; however, the GR mice exhibited regression in their escape times on days 2 and 3. They concluded that alternation in social interactions, learning and activity of mice due to GR could be mitigated by supplementation with the neurotrophic hormone leptin.

The risks of space radiation to astronauts and Alzheimer’s disease-related pathology were evaluated by Rudobeck et al.,2017. APP/PSEN1 transgenic mice and wild-type mice were irradiated with protons, and their performance was tested on the BM at 3 and 6 months after irradiation to evaluate spatial learning and memory.

Apparatus & Equipment

The Barnes Maze apparatus is composed of a circular platform that is raised above the floor to a height of approximately 100 cm. Holes of sizes ranging from 50 to 100 cm in diameter to accommodate mice, rats, and small primates are arranged serially and are equally spaced along the perimeter of the platform.

The holes are placed several centimeters from the edge of the maze, and the number of holes can vary from anywhere between 18 to 20 or more. The designated target hole has a recessed small, dark chamber under the platform. This hole serves as a safe space for the subject and is not visible from the platform to the subject. The entire platform is rotatable around the center point allowing the location of the escape chamber to be easily varied.

The platform of the maze is colored in contrast to the color of the animals being tested. For ease of observation, the holes of the maze are numbered starting from the target hole; the holes to the right of the target hole are numbered starting with +1, and those to the left are numbered starting with -1, with an opposite hole directly across from the target hole, labeled zero.

The visual cues used for the task can be either intra- or extra-maze cues and can be prepared using different colors and shapes. The cues serve as a reference point for the subject during the task.

To avoid shadows in the maze, the BM should be well-lit from above. Proper lighting also ensures that the subject can see the rewards or other cues. Tracking software and video camera, such as Noldus Ethovision XT or ANY-Maze, mounted above the maze can assist with live scoring and tracking, and recording the subject and its movements within the maze. The apparatus must be cleaned thoroughly before and after each trial to limit the influence of any residual stimuli from the previous trials.

Protocol

The Barnes Maze is a simple task used in measuring spatial learning and memory in rodents and small primates. The task measures these parameters by observing the ability of the subject to remember the location of the target hole leading to an enclosed escape chamber.

This test can provide information regarding hippocampal-dependent learning, specifically spatial memory. The BM task has been utilized to understand the effects of age and neurodegenerative diseases on the learning and memory capabilities of the subject. Typically, animals are capable of learning and remembering the location of the target hole using intra- and extra-maze cues.

Before every trial, the apparatus must be thoroughly cleaned to avoid the influence of residual stimuli, if any, from influencing the performance of the subjects.

Pre-Training for the Barnes Maze

The apparatus is set up, and the visual cues are placed in their respective locations. The cues remain constant throughout all training and testing trials. The subject is (usually) placed in a cylindrical dark start chamber in the middle of the circular platform and released after 10 seconds have elapsed.

The subject is gently guided towards the escape hole avoiding any force to prevent unnecessary stress on the animal. The subject is allowed to remain in the escape chamber for 2 minutes.

Evaluation of Spatial Memory Using the Barnes Maze

The apparatus is cleaned to remove residual olfactory cues from previous runs, and the platform is rotated on its central axis to control any remaining olfactory cues. The escape chamber is adjusted so that it is in the same position.

The video recording is started, and the subject is released from a cylindrical chamber from the center of the platform after 10 seconds. The trial lasts about 3 minutes, during which the subject freely explores the platform. Errors are recorded every time the subject pokes its head into a hole that is not the target hole, and the latency time is determined as the time the subject takes to reach the target hole. The trial ends when the subject has entered the escape chamber, or the 3 minutes have elapsed. In the event, the subject fails to find the escape chamber it is gently guided to it and allowed to remain in it for 1 minute. The subject is returned to its home cage until the next trial.

Each animal should perform four trials on each of the four testing days with approximately fifteen minutes inter-trial intervals.

Evaluation of Reference Memory Using the Barnes Maze

On the fifth testing day, the target hole is closed, or the escape chamber is removed. The trial is initiated as mentioned earlier and lasts for 90 seconds. The number of errors and the latency time are recorded. The subject is removed from the maze when the 90 seconds have elapsed.

The procedure is repeated 7 days later, on the twelfth day.

Modifications

Since its original design, the Barnes Maze has been adapted with several simple modifications. For example, a curtain surrounding the maze platform can be used to prevent the animals from making spatial associations between distal room cues and the location of the target hole (Harrison et al., 2006, Rosenfeld & Ferguson  2014). Protocol variations have also been made to increase task difficulty (Attar et al., 2013).

The escape chamber beneath the maze can optionally lead to an escape tube that allows the animal to reach a home cage or other safe space (Rosenfeld & Ferguson, 2014). A false floor can also be added beneath the maze platform to close off the holes that do not lead to the escape chamber.

The hole positioning has also seen modifications over the years to improve the spatial learning and memory measure of the Barnes Maze task. The Delayed Matching to Place Barnes Maze (DMP Barnes Maze), is a dry variant of the DMP water maze by Steele and Morris (Steele and Morris 1999) that was refined by Faizi et al. 2012. The DMP maze is a Patterned Barnes Maze (PBM) that has the escape platform frequently changed during trials. The apparatus includes an elevated circular platform having 40 holes arranged across the inner, middle and outer rings. Each of these holes is attached to an ABS tube of which only one tube acts as an escape tube.

Another variation of the Barnes Maze is the Randomized Barnes Maze which was designed to overcome the limitation of the BM. In Barnes Maze, the holes are arranged serially along the circumference of the platform that can be serially searched by the subject rather than using a spatial strategy. The Randomized Barnes Maze is modified such that the holes are placed in a pseudorandom order to discourage non-spatial strategies.

A combination of the classic Radial Arm Maze and Barnes Maze, the Radial Arm Barnes Maze combines the advantages of both mazes into one. The maze was first described by Paganelli’s et al. in their 2004 paper investigating the influence of ischemic brain damage on the acquisition and retention of cognition in mice.

Data Analysis

The data obtained from the Barnes Maze generally consists of two main measures: the number of error head pokes the animal makes, and the time it takes the animal to enter the target hole and the escape chamber. Other measures such as the total path length and movement speed can also be measured and obtained from video tracking software.

As the animal learns the relationship between local or distal spatial cues and the location of the target hole, the number of error head pokes and the latency time should decrease. These measures can be simply graphed and compared across a sham control group and a disease model.

The search strategy used by the animal must be analyzed manually using the video recording and tracking software. Generally, one of three search strategies is used by the animal to locate the target hole:

• Direct: The animal moves directly to the target hole or an immediately adjacent hole before entering the target hole
• Mixed: The animal searches random holes, crossing through the center of the maze
• Serial: The animal visits several adjacent holes in a clockwise or counter-clockwise manner before reaching the target hole

The exact position of each head poke error can also be counted and graphed to help visualize these strategies.

Using graphs to compare latency time, the number of error head pokes, the position of these errors and the total path length between different disease or treatment groups the effect on spatial memory and learning can be easily visualized.

Animals in the control groups should show significant improvements in reaching the target hole quickly and efficiently. Animals as disease models of neurodegenerative disorders, for example, should show a much slower learning curve with more errors and longer path lengths, even after several days. Generally, animal cohorts of 10-30 animals are sufficient to obtain p-values of <0.05 using ANOVA and step-down Bonferroni tests (Harrison et al., 2006, Attar et al., 2013, Sunyer et al., 2007).

Traslational Research

The Barnes Maze is a simple and straightforward task to assess spatial learning and memory in neurocognitive diseases, neurodegenerative diseases, and traumatic brain injury models.

The benefits of exercise have often been explored as a therapeutic intervention for cognitive improvement. In their study regarding the effects of Whole-brain irradiation (WBI) therapy, Wong-Goodrich et al. were able to show that running can abrogate the progressive learning and memory deficits induced by WBI and aid in the recovery of adult hippocampal plasticity. In the study that was conducted by Wu et al., swimming exercise was observed as a promising therapeutic option in the prevention of neurodegeneration in the elderly and/or AD population.

Space radiations present a health risk to astronauts spending long missions in space. Rudobeck’s et al. investigation aimed to understand the impact of protons, the main constituent of the space radiation spectrum, in accelerating the onset of Alzheimer’s disease and AD-related pathology.

Strengths and Limitations

In comparison to the Morris Water Maze and the Radial Arm Maze, the Barnes Maze is relatively less stressful. The MWM subjects the animal to significantly more stress as the subject must be submerged in water and swim in order to survive and search for the escape platform (Hodges 1996, Harrison et al., 2006). However, some groups report that there is little difference in stress and anxiety between the two mazes (Harrison et al., 2006).

Although the BM is considered a less anxiogenic alternative to other behavioral assays, aversive stimuli such as bright light and adverse noise can be used to encourage explorative drive in finding the escape hole. The absence of significant stressors, such as forced swimming and food/water deprivation, allows for better observations of working and reference memory in the animals as they perform in the maze.

In the absence of aversive stimuli to motivate the subject to seek the enclosed target chamber, the subject may simply explore the maze rather than completing the task. Further, if the maze is being used for multiple animals, proper cleaning of the apparatus is a must to ensure no olfactory cues from previous trials influence the performance of subsequent subjects. This can be easily achieved by cleaning the maze before and after each trial.

As with all mazes that measure aspects of learning and memory, it is important to remember that many different processes play into the behavior observed in the maze. Factors such as anxiety and exploratory activity should be considered when interpreting the results of a spatial memory task.

Summary

• The Barnes Maze was designed and developed by Carol Barnes in 1979.
• The Barnes Maze has similar principles as the MWM and RAM; however, unlike those behavioral assays, the BM does not use strong aversive stimulus.
• The Barnes Maze is considered as a less stressful alternative to the MWM.
• The Barnes Maze uses an elevated circular platform that has equally spaced holes placed at the perimeter of the platform.
• The subject’s natural tendency to seek sheltered, dark spaces is challenged by the open and brightly lit platform, forcing it to explore the maze for the target hole.
• Intra- and extra-maze cues are often used to assist the subject in remembering the location of the target hole.
• The difficulty of the maze can be varied by removing the maze cues.
• Animals in control groups show rapid learning as they remember the location of the target hole, while in comparison, animals as disease models of neurodegenerative disorders show a much slower learning curve.

Specifications

Features

1. Highest Quality – built from high-quality stainless steel (harder than acrylic, aluminum, and zinc), our matrices are guaranteed to withstand repeated autoclave sterilizing cycles and scrubbing without damaging the surfaces.
2. Highly Accurate – the durable stainless steel material drives the cutting blade firmly to provide the accuracy needed for your experiment/research.
3. Superfine – cutting-edge manufacturing techniques ensure the narrowest channels separated by the finest walls available in the market, allowing you to dissect your sample into fine slices that are as much as 30 percent thinner than previously attainable with standard 0.5 - 1.0 mm matrices.
4. Multiple Uses – our matrices are perfect for creating sections for biochemical analysis of various substances; for pharmacokinetic studies using the brain slice uptake technique; and for accurate blocking prior to microtome sectioning.

Specifications

Features

1. Highest Quality – built from high-quality stainless steel (harder than acrylic, aluminum, and zinc), our matrices are guaranteed to withstand repeated autoclave sterilizing cycles and scrubbing without damaging the surfaces.
2. Highly Accurate – the durable stainless steel material drives the cutting blade firmly to provide the accuracy needed for your experiment/research.
3. Superfine – cutting-edge manufacturing techniques ensure the narrowest channels separated by the finest walls available in the market, allowing you to dissect your sample into fine slices that are as much as 30 percent thinner than previously attainable with standard 1.0 mm matrices.
4. Multiple Uses – our matrices are perfect for creating sections for biochemical analysis of various substances; for pharmacokinetic studies using the brain slice uptake technique; and for accurate blocking prior to microtome sectioning.

Dimensions

Features

1. Our rodent warming pads are produced with the finest materials, by our expert manufacturers to provide a professional with a high-quality product with optimal performance.
2. Ideal for use before, during, and after surgical procedures on mice and rats.
3. Animals frequently become hypothermic when exposed to anesthesia for a variety of reasons. Therefore, the use of our heating pads before any surgical procedure can minimize heat loss during anesthetic administration. Our heating pads are also optimal to maintain temperature during surgical procedures and they can be placed in the rodent’s cage to provide faster recovery after any surgical procedure.
4. The best option to obtain the best surgical outcomes warming animals quickly, safely, and efficiently.
5. Our laboratory heating pads are designed to fit all your surgical needs as they are available in three different sizes: mouse, rat, and home cage, and they are able to fit in standard stereotaxic instruments.

Specifications

1. Warms Animals quickly, safely, and efficiently.
2. Produced with High-Quality Materials.
3. Available in Three Different Sizes: Mouse, Rat, and Home Cage.
4. Ideal for use before, during, and after surgical procedures.
5. Temperature Control Range: 25-45⁰C
6. Temperature Resolution : 0.1⁰C

Introduction

Heating pads are extensively used for thermoregulation before, after, and amidst rodent surgeries. The ambient temperatures (20-24oC) of laboratories and vivariums where rodents are housed can cause hypothermia in the animals. Moreover, rodents are also prone to cold stress due to anesthesia and while recovering from anesthesia. Heating pads create thermoneutral environments in the cages, during and after surgery, preventing inadvertent hypothermia. 

The use of anesthetics for sedating animals during surgery interferes with normal temperature regulation mechanisms of the body. Exposure of skin to drugs and injection of large volumes of intravenous and irrigation fluids can result in significant loss of body heat, ultimately causing perioperative and postoperative hypothermia. This hypothermia is the leading cause of mortality in rodents due to their "high surface area to body mass ratio." Therefore, the animal is provided with warmth via a heating pad during experiments to save them from cold stress.

In a general stereotaxic surgery (such as for implanting a probe or cannula in rodents' brains), the heating pad is pre-warmed for 30 minutes before use. The rodents are then anesthetized using an intraperitoneal or respiratory drug. The animal is placed on a stereotaxic instrument, and loss of reflex is confirmed by toe pinching. The heating pad is then placed beneath the animal with a towel on the top to avoid thermal burns. The surgery is performed and the animal is shifted to the recovery cage. The heating pad is then placed underneath the recovery cage to prevent postoperative hypothermia. The duration for which a heating pad is used depends on the effect of the anesthetic/drug used. The heating pad should be warm enough not to let the temperature drop by 37oC. However, in hyperthermia (where the temperature exceeds 38oC), one must turn the heating pad off. 

Apparatus and Equipment

Conduct Science’s heating pad is made of non-toxic silicone. It offers temperature control between 25 and 45oC with a resolution of 0.1 degree Celsius. The heating pad is a part of the Homeothermic Monitoring System designed for optimal performance before, during, and after surgery. It can adapt well to multiple experimental platforms and is easy to use. You need to connect it to the Temperature Controller. It is available in three different sizes: rat, mouse, and cage, and is compatible with various stereotaxic instruments. The experimenter can easily clean the apparatus after use. 

Applications

Maintenance of Normothermic conditions during Dexmedetomidine administration 

Lavon et al. (2017) studied the effect of the drug Dexmedetomidine on metastasis in rodent models of breast, lung, and colon cancers. However, this drug is reported to cause potential hypothermia in rodent models. The animal can face cold stress for up to 8 hours after Dexmedetomidine administration. The researchers used heating pads (temperature adjusted at 40oC) for about 8 hours to resolve this problem. The hypothermic effects of the drug were easily overcome by the heating pad that induced normothermic conditions in the subject. They injected the subjects with the drug of choice, studied its effects, and concluded that dexmedetomidine increases metastasis and tumor cell retention in mammary and colon adenocarcinoma.

Prevention of Hypothermia in Stereotaxic Surgeries

Poole et al. (2019) anesthetized the animals for stereotaxic surgery to implant guide cannulas in rodent brains for administering drugs to the targeted regions and used a heating pad for thermoregulation. They took 28 to 30 days old postnatal male Sprague-Dawley rats weighing up to 120g housed in normal cages and shifted them into an induction chamber filled with 0.5% isoflurane. The isoflurane percentage was increased by 0.5% every 20 seconds until it reached 3.5%. The animals were allowed to inhale 3.5% isoflurane for approximately 3 minutes to get anesthetized. The animal was fixed into the stereotaxic frame. To prevent hypothermia, they placed the animal on a heating pad. A towel was placed as a barrier to avoid the animal's direct contact with the heating pad. The lack of sensation was confirmed by 'the pinch withdrawal reflex. They decreased the isoflurane concentration to 3% and then used a lubricated rectal thermometer for a temperature check. Researchers also ensured that the heating pad did not let the temperature drop below 37oC and increase above 37.5oC. They shaved the skin of the animal; the skull was incised and open for full exposure. And then removed pericranial tissues via cotton swabs, and the cannula was mounted on the bregma. In a nutshell, the researchers successfully implanted the cannulas with an overall mortality rate of 0%.  

Heating Pad’s Efficiency Assessment

Zhang et al. (2017) assessed the efficiency of the heating pad after Isoflurane administration in Sprague-Dawley rats. They aimed to ascertain an effective 'warming period' for maintaining normothermia during recovery. The researchers took eight male and nine female rats free from any bacterial or viral infection, habituated in polycarbonate cages in the form of pairs in a controlled housing environment (22% humidity, 23oC temperature, and 12:12h light and dark reactions). The animals were provided with food and water ad libitum. They pre-warmed the heating pad for 30 minutes and assessed its performance by measuring the temperature at different points on its surface. The rats were also acclimated to the recovery cages before testing, where they were handled for 15 minutes by the experimenter and presented with a reward. The animals were exposed to 5% isoflurane for 40 minutes until the loss of reflex and then shifted into the recovery cages. They were placed into two treatment groups: a) 30 minutes warming post-recovery and b) 60 minutes warming post-recovery. Experimenters measured the rectal and cage floor temperatures every 10 minutes, and the time spent in the recovery cage was 2 hours for each group.  This duration for the first group was divided into “30 min in the recovery cage and 90 min in the home cage”. For the second group, this time was divided into “60 min in the recovery cage and 60 min in the home cage”. They concluded that 60 minutes is an effective warming period for preventing rodents from hypothermia while recovering from general anesthesia.

Strengths and Limitations

The heating pads are easy to use and cost-effective. They can be adjusted according to the experimental setup and can be easily cleaned after use. They efficiently prevent rodents from hypothermia before, during, and after surgeries. However, direct contact of the heating pad with the rodents can cause thermal burns. The heating pads are placed partially under the cage such that only 50% of the cage is above the heating pad to avoid this issue. Also, an insulator like a towel is placed over the heating pad to avoid direct contact with the subject. 

Summary

• Heating pads are extensively used for thermoregulation before, after, and amidst rodent surgeries.
• Heating pads prevent pre-operative, postoperative, and perioperative hypothermia.
•  The heating pad should be warm enough not to let the temperature drop by 37oC. However, in hyperthermia (where the temperature exceeds 38oC), the heating pad is turned off. 
• Overheating of the pad can result in thermal burns.

References

1. Lavon, H., Matzner, P., Benbenishty, A., Sorski, L., Rossene, E., Haldar, R., Elbaz, E., Cata, J. P., Gottumukkala, V., & Ben-Eliyahu, S. (2018). Dexmedetomidine promotes metastasis in rodent models of breast, lung, and colon cancers. British journal of anesthesia, 120(1), 188-196.
2. Zhang, E. Q., Knight, C. G., & Pang, D. S. (2017). Heating pad performance and efficacy of 2 durations of warming after isoflurane anesthesia of Sprague–Dawley rats (Rattus Norvegicus).  Journal of the American Association for Laboratory Animal Science, 56(6), 786-791.
3. Poole, E. I., McGavin, J. J., Cochkanoff, N. L., & Crosby, K. M. (2019). Stereotaxic surgery for implantation of guide cannulas for microinjection into the dorsomedial hypothalamus in young rats. MethodsX, 6, 1652-1659.
4. Chaejeong Heo, Hyejin Park, Yong-Tae Kim, Eunha Baeg, Yong Ho Kim, Seong-Gi Kim, Minah Suh. (2016). A soft, transparent, freely accessible cranial window for chronic imaging and electrophysiology. Scientific Reports, 6: 27818.
5. Danny Florez-Paz, Kiran Kumar Bali, Rohini Kuner, Ana Gomis. (2016). A critical role for Piezo2 channels in the mechanotransduction of mouse proprioceptive neurons. Scientific Reports, 6: 25923.
6. Megan E. Poorman, Vandiver L. Chaplin, Ken Wilkens, Mary D. Dockery, Todd D. Giorgio, William A. Grissom, Charles F. Caskey. (2016). Open-source, small-animal magnetic resonance-guided focused ultrasound system. Journal of Therapeutic Ultrasound, 4:22.
7. Gregor-Alexander Pilz, Stefano Carta, Andreas Stäuble, Asli Ayaz, Sebastian Jessberger, Fritjof Helmchen. (2016). Functional Imaging of Dentate Granule Cells in the Adult Mouse Hippocampus. Journal of Neuroscience, 36 (28) 7407-7414.
8. David P. Ferguson, Lawrence J. Dangott, J. Timothy Lightfoot. (2014). Lessons learned from vivo-morpholinos: How to avoid vivo-morpholino toxicity. Biotechniques, 56(5): 251–256.

Dimensions

Information

Broome Rodent Restrainer is one of the most convenient types of restrainer for ease of use as it allows for both hands to be used when injecting the test subjects' caudal vein. Our Broome restrainer is made up of the finest quality acrylic and comes in a variety of sizes. Choosing the right size of the restraint is vital; if the restraint is too large the animal can easily turn around, and if the restraint is too small the animal might not be able to breathe properly.

A broome rodent restrainer is a cylindrical device that is closed on one end. An open groove extends from the opening to the center of the endplate for the adjustment of the nosepiece. On the adjacent side of the open groove, a slit that runs the entire length of the tube aids to get access to the rodent’s tail.

Broome Rodent Restrainer is one of the most convenient types of restrainer when the ease of the experiment is in question; the experimenter can use both his/her hand when injecting the caudal vein. Our Broome restrainer is made up of the finest quality acrylic and comes in a variety of sizes. Choosing the right size of the restraint is vital; if the restraint is too large the animal can easily turn around, and if the restraint is too small the animal might not be able to breathe properly.

A broome rodent restrainer is a cylindrical device that is closed on one end. An open groove extends from the opening to the center of the endplate for the adjustment of the nosepiece. On the adjacent side of the open groove, a slit that runs the entire length of the tube aids to get access to the rodent’s tail.

First, the nosepiece should be removed from the body of the Broome-style restrainer by loosening the screw. Position the restrainer so that the slit that runs the entire length of the tube is facing up. Hold the subject by the base of the tail and gently introduce the subject to the restraint device.

Note: Hind limbs should be introduced first. Furthermore, it is always beneficial to place the animal on a smooth surface to expedite its placement into the Broome-style restrainer.

Once the subject is introduced securely, the nosepiece is placed in the groove and tightened so that the open end is sealed. Ideally, the nose of the subject should be projected at the center of the nosepiece. Caution: To avoid suffocation caused to the subject, the nosepiece should not be over-tightened.

Our Broome style restrainer is exceptionally advantageous:

1. Minimum handling is required; once the subject is secured, the experimenter can simply immerse the tail in warm water for vasodilation.
2. Minimum stress evoked in subjects; the Broome style handler calms the animals as they will not try hard to escape as an escape is blocked.
3. The cylindrical design impedes the movement of the rodents once they are safely secured within the restrainer.
4. Equally favorable for injection delivery and blood sampling techniques.

Proper handling and management of experimental animals is an important aspect of various research procedures. Minimal handling, such as cage changing and other noninvasive procedures have been found to cause physiological manifestations of stress in rodents, including elevated heart rate and blood pressure levels (Stewart and Schroeder., 2017). Given that prolonged effects of stress can influence research outcomes, it is imperative that researchers are equipped with the right knowledge, training, and tools for proper animal handling and restraint methods.

The Broome restrainer is one of many types of restraint devices that provide a tightly enclosed fit around the rodent, designed to give access to the tail for intravenous injections. The Broome handler is a cylindrical device that is closed on one end, with an open groove extending from the open end to the center of the endplate for the adjustment of the nosepiece.

A slit runs the entire length of the tube, serving to accommodate the rodent’s tail. Injecting the rodent through the caudal vein usually requires more than one pair of hands to hold the animal down and to secure the area for injection, but the Broome restrainer successfully addresses this commonly-faced difficulty through its novel design. The restraint devices such as the Broome handler are useful for injections and blood sample collection.

Apparatus and Equipment

The Broome restrainer is a cylindrical, close-fitting restraint device made from fine quality acrylic that comes in different sizes to accommodate different rodent types. The device is transparent and closed on one end, with a slit running the entire length of the tube, giving access to the rodent’s tail. An open groove on the cylinder extends from the opening to the center of the endplate, to fix the adjustable nose piece in place.

Choosing the right size of the restraint is vital; if the restraint is too large the animal can easily turn around, and if the restraint is too small the animal might not be able to breathe properly.

Protocol

In the usage of a Broome restrainer, it is important to prepare the correctly-sized device for each rodent. An appropriate fit would prevent the animal from fidgeting easily or turning around but is loose enough to allow the animal to breathe properly.

First, the nosepiece should be removed from the body of the Broome-style restrainer by loosening the screw. Position the restrainer so that the slit that runs the entire length of the tube is facing up. Hold the subject by the base of the tail and gently introduce the subject into the restraint device.

Note: Hind limbs should be introduced first. Furthermore, it is always beneficial to place the animal on a smooth surface to expedite its placement into the Broome-style restrainer.

Once the subject is introduced securely, the nosepiece is placed in the groove and tightened so that the open end is sealed. Ideally, the nose of the subject should be projected at the center of the nosepiece. Caution: To avoid suffocation caused to the subject, the nosepiece should not be over-tightened.

Applications

The use of the right techniques in the proper handling of mice and rats is a necessity to minimize the effects of stress and discomfort on research outcomes such as fluctuations in physiological parameters. Rigid restraint devices are practical tools that make animal handling easier, safer, and more trouble-free. The Broome style handle, in particular, is designed to provide convenient access to the rodent’s tail for intravenous injections as well as blood sample collection. Modified Broome handlers have found many applications, such as in the monitoring of fetal breathing movements through abdominal ultrasound measurements in rats (Kobayashi et al., 2001), development of rodent models for intraocular pressure fluctuations in patients with glaucoma (Joos et al., 2010), injections through the lateral tail vein (Dorsey et al., 2009), and the regulation of restraint stress (Seifi et al., 2014; Jamieson et al., 2016).

Strengths and Limitations

- The Broome restrainer is one of many types of restraint devices that provide a tightly enclosed fit around the rodent, designed to give access to the tail for intravenous injections

- The Broome restrainer is a cylindrical, close-fitting restraint device made from fine quality acrylic that comes in different sizes to accommodate different rodent types

- The device is closed on one end, with an open groove extending from the open end to the center of the endplate for the adjustment of the nosepiece, and a slit runs the entire length of the tube, serving to accommodate the rodent’s tail

- Restraint devices such as the Broome restrainer are useful for injections and blood sample collection

- Choosing the right size of the restraint is vital; if the restraint is too large the animal can easily turn around, and if the restraint is too small the animal might not be able to breathe properly

References

1. Karen M. Joos, Chun Li, Rebecca M. Sappington. (2010). Morphometric Changes in the Rat Optic Nerve Following Short-term Intermittent Elevations in Intraocular Pressure. Investigative Ophthalmology & Visual Science, Vol.51, 6431-6440
2. Kay Stewart, Valerie A. Schroeder. University of Notre Dame. (2017). Rodent Handling and Restraint Techniques. Retrieved from https://www.jove.com/science-education/10221/rodent-handling-and-restraint-techniques
3. Koichi Kobayashi, Robert P. Lemke, John J. Greer. (2001). Ultrasound measurements of fetal breathing movements in the rat. Journal of Applied Physiology, vol. 91 no. 1, 316-320
4. Mohsen Seifi, James F. Brown, Jeremy Mills, Pradeep Bhandari, Delia Belelli, Jeremy J. Lambert, Uwe Rudolph and Jerome D. Swinny. (2014). Molecular and Functional Diversity of GABA-A Receptors in the Enteric Nervous System of the Mouse Colon. Journal of Neuroscience, 34 (31) 10361-10378
5. Pauline M. Jamieson, Chien Li, Christina Kukura, Joan Vaughan, Wylie Vale. (2016). Urocortin 3 Modulates the Neuroendocrine Stress Response and Is Regulated in Rat Amygdala and Hypothalamus by Stress and Glucocorticoids. Endocrinology, 147 (10): 4578-4588
6. Susan G. Dorsey, Carmen C. Leitch, Cynthia L. Renn, Sherrie Lessans, Barbara A. Smith, Xiao M. Wang, Raymond A. Dionne. (2009). Genome-Wide Screen Identifies Drug-Induced Regulation of the Gene Giant Axonal Neuropathy (Gan) in a Mouse Model of Antiretroviral-Induced Painful Peripheral Neuropathy. Biol Res Nurs, 11(1): 7–16.

Apparatus and Equipment

Procedure

Applications

Strengths and Limitations

Summary

• An injection cone is used to restrain the rodents for administering intravenous tail injections during stereotaxic rodent surgeries.
• It can be used for anesthesia administration, intravenous injections, and monitoring rodent stress responses.
• The transparent cone design ensures a complete and accessible vision of the restrained animal.
• The injection cone is the easiest and quickest rodent restraint for intravenous, intraperitoneal, or subcutaneous drug/anesthesia administration.

References

• Weigh the subject and determine the volume of drug to be administered.
• The light color is adjusted to meet the best contrast of the caudal vein.
• Fill the syringe with the drug under consideration and remove air bubbles that may evolve.
• Critical: The injection must not be cold, administering large quantities of the cold solution may cause hypothermia which may alter the efficacy of the drug under consideration.
• Carefully introduce the subject to the Broome handler. (Refer to Broome handler’s protocol section for detailed instructions).
• After restraining the subject, immerse its tail in the warm water bath set at 42°C for approx. 40 to 45 seconds for vasodilation. Note: A beaker should be placed in the water bath to restrict contamination by urine and feces.
• Furthermore, warming up the whole subject with heat lamps may prove to be more efficient, but it takes longer and carries a risk of heatstroke.
• Dry the tail and try to locate the left or right caudal vein. The restrainer is fixed to the stand at an angle of 90° so that the chosen caudal vein faces the experimenter.
• The tail should then be held with the thumb of one hand, and the needle should be inserted gently with the preferred hand. Note: The needle should be parallel to the vein; almost a 0° angle, and should not be penetrated more than 6-7mm inside. Critical: Do not attempt to extract the blood back into the needle as this may cause the vein to rupture.
• The plunger should be pushed be pressed gently and slowly. Critical: If you feel resistance or see a lump, immediately stop and retract the needle. Insert the needle approx. 1 to 2 cm above the same vein.
• Cover the injection site with a swab before retracting the needle to limit excessive blood and drug loss. Keep pressing the swab until you remove the subject from the restrainer.
• Rinse the restrainer with a sufficient quantity of water before injecting the next subject.

Specifications

1. Ideal for Large Quantity Samplings
2. Produced with High-Quality Acrylic
3. Transparent, Durable, and Easy to Clean

Documentation

Apparatus and Equipment

Protocol

Applications

Strengths and Limitations

Summary

References

1. Messer, Juerg. (2015). Improving intravenous injection in black mice.

Specifications

Anesthesia Induction Chambers

Each model contains 1 plexiglass box, 1.2m bellows and 1 gas filter canister (R510-31S)

Low Stress Anesthesia Induction Chamber

The pipeline connecting the Gas Evacuation. Apparatus is already included

Low stress Anesthesia Induction chamber

Rodents lack cone cells that sense red light and are insensitive to red light.The red transparent appearance is convenient for the experimenter to observe the stateof the animal while reducing the stress response of the animal and reducing the impacton the biological rhythm of the animal. Comply with animal welfare.

Anesthesia Induction Chambers

ConductScience’s Anesthesia Induction Chamber is a state-of-the-art apparatus contrived from supreme quality acrylic. It is used to confine subjects during an induction procedure. The box is provided with two adjacent orifices; an inlet for the entry of the fresh anesthetizing gas and an outlet for scavenging the waste gas material. The transparent material helps in visualizing the physical state of the subject; thus preventing accidental overdosing. The Anesthesia Induction Chamber can be controlled by a flowmeter with inlet air ranging from 0.1-4L/min.

The subject(s) should be carefully placed into the box, and the supply of fresh anesthetizing should be initiated and maintained for approximately 2 to 5 minutes. After the subject is fully anesthetized, the anesthesia supply should be discontinued.

Critical: Do not open the anesthesia box immediately, wait for about 10 to 15 seconds so that oxygen in the box neutralizes the gas concentration.

Finally, the chamber should be opened slightly, just enough to remove the anesthetized subject from the box, and closed immediately.

Caution: Do not wide open the chamber, just enough so that a hand can be introduced for the removal of the subject.

By keeping the opening far away from the experimenter and limiting the time that the box is open, the risk of exposure to the experimenter will be diminished.

Optimal Benefits Achieved Through Our Exceptional Design

• Our design is preferred over the conventional mask and circuits because it is less time-consuming and it can hold and anesthetize more than one subject simultaneously.

• Our special design provides extra protection to the experimenter by keeping the opening far away from the experimenter.

  Our device is compatible with almost all (non-explosive) gas mixtures. However, it should not be used with liquid organic solvents.

• Our scavenging tube efficiently removes the waste gas materials.

A fundamental component of surgical procedures in animal research is the usage of anesthetics. Anesthesia is an important tool to restrain animals during procedures that may either cause excessive stress to the animal or expose the researcher to unavoidable hazards, as well as post-surgery pain management. The surgical methods that are sure to cause pain or discomfort in animals must, therefore, be performed under general anesthesia (“Anesthesia and Analgesia in Laboratory Animals,” 2010). Anesthesia may be inhaled or injected depending on factors such as type and duration of surgery and animal species.

The process of putting animals under inhalant anesthesia is made easier with the use of an anesthesia box that confines subjects during the anesthesia induction procedure. It is also provided with two separate openings, an inlet for the entry of fresh anesthetizing gas and an outlet for scavenging the waste gas material. To prevent accidental overdosing, the apparatus is made of transparent acrylic for easy observation of the animal

Apparatus and Equipment

The anesthesia box is a state-of-the-art apparatus manufactured from supreme quality acrylic, used to enclose animals during inhalant anesthesia induction procedures. The transparent acrylic material is used for convenient inspection of the animal’s physical state, in order to prevent any possibility of accidental overdose.

Protocol

The anesthesia box must be carefully placed on an even and stable surface. Depending on its size, the anesthesia box may accommodate more than one animal.

Once the animal is carefully placed inside the empty anesthesia box, the supply of fresh anesthetizing gas is then initiated and maintained for approximately 2 to 5 minutes. The supply should then be discontinued once the animal is fully anesthetized. A critical step is to wait for about 10 to 15 seconds after the anesthesia supply is cut off so that the oxygen in the box neutralizes the gas concentration. Once the animal is retrieved through a slight opening in the top lid, the anesthesia box is immediately closed to prevent the unwanted escape of gas. Once the animal is moved out of the chamber, anesthesia may be maintained via other tools, such as the cone device.

Applications

The discovery of general anesthesia during the middle of the 19th century can be considered one of the most important developments in the history of medicine (Werner et al., 2011), and since then, its usefulness has found other important applications, such as animal research.

Much of research in the neurosciences is benefited from the use of experimental animals. A huge aspect of animal research is the employment of surgical procedures, and ethical considerations necessitate anesthesia as a tool for eliminating any unnecessary stress, pain, or discomfort the animal may experience during and after surgery. Today, there are many novel and sophisticated methods for administering different types of anesthesia, but the mechanisms have not changed. The anesthesia box presents a convenient way of employing gas induction procedures, with such substances as isoflurane, sevoflurane, and other inhalant agents, that, through inlet and outlet openings of gas and a transparent, sealed design, ensures the safety of everybody involved.

Strengths and Limitations

The anesthesia box’s design is much preferred over the conventional mask and circuit method because it requires less time for the inhalant anesthesia to take effect. Further, the anesthesia box can hold and anesthetize more than one animal simultaneously, depending on its size.

It is a reality that exposure to waste anesthetic gas is a serious occupational hazard (“Anesthesia and Analgesia in Research Animals,” 2012). The experimenter’s safety is, therefore, another concern that the anesthetic box’s design succeeds in addressing. The anesthesia box’s special design provides extra protection to the experimenter by keeping the opening on the farther side to avoid exposure. Additionally, the anesthesia box is compatible with most non-explosive gas mixtures. The outlet scavenging tube is also an efficient addition to the anesthesia box’s design, effectively removing toxic waste gas materials.

Summary

• An anesthesia box is a tool used for confining animal subjects in a closed space during anesthesia induction procedures.
• To prevent accidental overdosing, the apparatus is made of transparent acrylic for easy observation of the physical state of the animal.
• The anesthesia box ensures the safety of experimenters from accidental exposure to waste anesthetic gas.

References

Anesthesia and Analgesia in Laboratory Animals. (2010, November 08). Retrieved from Penn State Animal Research Program, https://www.research.psu.edu/arp/anesthesia.html

Anesthesia and Analgesia in Research Animals. (2012, December 01). Retrieved from https://lar.indiana.edu/doc/Anesthesia_and_Analgesia_in_Research_Animals.pdf

Werner, D. F., Swihart, A., Rau, V., Jia, F., Borghese, C. M., McCracken, M. L., ... & Eger, E. I. (2011). Inhaled anesthetic responses of recombinant receptors and knockin mice harboring α2 (S270H/L277A) GABAA receptor subunits that are resistant to isoflurane. Journal of Pharmacology and Experimental Therapeutics, 336(1), 134-144.[/vc_column_text][vc_column_text]Quan Ren, Mian Peng, Yuanlin Dong, Yiying Zhang, Ming Chen, Ning Yin, Edward R. Marcantonio, Zhongcong Xie. (2015). Surgery plus anesthesia induces loss of attention in mice. Front Cell Neurosci, 9: 346

Amy Miller, Gemma Kitson, Benjamin Skalkoyannis, Matthew Leach. (2015). The effect of isoflurane anaesthesia and buprenorphine on the mouse grimace scale and behaviour in CBA and DBA/2 mice. Appl Anim Behav Sci, 172: 58–62

Lundt A, Wormuth C, Siwek ME, Muller R, Ehninger D, Henseler C. (2016). EEG Radiotelemetry in Small Laboratory Rodents: A Powerful State-of-the Art Approach in Neuropsychiatric, Neurodegenerative, and Epilepsy Research. Neural Plast, 8213878

F. Werner, A. Swihart, V. Rau, F. Jia, C. M. Borghese, M. L. McCracken, S. Iyer, M. S. Fanselow, I. Oh, J. M. Sonner, E. I. Eger, N. L. Harrison, R. A. Harris, G. E. Homanics. (2011). Inhaled Anesthetic Responses of Recombinant Receptors and Knockin Mice Harboring α2(S270H/L277A) GABAA Receptor Subunits That Are Resistant to Isoflurane. Journal of Pharmacology and Experimental Therapeutics January, 336 (1) 134-144

Michael A. Makara, Ky V. Hoang, Latha P. Ganesan, Elliot D. Crouser, John S. Gunn, Joanne Turner, Larry S. Schlesinger, Peter J. Mohler, Murugesan V.S. Rajaram. (2016). Cardiac Electrical and Structural Changes During Bacterial Infection: An Instructive Model to Study Cardiac Dysfunction in Sepsis. Journal of the American Heart Association, 5:e003820

• Input power range: 100 - 240V, 50 /60Hz, 24V, 40W, 1.67A adapter DC power supply
• Provide safes and effective intermittent positive pressure ventilation (IPPV)
• Volume and pressure modes are available;
• Sound, intelligent text, error code and other fault alarm alerts
• Small & compact.
• 7 inch LCD touch screen
• The pressure-time graph can be displayed in real time in volume and pressure modes
• 10 stored parameters can be entered and recalled for maximum convenience.
• The respiration rate (RR) can be switched between 10 – 300 rpm
• Tidal volume range:0.05ml ~ 5ml
• Airway pressure range: 1-50cm H₂O, the accuracy range: ±0.7 cmH₂O; Positive End Expiratory Pressure (PEEP) range is 0-10cmH₂O
• Set the respiratory parameters such as PIP/PEEP/Plateau Pressure/INSP.Hold/EXP. Hold;
• After entering the animal weight, the device automatically sets the reference breathing parameters
• The adjustable range of I:E ratio is from 20% to 80%; the plateau pressure increases the tidal volume from 0 to 20%, and can set auto-peep once after every 10 to 999 breaths or manual control

Information

Apparatus and Equipment

Specifications

Mode of Operation

Protocol

• Place the subject on its back.
• Hold the laryngoscope in your left hand.
• Extend the subject’s tongue forward and to the left.
• Slowly insert the blade into the right side of the subject’s mouth.
• Advance the blade inward and midline towards the base of the tongue.
• Place the tip of the blade under the epiglottis.
• Apply pressure caudally and upward with the handle placed at an angle of 45 degrees.
• Lift the handle until vocal cords are visualized.
• Perform intubation under direct visualization of vocal cords.
• Withdraw the blade while firmly holding the endotracheal tube in place.

• Adequately anesthetize the subject and place it in the supine position with its limbs restrained.
• Extend the neck. Remove the hair below the mandible and ensure local asepsis of the area.
• Make an incision midline below the neck.
• Separate the salivary glands and fix the muscles surrounding the trachea with sutures.
• Once the trachea is visualized, make an incision between the fourth and the fifth tracheal ring and insert the cannula.

Applications

Strengths and Limitations

Summary

• Mechanical ventilation is used to support or replace spontaneous breathing. They often are a part of anesthetization process, especially when neuromuscular blocking agents are used.
• Mechanical ventilators can potentially cause ventilator-induced lung injury, lead to the rapid type of disuse atrophy in respiratory-related muscles, barotrauma, and impairment of mucociliary motility in the airways.
• The use of mechanical ventilators has been extended to the research of ventilator-induced lung injuries and for understanding the relationship between ventilator settings and certain biological responses.
• Prolonged ventilation of neonates and infants raises their risk of developing long-term clinical complications.
• Mechanical ventilation is performed either via endotracheal intubation or by cannulating the trachea (tracheotomy).
• It is important that the animal is sufficiently anesthetized before ventilation and is placed in an appropriate position to prevent injury.
• Good anesthesia practice should involve appropriate anesthesia management and monitoring techniques, and recovery care.
• Neuromuscular blocking agents only produces paralysis. Thus efforts to monitor the depth of anesthesia is critical for the humane treatment of the animals.
• Anesthetization of pregnant animals and neonates should be done with extreme caution to prevent undesirable effects.

Features:

• Adopt pyroelectric probe, applicable to multiband measurement with a broad-spectrum range.
• Short response time, good thermal stability, small volume, convenient installation, and fixed.
• Convenient operation with a digital display, applicable to the measurement on a variety of laser power.
• Automatic shutdown (about 30 minutes later) facilitates saving battery power.

Documentation

Introduction

1. Thermal detectors
2. Quantum detectors

Principle

Apparatus and Equipment

Applications

Strengths and Limitations

Summary

• Laser power meters are used to measure the laser power in laser-equipped labs.
• There are two kinds of laser power meters based on the type of detectors used. The detectors can be thermal as well as quantum detectors.
• The most commonly used quantum detectors are photodiodes. Photodiodes are preferred over thermal sensors because of several reasons.
• A silicon photodiode-based laser power meter works by converting incident laser light into a proportionate current.
• Silicon PDs have high quantum efficiency and, therefore, can measure low laser powers.
• Photodiode-based laser power meters are durable, compact, and lightweight.

References 

1. Guo, Y., Mehrabian, Z., & Bernstein, S. L. (2016). The rodent model of nonarteritic anterior ischemic optic neuropathy (rNAION). JoVE (Journal of Visualized Experiments), (117), e54504.

• 20 MHz sine, 10 MHz pulse waveforms provide coverage for your most common applications
• 250 MS/s sampling rate and 14-bit vertical resolution enable the creation of high-fidelity signals

Key features

• The innovative UI reduces setup and evaluation time with direct access to frequently used functions and parameters
• The internal 4 × 16 kS memory and the USB memory expansion capability provide substantial capacity for defining complex waveforms
• USB host port on the front panel for saving/reloading arbitrary waveforms and instrument settings
• Built-in Modulation, Noise Generator, Burst, and Sweep modes for greater versatility
• Built-in waveforms provide quick access to commonly used signals
• Large 3.5-inch color screen displays both graphical and numeric waveform information simultaneously
• Menu and online help in Simplified Chinese and English
• 2U height and half-rack width fits benchtop applications
• Free ArbExpress software makes waveform editing extremely easy

Applications

• Electronic test and design
• Sensor simulation
• Education and training
• Functional test

• A simple binocular coaxial illumination surgery microscope with a compact design and high flexibility to meet the requirements of general animal microsurgery
• Eyepiece magnification: 12.5X
• Objective lens focal length: 200mm
• Working distance: 190mm
• Total magnification: 5.3X, 8X, 12X
• Field of view diameter: 38mm, 25mm, 17mm
• Diopter adjustment range: ±5D
• Interpupillary distance adjustment range: 50mm-70mm
• Lighting source: 12V/100W, cold reflection medical halogen bulb
• Illumination type: 6°+0° cold light source coaxial illumination
• Coaxial illumination object surface illuminance: ≧20000lx
• Cross arm extension radius: 870mm
• Fine focusing stroke: 30mm
• Voltage: AC220V±10%, 50Hz±1Hz
• Power: 120VA
• Fuse: AC250V T1.25A
• Whole machine weight: 41kg

Introduction

Principles

History

Origin

Development

Apparatus and Equipment

• Eyepieces 12.5X focus length of objective: 200mm
• Working distance 190mm
• Total magnification 5.3X, 8X, 12X

• Diopter adjustment range +5D
• Pupil distance range 50mm~70mm
• Bulb 12V/100W (Medical halogen lamp Bulb with cold reflection)
• Type of Illumination 6°+ 0°cold light source coaxial illumination
• The intensity of Illumination 20000Lx or up
• The maximum stretching radius of the arm is 870mm
• Vertical movement range (from floor to front surface of the objective) 700mm~1100mm
• Range of fine focusing adjustment 30mm
• Input voltage AC220V±10%, 50Hz±1Hz
• Input power 120VA
• Power AC250V T1.25A
• Net weight 41kg

Training Protocol

• First, complete the installation of the device and shift the surgical microscope to an appropriate location for the surgery and bolt the castors.
• Adjust the counter-balanced arm and the main microscope to a convenient position and screw the corresponding lock shut.
• Next, hold the main surgical microscope with the hand and release the up-down positioning knob and rotate the balance knob until the main microscope has free up and down movement.
• Then, switch on the power button, turn the brightness switch on and loosen the corresponding lock to move the device.
• Pull the main microscope over the surgical field within the height of the working distance (which relies upon the focal length of the objective lens), until the images can be viewed in the binoculars and tighten the corresponding lock.
• Adjust the position of the binoculars by hand to achieve the appropriate papillary distance.
• Next, rotate the magnification changer knob to achieve the required magnification level.
• Adjust the illuminator module to achieve the required illumination on the surgical site and get the best clear visual image by the motorized up-down focus method or activate the focus function on the foot control.
• After adjusting the focus, check if the two eyes have varying diopters and alter the diopter on the eyepiece for each eye.
• Use the back-forth inclination function by rotating the inclination knobby back and forth and positioning the main microscope at a proper place.
• In the same manner, adjust the right and left inclination adjustment and bring the microscope to an appropriate position.
• After the procedure is over and the surgical microscope is not in use, switch the bright switch off, and after a few minutes turn off the power switch of the unit.
• Release the corresponding lock and move the arm and main microscope to a suitable spot and wrap it with a plastic-proof cover.

Applications In Surgical Experiments

Maintenance and Precautions

• The objective lenses, eyepieces, and accessories that are not being utilized must always be kept in dust-free containers.
• The external surfaces of optical parts like eyepieces and objective lenses only need to be cleaned when necessary without utilizing any chemical cleaning agent.
• The dust on the surface of optical components may be blown off by utilizing a squeeze blower, or it can be eliminated by utilizing a clean grease-free brush.
• Use an anti-fogging agent to protect the eyepiece optics from fogging.
• All mechanical surfaces of the instrument can be simply cleaned with a damp cloth without the use of aggressive agents.
• Wrap the foot pedal with a clear plastic cover to avoid damage to the electronics from surgical and cleaning fluids.
• Abstain from looking directly into the light source, e.g., into the objective lens or a light guide.
• Do not run the instrument in locations at risk of explosives or containing inflammable anesthetics or volatile chemicals.
• Do not place or utilize the equipment in damp rooms and avoid exposing the instrument to water.
• Promptly unplug any instrument that emits smoke, sparks, or odd sounds.

Strengths

• The objective lenses, eyepieces, and accessories that are not being utilized must always be kept in dust-free containers.
• The external surfaces of optical parts like eyepieces and objective lenses only need to be cleaned when necessary without utilizing any chemical cleaning agent.
• The dust on the surface of optical components may be blown off by utilizing a squeeze blower, or it can be eliminated by utilizing a clean grease-free brush.
• Use an anti-fogging agent to protect the eyepiece optics from fogging.
• All mechanical surfaces of the instrument can be simply cleaned with a damp cloth without the use of aggressive agents.
• Wrap the foot pedal with a clear plastic cover to avoid damage to the electronics from surgical and cleaning fluids.
• Abstain from looking directly into the light source, e.g., into the objective lens or a light guide.
• Do not run the instrument in locations at risk of explosives or containing inflammable anesthetics or volatile chemicals.
• Do not place or utilize the equipment in damp rooms and avoid exposing the instrument to water.
• Promptly unplug any instrument that emits smoke, sparks, or odd sounds.

Limitations

Summary

• The binocular surgical microscope improves the surgeon’s view by providing great magnification, powerful illumination, and a stereoscopic perspective during surgery.
• The basic surgical microscope typically consists of a binocular head, adjustable eyepieces, illumination, foot controls, objective lens, diopter lock button, diopter ring, and magnification changer.
• The device is used in various surgical disciplines such as microvascular surgery, plastic and reconstructive surgery, neurosurgery, otorhinolaryngology, transplantation procedures, oncology, urology, dentistry, etc.
• The binocular surgical microscope eliminates the problems that arise from standing for long durations during conventional surgeries such as fatigue, neck pain, eye strain, and posture-related problems.
• The device can be rather bulky and take up a lot of space as well as require proper and constant maintenance.

References

Principle

Specifications

• View field diameter 20mm
• 0.7X-4.5X Objective lens ratio 1：6.4
• Binocular Drawtube Pupil Distance 55-77mm
• Diopter adjustment ±5
• Diopter 45° Incline 360°Spin
• View field 31.2 mm -5.1mm
• Total Magnification 7X-45X (10XEyepiece), 3.5X-22.5X(10X Eyepiece +0.5X Assist lens)
• Working Distance 100mm (10X Eyepiece)170mm (10X Eyepiece +0.5X Assist lens)
• Package Items Universal support(1 unit), Binocular drawtube (1 unit)，10X Eyepiece(1 pair), 0.5X
• Objective Lens (1 unit), Blinder (1 unit), Dust cover (1 unit), Instruction Manual (1 unit)

Protocol

Applications

Maintenance And Precautions

Strengths

Limitations

Summary

References

Overview

Principle

History

Apparatus and Equipment

RFLSI Pro Laser Speckle Perfusion Imager comprises a micro-imaging system, software system, and a laser light source. The instrument does not require a contrast agent and is able to achieve real-time dynamic blood flow monitoring and video imaging. The monitoring distance is 110mm. The system allows the arbitrary addition of regions of interest (ROI) and vessel diameter measurements during the recording process or during off-line analysis to support any shape and number of ROI choices. The monitoring area and blood flow imaging spatial resolution can be adjusted accordingly as well. The monitoring records can be exported to AVI format video files, including curves, blood flow, and experimental process records, and the output video can be adjusted as required.

Protocol

Applications

Strengths and Limitations

Strengths

Limitations

Summary

• The laser speckle perfusion imager (LSPI) is a vascular imaging apparatus based on the laws of speckle contrast analysis and provides an index of blood flow.
• The LSPI uses a charge-coupled device (CCD) camera and an image processing method.
• The LSPI technique has extended to several medical fields like dermatology, plastic & reconstructive surgery, cardiology, vascular solution, diabetology, neuroscience, and ophthalmology, etc.
• LSPI has the advantage of better blood flow parameters, higher temporal resolution, and high reproducibility of data.
• LSPI has the drawback of LSPI delivering blood flow parameters in arbitrary units and generating overly large amounts of information.

References

Features

Introduction

Apparatus and Equipment

Mode of Operation

Precautions

References

• Braz, M. G., M. Carvalho, L. I., Chen, Y. O., Blumberg, J. B., Souza, K. M., Arruda, N. M., A. Filho, D. A., Resende, L. O., G. Faria, T. B., Canário, A., Corrêa, C. R., C. Braz, J. R., & Braz, L. G. (2020). High concentrations of waste anesthetic gases induce genetic damage and inflammation in physicians exposed for three years: A cross-sectional study. Indoor Air, 30(3), 512-520. https://doi.org/10.1111/ina.12643

• Lucio, L. M. C., Braz, M. G., Nascimento Junior, P. do ., Braz, J. R. C., & Braz, L. G.. (2018). Occupational hazards, DNA damage, and oxidative stress on exposure to waste anesthetic gases. Revista Brasileira De Anestesiologia, 68(1), 33–41.

• Smith F. D. (2010). Management of exposure to waste anesthetic gases. AORN journal, 91(4), 482–494.

• This apparatus generates white light for a clear view of surgical procedures.
• Input Power：230V AC/50Hz
• Bulb Power：21V, 150W
• Rated Life： Approx 300h
• Color temperature：3200K, color correction filters are optional（3000-6500k)
• Type of Cooling: Air cooling
• Weight: 2.76Kg
• Dimension: 28.0 cm x 10.0 cm x 16.5 cm

• The speed is continuously adjustable from 0-38,000rpm
• The direction of rotation can be switched between clockwise and counterclockwise
• There are 2 control modes, manual or foot pedal
• Can pass the cranial drill holder (68605)
• Fixed to the brain stereotaxic device, the depth is easy to control
• The range of 0.5-2.3mm can be selected according to the experimental requirements
• Various specifications of drill bits (rod length L: 44mm diameter Φ: 2.3m)

Comes with the following items, with key inclusion of MRI compatible supply

• (1) MRI Compatible mask with tubing for 1 animal (Your choice of Cone or Tubing Mask)
• (1) MRI Compatible Mounting Bracket for Tubing or Cone Mask

Includes:

Introduction

The T-maze is a widely employed operant task across various species. In zebrafish, it has served as a tool to investigate color discrimination (Colwill et al., 2005; Avdesh et al., 2012), memory (Braida et al., 2014; Echevarria et al., 2016), locomotion, exploratory behavior (Peitsaro et al., 2003; Vignet et al., 2013), and place preference (Swain et al., 2004). Studies indicate that zebrafish demonstrate robust long-term retention in spatial alternation (Williams et al., 2002) and active avoidance learning for color discrimination tasks (Pradel et al., 1999), highlighting their suitability for neurobehavioral assessments.

The T-maze setup typically involves a Plexiglas maze with customizable configurations and sizes. Various protocols can be implemented, such as assessing visual discrimination using colored or patterned sleeves in a designated goal area or an enrichment chamber within maze arms. Researchers can employ positive reinforcement like food rewards or environmental enrichment in the goal zones.

Experimental protocols commonly include measuring the time taken to reach the goal zone, percentage of correct choices to evaluate learning, or discrimination between pairs of visual stimuli. Following an initial acquisition phase, extinction and reversal protocols are often applied to eliminate potential biases (Colwill et al., 2005; Avdesh et al., 2012). Parameters such as training duration, session frequency, and number of trials per session can be adjusted according to experimental requirements.

The T-maze enables rapid and reliable toxicological and pharmacological assays, as well as investigations into the effects of genetic manipulations or genetic backgrounds. Results obtained from zebrafish T-maze experiments complement those from rodent models (Levin and Cerutti, 2009). Zebrafish offer advantages for high-throughput studies of neurodevelopment processes at a low cost and with potential for extrapolation to vertebrate systems. Moreover, the functionality and projections of discrete neural systems can be assessed effectively using this model.

Apparatus and Equipment

To conduct this test in zebrafish, essential equipment includes a transparent Plexiglas T-maze. We recommend using a cross design (70 cm x 50 cm x 10 cm) or symmetrical design (50 cm x 50 cm x 10 cm), with flexibility for dimension adjustments (Darland and Dowling, 2001; Colwill et al., 2005). Opaque Plexiglas doors should be removable to block maze arms from goal and start zones.

To enhance visual stimuli, colored or patterned sleeves (e.g., horizontal vs. vertical) can be placed along maze walls in arms or solely in goal boxes. Color choices should consider findings from Avdesh et al. (2012), indicating aversion to blue and preference for green and red over yellow. The maze should contain water filled to a depth of approximately 8-10 cm, using either tank water (deionized water with sea salts; Braida et al., 2014) or a blend of filtered tap water treated with conditioner and tank water (Colwill et al., 2005). Water temperature should be maintained between 25.5 ºC and 28.5 ºC (Colwill et al., 2005; Braida et al., 2014), facilitated by a 25-W heater placed on the maze floor as per Colwill et al. (2005).

Home tanks must have continuous filtration and aeration. During trials, a stopwatch is essential for timing. If the study involves rewards, stainless steel tweezers should be used to dispense food into the designated goal zones.

Training Protocol

T-maze protocols are versatile for testing latency variations in reaching the goal zone across different genotypes or under the influence of drugs, as well as for color or pattern discrimination studies involving acquisition, extinction, and reversal phases. For effective discrimination reversals, a crossover design is recommended, where the reward location and/or sides for colors (green and red) or patterns (horizontal and vertical stripes) are counterbalanced. Maintaining experimenter-blind conditions, as emphasized in other neurobehavioral methodologies, is crucial (Colwill et al., 2005).

Darland and Dowling (2001) initiated the maze exploration with a 5-minute free exploration habituation period for the fish. In contrast, Braida et al. (2014) conducted two daily habituation trials lasting 1 hour each over three days to mitigate procedural novelty and handling stress. Conversely, Colwill et al. (2005) implemented two sessions, each comprising two trials where fish were exposed to food rewards without colored sleeves. Multiple fish can undergo these trials simultaneously (Braida et al., 2014). Group sizes should be gradually reduced until testing is individualized within 3-4 days. The total number, frequency, and trials per individual training session can be adjusted. Colwill et al. (2005) conducted 4 trials per session, with two trials presenting green sleeves on one arm and purple sleeves on the other, and vice versa for the remaining two trials. Braida et al. (2014) conducted two individual training trials spaced 24 hours apart.

For studies involving color or pattern discrimination, half of the fish should have the goal box on the left and the other half on the right, with subsequent reversals. Each fish should spend 2-5 minutes in the start box with the door closed before being allowed to explore the maze. The door should be promptly closed once the fish leaves the start box (Colwill et al., 2005). A 10-minute limit can be set for reaching the goal box (Braida et al., 2014). In trials utilizing food rewards, the reward should be delivered upon correct choices. Fish should be netted 30 seconds after making a choice and returned to the start box or home tank at the end of the session. The experimenter should record whether the fish ate during correct training trials.

During the discrimination acquisition phase, it is important for the investigator to document: a) the latency to reach the goal box and remain there for at least 20 seconds; and b) the color or pattern associated with the choice made. The number of correct trials can also serve as a metric. Following each session, fish should be returned to their home tank, consistent with the training protocol. Upon completion of the acquisition phase, a test trial should be conducted. Depending on the study’s objectives, multiple trials at specific intervals may be necessary to evaluate learning.

For studies involving discrimination between two stimuli, extinction tests should be conducted subsequently. Colwill et al. (2005) conducted 7 tests similar to those used in acquisition, but without food rewards, recording comparable results. It is expected that preference for the side previously rewarded will gradually diminish during extinction (Colwill et al., 2005). Subsequently, a reversal protocol should be implemented, where the previously incorrect choice is now rewarded. Colwill et al. (2005) initiated the first reversal training session the day after the final extinction session, following the same procedures and recording methods as described for acquisition. Similar to the acquisition phase, a final test trial should be performed.

This protocol allows assessment of whether zebrafish exhibit color discrimination when choosing the arm with the goal box, whether this preference declines during extinction when food rewards are discontinued, and whether this preference reverses when reinforcement shifts to the other color or pattern during discrimination reversal.

Darland and Dowling (2001) initiated the maze exploration with a 5-minute free exploration habituation period for the fish. In contrast, Braida et al. (2014) conducted two daily habituation trials lasting 1 hour each over three days to mitigate procedural novelty and handling stress. Conversely, Colwill et al. (2005) implemented two sessions, each comprising two trials where fish were exposed to food rewards without colored sleeves. Multiple fish can undergo these trials simultaneously (Braida et al., 2014). Group sizes should be gradually reduced until testing is individualized within 3-4 days. The total number, frequency, and trials per individual training session can be adjusted. Colwill et al. (2005) conducted 4 trials per session, with two trials presenting green sleeves on one arm and purple sleeves on the other, and vice versa for the remaining two trials. Braida et al. (2014) conducted two individual training trials spaced 24 hours apart.

For studies involving color or pattern discrimination, half of the fish should have the goal box on the left and the other half on the right, with subsequent reversals. Each fish should spend 2-5 minutes in the start box with the door closed before being allowed to explore the maze. The door should be promptly closed once the fish leaves the start box (Colwill et al., 2005). A 10-minute limit can be set for reaching the goal box (Braida et al., 2014). In trials utilizing food rewards, the reward should be delivered upon correct choices. Fish should be netted 30 seconds after making a choice and returned to the start box or home tank at the end of the session. The experimenter should record whether the fish ate during correct training trials.

During the discrimination acquisition phase, it is important for the investigator to document: a) the latency to reach the goal box and remain there for at least 20 seconds; and b) the color or pattern associated with the choice made. The number of correct trials can also serve as a metric. Following each session, fish should be returned to their home tank, consistent with the training protocol. Upon completion of the acquisition phase, a test trial should be conducted. Depending on the study's objectives, multiple trials at specific intervals may be necessary to evaluate learning.

For studies involving discrimination between two stimuli, extinction tests should be conducted subsequently. Colwill et al. (2005) conducted 7 tests similar to those used in acquisition, but without food rewards, recording comparable results. It is expected that preference for the side previously rewarded will gradually diminish during extinction (Colwill et al., 2005). Subsequently, a reversal protocol should be implemented, where the previously incorrect choice is now rewarded. Colwill et al. (2005) initiated the first reversal training session the day after the final extinction session, following the same procedures and recording methods as described for acquisition. Similar to the acquisition phase, a final test trial should be performed.

This protocol allows assessment of whether zebrafish exhibit color discrimination when choosing the arm with the goal box, whether this preference declines during extinction when food rewards are discontinued, and whether this preference reverses when reinforcement shifts to the other color or pattern during discrimination reversal.

Modification

In contrast to color or pattern discrimination, researchers have the option to promote reinforced choices by offering an enrichment chamber equipped with deep water, artificial grass, and marbles (Darland and Dowling, 2001).

The T-maze test has been widely employed across various research fields, including investigations involving flies (Jiang et al., 2016), mice, and rats (Lalonde, 2002).

Sample Data

The T-maze test typically presents results as percentage of correct trials (A), latency to reach the goal zone (B), and/or changes in latency relative to the initial trial (C). These metrics are commonly depicted across test days, encompassing extinction and reversal phases in visual discrimination studies (A). Alternatively, group averages across multiple trials (B and C) are utilized to assess learning effects or the impact of drugs/toxic compounds. Results for metrics B and C are reported as mean ± standard error of the mean.

Strengths and Limitations

The T-maze represents a high-throughput method for swiftly screening potential toxicological and therapeutic agents (Peitsaro et al., 2003; Swain et al., 2004; Echevarria et al., 2016), along with evaluating the impact of genetic manipulation or different genetic backgrounds (Darland and Dowling, 2001).

Zebrafish are advantageous for studying neurodevelopmental processes due to their vertebrate nature, facilitating enhanced extrapolation. Moreover, researchers can analyze the proliferation, differentiation, and migration of discrete neural systems. Zebrafish research also enhances experimental throughput and reduces experimentation costs compared to mammals, as these fish breed prolifically, develop quickly, and require minimal housing space.

When conducting T-maze experiments, careful consideration of color choice is crucial due to its potential impact on outcomes (Colwill et al., 2005; Avdesh et al., 2012). Furthermore, studies have highlighted significant inter-strain variability in this context (Vignet et al., 2013).

Unlike mammals, neurobehavioral studies in fish do not provide insights into the roles of frontal cortical and hippocampal structures in learning and memory processes.

The T-maze represents a high-throughput method for swiftly screening potential toxicological and therapeutic agents (Peitsaro et al., 2003; Swain et al., 2004; Echevarria et al., 2016), along with evaluating the impact of genetic manipulation or different genetic backgrounds (Darland and Dowling, 2001).

Zebrafish are advantageous for studying neurodevelopmental processes due to their vertebrate nature, facilitating enhanced extrapolation. Moreover, researchers can analyze the proliferation, differentiation, and migration of discrete neural systems. Zebrafish research also enhances experimental throughput and reduces experimentation costs compared to mammals, as these fish breed prolifically, develop quickly, and require minimal housing space.

When conducting T-maze experiments, careful consideration of color choice is crucial due to its potential impact on outcomes (Colwill et al., 2005; Avdesh et al., 2012). Furthermore, studies have highlighted significant inter-strain variability in this context (Vignet et al., 2013).

Unlike mammals, neurobehavioral studies in fish do not provide insights into the roles of frontal cortical and hippocampal structures in learning and memory processes.

Summary

• The T-maze in zebrafish is a high-throughput assay that enables rapid screening of toxic and therapeutic candidates, as well as testing the influence of genetic manipulation or diverse genetic backgrounds.
• This test offers the possibility of studying color discrimination, memory, locomotion, exploratory behavior and place preference.
• The equipment required consists of a T-maze with variable configurations. Colored or patterned sleeves can be used to assess visual discrimination; reinforcement can be included using either food rewards in goal boxes or a reservoir with environmental enrichment
• The experimental protocol typically consists of a training period where the fish should learn the location of the rewarded/enriched zone, followed by test trials. If assessing discrimination between colors or patterns, the acquisition period should be followed by extinction and reversal periods in a counterbalanced design.
• Results are typically reported as percentage of correct trials, latency to find goal zone/enrichment chamber with or without assessment of visual discrimination, or change in latency relative to initial trial in order to assess learning.

References

Avdesh A, Martin-Iverson MT, Mondal A, Chen M, Askraba S, Morgan N, Lardelli M, Groth DM, Verdile G, Martins RN (2012) Evaluation of color preference in zebrafish for learning and memory. Journal of Alzheimer’s disease : JAD 28:459-469.

Braida D, Ponzoni L, Martucci R, Sparatore F, Gotti C, Sala M (2014) Role of neuronal nicotinic acetylcholine receptors (nAChRs) on learning and memory in zebrafish. Psychopharmacology 231:1975-1985.

Colwill RM, Raymond MP, Ferreira L, Escudero H (2005) Visual discrimination learning in zebrafish (Danio rerio). Behavioural processes 70:19-31.

Darland T, Dowling JE (2001) Behavioral screening for cocaine sensitivity in mutagenized zebrafish. Proceedings of the National Academy of Sciences of the United States of America 98:11691-11696.

Echevarria DJ, Caramillo EM, Gonzalez-Lima F (2016) Methylene Blue Facilitates Memory Retention in Zebrafish in a Dose-Dependent Manner. Zebrafish 13:489-494.

Jiang H, Hanna E, Gatto CL, Page TL, Bhuva B, Broadie K (2016) A fully automated Drosophila olfactory classical conditioning and testing system for behavioral learning and memory assessment. Journal of neuroscience methods 261:62-74.

Lalonde R (2002) The neurobiological basis of spontaneous alternation. Neuroscience and biobehavioral reviews 26:91-104.

Levin ED, Cerutti DT (2009) Behavioral Neuroscience of Zebrafish. In: Methods of Behavior Analysis in Neuroscience (Buccafusco, J. J., ed) Boca Raton (FL).

Peitsaro N, Kaslin J, Anichtchik OV, Panula P (2003) Modulation of the histaminergic system and behaviour by alpha-fluoromethylhistidine in zebrafish. Journal of neurochemistry 86:432-441.

Pradel G, Schachner M, Schmidt R (1999) Inhibition of memory consolidation by antibodies against cell adhesion molecules after active avoidance conditioning in zebrafish. Journal of neurobiology 39:197-206.

Swain HA, Sigstad C, Scalzo FM (2004) Effects of dizocilpine (MK-801) on circling behavior, swimming activity, and place preference in zebrafish (Danio rerio). Neurotoxicology and teratology 26:725-729.

Vignet C, Begout ML, Pean S, Lyphout L, Leguay D, Cousin X (2013) Systematic screening of behavioral responses in two zebrafish strains. Zebrafish 10:365-375.

Williams FE, White D, Messer WS (2002) A simple spatial alternation task for assessing memory function in zebrafish. Behavioural processes 58:125-132.

Specifications

Description

1. Quickest and easiest restraint for tail vein injections.
2. Unique design cone for rodents which are manufactured by our professional team with high-quality materials, which guarantees a product with great durability.
3. An extremely useful tool for large quantity samplings and their unique design allows an efficient positioning of tailed rodents for maximum results.
4. Our injection cones are an essential tool for any researcher that aims for efficiency and high speed in tail vein injections.
5. With a completely transparent design, the injection cone provides a complete vision of the animal, and its four suction cup feet secure the unit firmly to the countertop.
6. The rodent is held secure in the clear plastic cone and the researcher can realize the procedure with maximum efficiency, limiting the chances of stressing the animal.
7. In two sizes, for mice and rats, the injection cone has a unit that possesses a bright light illumination system for optimal results.
8. Ideal for large quantity samplings.
9. Produced with high-quality acrylic.
10. Transparent, durable, and easy to clean.
11. For use with 110V and 220V power supplies.

Injection Cone with Light

Advantages

Quick and easy restraint for tail vein injections. Unique cone design for rodents, manufactured with high-quality materials for durability.

Efficient positioning for large quantity samplings, ensuring maximum results. Essential tool for researchers aiming for efficiency and speed in tail vein injections.

Completely transparent design for a clear view of the animal; four suction cup feet secure it firmly. Provides secure restraint, minimizing stress on the animal during the procedure.

Available in two sizes for mice and rats, equipped with a bright light illumination system. Ideal for large quantity samplings due to its efficient design.

Produced with high-quality acrylic, ensuring transparency, durability, and easy cleaning. Compatible with both 110V and 220V power supplies.

Principle of Technique

The injection tail technique in rodents, often referred to as "tail vein injection," is a common method used in research to administer substances such as drugs, contrast agents, or other compounds into the bloodstream. This technique is often facilitated by visualizing the tail vein, and one method involves using a cone with light to enhance visibility. Here's an overview of the technical principle:

Animal Restraint.

The rodent is typically restrained to minimize movement during the injection process. Common methods include using a restrainer or a tube that allows access to the tail while keeping the animal stil Vein VisualizationTo visualize the tail vein, a transparent or translucent cone with a light source is often used. The cone is placed over the tail, and the light helps to illuminate the blood vessels, making it easier to identify the vein.

Tail Heating.

Sometimes, gently heating the tail can enhance vein visibility. This is done using a warm water bath or an infrared lamp. The increased blood flow to the tail can make the veins more

Vein Identification.

The researcher identifies the target vein, usually by its color and size. The tail vein is a common choice due to its accessibility and size in rodents.

Injection.

Once the vein is identified, a needle connected to a syringe containing the substance to be injected is carefully inserted into the vein. Precision is crucial to avoid injury and ensure proper administration.

Monitoring.

After injection, the researcher may monitor the animal for any adverse reactions or to ensure that the substance is properly administered.

It's essential to note that the use of restraint and the injection procedure should follow ethical guidelines and regulations for the humane treatment of animals in research. Researchers typically undergo training to perform these procedures with care and precision.
Keep in mind that the specific details of the technique may vary depending on the laboratory, the type of study, and the substances being injected. Always follow the appropriate ethical and regulatory guidelines when working with laboratory animals.

Summary

The injection tail technique in rodents involves administering substances into the bloodstream using the tail vein. A common method includes using a cone with a light source to enhance visibility of the vein.

The procedure typically involves restraining the rodent, visualizing the tail vein with the aid of a cone and light, and carefully injecting the substance into the identified vein.

Researchers follow ethical guidelines and regulations to ensure the humane treatment of animals in research.

Apparatus and Equipment

Protocol

Strengths and Limitations

Strengths

Limitations

Summary

1. The device uses a removable pin to both support the weight at the ideal height and to function as a release mechanism
2. The impounder is held in place using a horizontal pin which guides the weight and also prevents it from bouncing on impact
3. Retrieval of the weight is done using a rod magnet
4. The impounder must be placed perpendicular to the spinal cord to induce symmetrical injuries

References

1. Koozekanani SH, Vise WM, Hashemi RM, McGhee RB. (1976). Possible mechanisms for observed pathophysiological variability in experimental spinal cord injury by the method of Allen. J Neurosurg.  Apr; 44(4):429-34.
2. Ahdeah Pajoohesh-Ganji, Kimberly R. Byrnes, Gita Fatemi, Alan I. Faden. (2010). A combined scoring method to assess behavioral recovery after mouse spinal cord injury. Neurosci Res; 67(2): 117–125.
3. Akhtar AZ, Pippin JJ, Sandusky CB. (2008). Animal models in spinal cord injury: a review.Rev Neurosci; 19:47–60.
4. McEwen, M.L., Springer, J.E. (2006). Quantification of locomotor recovery following
5. spinal cord contusion in adult rats. J. Neurotrauma 23, 1632–1653.

Introduction

• Clear cone for full visualization of the muzzle.
• The Cone Masks can be installed onto the 68620 anesthesia operation platform(300*210*75mm) to secure the animal.
• The height of the mask can be adjusted to meet different experimental needs.
• A special semi-enclosed mask design coordinates with the evacuation apparatus to completely absorb and remove the waste gas outside the mask.

Introduction

Chemical And Physical Properties

Pharmacodynamics

Pharmacokinetics

Strengths & Limitations

• Produces rapid induction and recovery
• The depth of anesthesia can be easily and rapidly altered
• Non-explosive and non-flammable
• Less pungent than other agents
• Mask induction tolerated well in many species

• Relatively expensive
• Unstable in the presence of soda lime
• Breakdown products can cause renal injury

References

Chemical And Physical Properties

Introduction

Pharmacodynamics

Pharmacokinetics

Strengths & Limitations

• Produces rapid induction and recovery from anesthesia
• The depth of anesthesia can be easily and rapidly altered
• Non-explosive and non-flammable
• Non-irritant

• Produces moderate respiratory depression
• Produces moderate cardiovascular depression
• Pungent odor

References

*Flow regulator_Tube: 304 stainless steel material. Infusion_Syringe:Flat head for the injection of liquid.

Introduction

The implantable osmotic release pump is an inexpensive cutting-edge drug delivery system. You can use this small implantable infusion pump for preclinical pharmaceutical research in mice, rats, and other laboratory animals. The mini-pump delivers drugs, hormones, and other test compounds at continuous and controlled rates, for terms extending from one day to a month and a half, without the requirement for external interference. The implantable pumps utilize osmosis for continuous infusion of unrestrained laboratory animals.

In the previous three decades, drug delivery research has made critical headways because of the recent developments and innovations in the fields of pharmaceutical sciences including pharmacokinetics, pharmacodynamics, and biopharmaceutics. Low development costs and regulated drug delivery pushed the research. Hence a significant portion of the novel drug delivery frameworks is upgraded such that the drug dose and dosing interim are reduced thereby maintaining optimum therapeutic dose and efficacy. These innovative drug delivery systems have been designed to regulate drug release over an extended time frame. Furthermore, recent advances have been made to make the rate and extent of the drug release independent of physicochemical properties of drugs and excipients, and physiological factors like pH of the gastrointestinal tract, the presence of food, and nutritional health.

Osmotic pumps are the most promising procedure based systems for controlled drug delivery. The controlled drug delivery system is mediated by osmosis which can be characterized as the net movement of water molecules over a selectively permeable film driven by a difference in osmotic gradient over the layer. The difference in solute concentration over the membrane permits entry of water, however, rejects most solute particles. Osmogens create osmotic pressure to stimulate drug release from the pump.

Apparatus And Equipment

In cross-section, the implantable osmotic release pumps are composed of a drug core (reservoir), the diffusion agent, and the tissue layer (rate controller). Additionally, a flow moderator is inserted into the body of the diffusion pump. It is a kind of an implantable system within which you can load a solution or suspension contained in a cylindrical reservoir shaped from an artificial collapsible, impervious stuff wall (e.g., polyester) that is open to the external surroundings via a single orifice.

The main components of an implantable osmotic release pump include the drug compound (water-soluble or insoluble), osmotic agents (ionic compounds of inorganic salts), a stable semi-permeable membrane, plasticizers, flux regulators, wicking agents, pore forming agent, and coating solvent.

Our implantable osmotic release pump is available in three different sizes to offer the researchers a variety of dosing capacities, release rates, and duration. You can load it with 100 µl, 200 µl, and 2 ml of the test compound.

The implantable osmotic pump employs an osmotic gradient inside the lumen called the salt sleeve and the tissue condition in which the pump is implanted. The high osmolality of the salt sleeve makes water flow into the pump through a semipermeable film which encases the external surface of the pump. As the water enters the salt sleeve, it compresses the flexible reservoir, dislodging the test substance from the tube at a controlled rate. The rate of drug delivery is directly proportional to the osmotic pressure of the core. Since the delivery system cannot be refilled, these pumps are intended for single use only.

The rate of the drug delivery by our osmotic pump is controlled by the water penetrability from the pump’s external film. In this manner, the drug delivery is independent of the drug formulation and excipients. Drugs of different atomic arrangements, including ionized medications and macromolecules, can be administered persistently at controlled rates. The sub-atomic weight of a compound, or its physical and chemical properties, does not affect the rate of the drug delivery by the implantable osmotic release pumps.

Procedure

The osmotic pumps can be implanted subcutaneously or intraperitoneally depending on the size of the animal. For targeted drug delivery, a catheter can be attached to the osmotic pump to gain access to the tissues of interest. Subcutaneous implantation is technically the easiest and least intrusive procedure. Follow these steps for subcutaneous implantation:

1. Anesthetize the animal with ketamine or any other rodent anesthetizer.
2. Shave and wash the skin over the implantation site.
3. Make an incision adjacent to the implantation site. If the implantation is planned on the backside of the animal, then the mid-scapular incision is made.
4. To avoid excessive blood flow, place a hemostat onto the incision. Make an appropriate pocket for the pump by manipulating the subcutaneous tissues by hemostat. Keep in mind that the pocket should be large enough to allow the pump movement.
5. Insert the pump containing the drug into the pocket in the incision.
6. With the help of incision clips, close the wound.

The osmotic system can also be implanted in the peritoneal cavity of the larger animals. Follow the below-mentioned protocol for intraperitoneal injection:

1. After anesthetizing the animal, shave and wash the skin over the implantation site.
2. In the lower abdomen, make a 1 cm long midline skin incision.
3. Tent the peritoneal muscles and carefully incise the peritoneal wall keeping the bowel safe.
4. Insert the osmotic pump into the cavity.
5. Close the abdomen with the help of sutures and incision clips.

Keep in mind that the pumps should be explanted if the animals survive after active infusion. The pumps must be removed no later than the first half-life of the test compound.  The pump should be removed to measure the residual volume to confirm delivery, verify a drug’s stability, and assure the bioactivity of the test compound.

Applications

Implantable consistent infusion osmotic pumps have many applications in preclinical drug delivery improvement. The system empowers continuous and controlled dosing permitting the accomplishment of enduring state conditions and precise drug delivery. The system offers the researchers temporal and spatial control over the drug release. It circumvents poor-availability hurdles in the challenging therapeutic studies of hormones and growth factors. Also, the drugs with faster clearance rates and shorter half-lives require a consistent dosage that can be easily achieved by the implantable osmotic release pumps. Furthermore, it can assist the drug delivery of the compounds having a lower therapeutic index by continuous infusion while avoiding toxic concentrations. Moreover, the osmotic systems not only deliver the drugs with moderate solubility but also with extreme solubility.

Continuous infusion of the therapeutic compounds with the help of the implantable osmotic release pump assists the experimenters to establish parameters of the drugs with unknown pharmacokinetics. These pumps can also be used for a comparative study of the efficacy of different drugs administered through different routes. In addition to the drug efficacy analysis, these drug delivery systems can also be used for the targeted delivery of chemotherapeutic agents to the tumors with the help of an attached catheter. The osmotic release pumps can not only be used for chemotherapeutic preclinical studies but also to monitor cell proliferation to assess the carcinogenic potential of the cells. Also, these pumps improve bio-luminescence imaging studies by continuously delivering bio-luminescent substrates.

Applications of the osmotic release pump include the disciplines of oncology, stem cell research, gene transfection, gene silencing, neuroscience research, and preclinical pharmacological studies.

Strengths And Weaknesses

• The osmotic release pump ensures round-the-clock delivery of the test molecules at a controlled rate.
• Zero-order drug release after an initial tag is one of the significant advantages of the implantable osmotic release pump.
• The tool offers drug delivery independent of physiological factors like gastric health, pH, and hydrodynamic conditions.
• The drug release can be easily controlled by regulating the amount of water filled in the pump.
• The equipment permits continuous and regulated administration of short peptides and proteins.
• It offers a convenient method for the persistent dosing of laboratory animals.
• It minimizes unwanted experimental variables and makes sure reproducible, constant results.
• It eliminates the need for nighttime or weekend dosing.
• Its small size increases its popularity for use in mice and small rats.
• The osmotic drug delivery system allows for focused and targeted drug administration.
• It can be used for both in-vivo and in-vitro studies.
• It provides the biomedical researchers with cost-effective research tool.
• There may be a chance of dose dumping if the film coating is not correct.
• The size of the orifice is critical for controlled drug release.
• It may cause ulceration or irritation at the site of implantation.

Osmotic pumps are one of the novel pharmaceutical tools for controlled and consistent drug delivery. Osmotic drug delivery pumps commonly comprise a drug center containing osmogen that is covered with a semipermeable layer. This covering has at least one transportation ports through which the test compound or suspension of the medication is discharged after some time. Different endeavors are underway to make an effective osmotic drug delivery system like pulsatile delivery with an expandable hole, and a lipid osmotic pump containing a smaller capsules than the usual osmotic pump for continuous and extended-release.

References

1. Continuous Drug Infusion Model with an Implantable Osmotic Pump. Retrieved from: https://noblelifesci.com/uploads/file/Noble%20Life%20SciencesContinuous%20Infusion%20Model.pdf
2. Keraliya, R. A., Patel, C., Patel, P., Keraliya, V., Soni, T. G., Patel, R. C., & Patel., M. M. (2012). Osmotic Drug Delivery System as a Part of Modified Release Dosage Form. ISRN Pharmaceutics.
3. Mathur, M., & Mishra, R. A review on osmotic pump drug delivery system. International journal of pharmaceutical sciences and research.
4. Popesko, P., et al. (1992) A color atlas of small laboratory animals, Volume Two, Rat, Mouse & Hamster, London: Wolf Publishing Ltd.
5. Prescott LF. Novel Drug Delivery and Its Therapeutic application. West Susset, UK: John Wiley and Sons; 1989. The need for improved drug delivery in clinical practice; pp. 1–11.
6. Stepkowski, S. M., Tu, Y., Condon, T. P., Bennett, C. F. (1994) ‘Blocking of heart allograft rejection by intercellular adhesion molecule-1 antisense oligonucleotides alone or in combination with other immunosuppressive modalities‘, Journal of Immunology, 153, 5336-5346.
7. Tu, Y., Stepkowski, S. M., Chou, T.-C., Kahan, B. D. (1995) ‘The synergistic effects of cyclosporine, sirolimus, and brequinar on heart allograft survival in mice,’ Transplantation, 59(2), 177-183.

Rat Kit

Introduction

Apparatus and Equipment

Orthopedic Surgery Protocols

Tibial Fracture Protocol (Xiong Et Al., 2018):

• Place the subject in a supine position on a heated mat.
• Administer analgesia before and after surgical manipulation.
• Perform a skin incision along the medial aspect of the hind limb.
• Visualize the diaphysis and tibial plateau.
• Insert a stainless steel pin into the medullary cavity.
• Induce a fracture at the midshaft of the tibia using Bonn scissors.
• Stabilize the fracture site and close the incision.

Distraction Osteogenesis Protocol (Lybrand Et Al., 2015):

• Position the subject with the operative extremity up.
• Make a longitudinal incision along the femur.
• Expose and dissect muscle fibers attached to the femur.
• Place wires around the femur and secure them with a distraction osteogenesis device.
• Create a transverse osteotomy of the femoral shaft using a circular saw.
• Close the incision and fascia.

Marrow Ablation Protocol (Lybrand Et Al., 2015):

• Place the subject on its back with the operative leg flexed.
• Make an incision over the knee joint and expose the tibial medullary canal.
• Insert spinal needles of increasing diameter to ablate the marrow.
• Flush the bone marrow cavity with sterile saline.
• Close the incision.

Destabilization Of The Medial Meniscus Protocol:

• Position the subject in the supine position.
• Make a skin incision from the patella to the tibial plateau.
• Open the joint capsule and section the meniscotibial ligament.
• Close the wound.

Calvarial Defect Protocol (Spicer Et Al., 2012):

• Induce anesthesia and make an incision over the calvarium.
• Score and remove the calvarium to expose the dura.
• Clean the defect and place the implant material.
• Close the periosteum and skin.

Total Hip Replacement Arthroplasty (Powers Et Al., 1995):

• Make an incision and expose the lateral hip muscles.
• Excise the femoral head and ream the intramedullary cavity.
• Place the femoral component and fix the acetabular component.
• Reduce the hip and close the incision.

Standard Closed Femoral Fracture Protocol (Bonnarens And Einhorn, 1984):

• Flex the knee and make a small incision near the patella.
• Dislocate the patella to expose the femoral condyles.
• Insert a pin retrograde into the femoral canal.
• Cut and bury the pin beneath the bone.
• Fracture the femoral diaphysis by dropping weight.

Post-Operative Care And Pain Management

Applications

Evaluation of the effects of Nrf2 deficiency in fracture healing

Evaluation of the effects of a compound on bone healing during distraction osteogenesis

Evaluation of the effect of a protein on fracture repair

Evaluation of the effect of local vibration and pulsed electromagnetic field on bone fracture

Investigation of bone fracture healing in splenectomized rats

Investigation of conjugated linoleic acid in promoting fracture healing

Evaluation of whole-body vibrations in improving fracture healing in ovariectomy-induced osteoporosis rats

Advantages And Disadvantages Of Rodent Models

Summary

References

1. Bilgin HM, Çelik F, Gem M, Akpolat V, Yıldız İ, Ekinci A, Özerdem MS, Tunik S (2017). Effects of local vibration and pulsed electromagnetic field on bone fracture: A comparative study. Bioelectromagnetics. 38(5):339-348. doi: 10.1002/bem.22043.
2. Bonnarens F, Einhorn TA (1984). Production of a standard closed fracture in laboratory animal bone. J Orthop Res. 1984;2(1):97-101.
3. Bove SE, Laemont KD, Brooker RM, Osborn MN, Sanchez BM, Guzman RE, Hook KE, Juneau PL, Connor JR, Kilgore KS (2006). Surgically induced osteoarthritis in the rat results in the development of both osteoarthritis-like joint pain and secondary hyperalgesia. Osteoarthritis Cartilage. 14(10):1041-8.
4. Butezloff MM, Zamarioli A, Leoni GB, Sousa-Neto MD, Volpon JB (2015). Whole-body vibration improves fracture healing and bone quality in rats with ovariectomy-induced osteoporosis. Acta Cir Bras. 2015 Nov;30(11):727-35. doi: 10.1590/S0102-865020150110000002.
5. Gomes PS, Fernandes MH (2011). Rodent models in bone-related research: the relevance of calvarial defects in the assessment of bone regeneration strategies. Lab Anim. 45(1):14-24. doi: 10.1258/la.2010.010085.
6. Haffner-Luntzer M., Kovtun A., Rapp A.E, Ignatius A. (2016). Mouse Models in Bone Fracture Healing Research. Curr Mol Bio Rep. 2: 101. https://doi.org/10.1007/s40610-016-0037-3
7. Jung YJ, Kim R, Ham HJ, Park SI, Lee MY, Kim J, Hwang J, Park MS, Yoo SS, Maeng LS, Chang W, Chung YA (2015). Focused low-intensity pulsed ultrasound enhances bone regeneration in rat calvarial bone defect through enhancement of cell proliferation. Ultrasound Med Biol. 41(4):999-1007. doi: 10.1016/j.ultrasmedbio.2014.11.008.
8. Kogan NM, Melamed E, Wasserman E, Raphael B, Breuer A, Stok KS, Sondergaard R, Escudero AV et al., (2015) Cannabidiol, a Major Non-Psychotropic Cannabis Constituent Enhances Fracture Healing and Stimulates Lysyl Hydroxylase Activity in Osteoblasts. J Bone Miner Res. 30(10):1905-13. doi: 10.1002/jbmr.2513.
9. Lippross S, Beckmann R, Streubesand N, Ayub F, Tohidnezhad M, Campbell G, Kan YW, Horst F, Sönmez TT, Varoga D, Lichte P, Jahr H, Pufe T, Wruck CJ (2014). Nrf2 deficiency impairs fracture healing in mice. Calcif Tissue Int. 95(4):349-61. doi: 10.1007/s00223-014-9900-5.
10. Lybrand K, Bragdon B, Gerstenfeld L (2015). Mouse models of bone healing: fracture, marrow ablation, and distraction osteogenesis. Curr Protoc Mouse Biol. 5(1):35-49. doi: 10.1002/9780470942390.mo140161.
11. Moran CJ, Ramesh A, Brama PA, O’Byrne JM, O’Brien FJ, Levingstone TJ (2016). The benefits and limitations of animal models for translational research in cartilage repair. J Exp Orthop. 3(1):1. doi: 10.1186/s40634-015-0037-x.
12. Powers DL, Claassen B, Black J (1995). The rat as an animal model for total hip replacement arthroplasty. J Invest Surg. 8(5):349-62.
13. Shan Z, Luo ZP, Shen X, Chen L (2017). Promotion of fracture healing by conjugated linoleic acid in rats. J Orthop Surg (Hong Kong). 25(2):2309499017718910. doi: 10.1177/2309499017718910.
14. Spicer PP, Kretlow JD, Young S, Jansen JA, Kasper FK, Mikos AG (2012). Evaluation of bone regeneration using the rat critical size calvarial defect. Nat Protoc. 7(10):1918-29. doi: 10.1038/nprot.2012.113.
15. Stine KC, Wahl EC, Liu L, Skinner RA, Vanderschilden J, Bunn RC, Montgomery CO, Suva LJ, Aronson J, Becton DL, Nicholas RW, Swearingen CJ, Lumpkin CK Jr (2014). Cisplatin inhibits bone healing during distraction osteogenesis. J Orthop Res. 2014 Mar;32(3):464-70. doi: 10.1002/jor.22527
16. Wang D, Gilbert JR, Cray JJ Jr, Kubala AA, Shaw MA, Billiar TR, Cooper GM (2012). Accelerated calvarial healing in mice lacking Toll-like receptor 4. PLoS One. 7(10):e46945. doi: 10.1371/journal.pone.0046945.
17. Xiao W, Hu Z, Li T, Li J (2017). Bone fracture healing is delayed in splenectomic rats. Life Sci. 173:55-61. doi: 10.1016/j.lfs.2016.12.005.
18. Xie Y, Luo F, Xu W, Wang Z, Sun X, Xu M, Huang J, Zhang D, Tan Q, Chen B, Jiang W, Du X, Chen L (2017). FGFR3 deficient mice have accelerated fracture repair. Int J Biol Sci. 2017 Jul 18;13(8):1029-1037. doi: 10.7150/ijbs.19309.
19. Xiong, C., Zhang, Z., Baht, G. S., Terrando, N (2018). A Mouse Model of Orthopedic Surgery to Study Postoperative Cognitive Dysfunction and Tissue Regeneration. J. Vis. Exp. (132), e56701, doi:10.3791/56701.

Rat Kit

ConductScience's Digital Microscope is a professional digital microscope with a precise operation such as thousand brain positioning or surgical experiment microscopy. It can deliver real-time experimental pictures, high magnification, a clear image, it's economical and reliable, especially for teaching experiments.

Our 77001D released the eyes of the experimenter, greatly alleviating eye and cervical discomfort caused by long-term observation with a traditional microscope.

The focusing distance of the digital microscope is 180mm, which provides more space for the surgeon's surgical operation. At the same time, it can also perform vertical or oblique angle imaging according to the experimental conditions to find a better viewing angle.

Specifications

Camera parameters

● 3.5MP, 1/2.3" CMOS sensor

● 20x - 120x magnification

● The shutter can be controlled manually or remotely

● The focusing distance is 18cm

● Support manual fine-tuning focal length 8mm

● LED ring illuminator

Stage, Focusing, and Construction

● Pillar-type frame: stable base and metal clips

● Rod allows adjusting the height (working distance) and horizontal position

● Coarse-focusing adjustment with a knurled knob on the lens

Photo/Video Capture, Memory, & Ports

   Streams 1080p videos via a mini-HDMI port at 60 fps

● Captures 60 fps AVI videos: HD 1080p, 1920 x 1080

● Saves visual data to a built-in microSD card

● Captures interpolated JPEG photos with a resolution of up to 16.0 MP

● Mini-HDMI video out port for direct streaming to HDTV displays

The Morris water maze test is a widely used and reliable method for assessing spatial learning and memory in rodents, and it has been used in a wide range of neuroscience research, including studies on aging, brain injury, and neurological disorders.

Conduct Science offers the Morris Water Maze. Customizations are available upon request. The hidden platform is included in your order.

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Accessories

Gantry

Adjustable platform

Steel Frame with Casters

Floating Platform

Radial Arm Water Maze

Rondi Reig

Y Maze Inserts

Snowcone Insert

Plus Insert

T Insert

Multiple Beacons

Radial Arm Tread

Heater

Swim Channel: Full, Half Lenght

Release Device

Tower Inserts

Preswim Chamber

Round Arena

Introduction

The Morris Water Maze (MWM) is a water navigation task developed by Richard G. Morris in 1981 as a response to the Radial Arm Maze (RAM). The task was developed to show that local clues (auditory, visual or olfactory) were not necessary for spatial learning (Morris 1981).  The maze utilizes the averseness of water in motivating the subject to learn and find the escape platform rapidly. The purpose of the MWM is the same as that of the RAM, that is, the assessment of spatial learning. However, the tasks are achieved under different environments and motivations.

The Morris Water Maze eliminates choice-point decisions as seen in the RAM and other mazes that were modeled on nature’s burrow and trail systems. Further, the MWM forces the animal to use its own spatial localization system to guide it to the goal location. The maze since its development has been a popular water navigation task in behavioral neuroscience to study spatial learning and memory. The task enables accurate assessment of spatial working, and other learning and memory aspects, making it an effective tool in measuring the effects of neurocognitive disorders, lesions, and age. The task can also be extended to studying the development of possible neural treatments.

The original design of the apparatus created by Morris (1981) included a 1.3 m diameter large pool of height 0.6 m, which was filled with water to a height of 0.4 m above the base. Two distinguishable escape platforms 0.4 m and 0.39 m in height were used to create a visible platform (painted black) and an invisible platform (painted silvery-white), respectively. The water was also made opaque by the addition of powder, paint, or milk to it, such that the invisible platform was concealed. The apparatus can easily be adapted and modified using simple inserts to create hybrid mazes such as the Water T-Maze, Water RAM, Water Y-Maze and Water Plus Maze.

See our citation list

History

Origin

In 1981, Morris R. developed the Morris Water Maze, a simple and inexpensive navigational task that required continuous decision-making by the subject to escape from the water. The task involved a large pool filled with water, into which the rats were immersed and forced to swim to find one of the two, visible and invisible, escape platforms. Due to the limitations in local olfactory, auditory and visual clues, the subject is forced to depend on its own spatial learning system to reach the goal. Morris used 4 groups of rats and tested them in a setup wherein they were presented with an escape platform that was above or just below the surface and in a fixed or varied location. The experiment following the initial test, investigated the performance of the rats when they were placed in a novel starting position.

Further papers published by Morris evaluated hippocampal-dependent learning over several years (Morris 1981, Morris 1982, Morris 1984, Morris 1986). The maze also gained popularity when it was used by Ian Whishaw’s group in Canada (Kolb et al., 1982, Kolb et al., 1983).

Developments

Since its conception and development by Richard Morris at the University of St. Andrews in Scotland, the Morris Water Maze became a viral behavioral assay for spatial learning and memory. The MWM allowed testing of different variables of behavioral investigations, including pharmacological assessment and cerebral function, making it a popular choice for research in the domain of neurodegenerative and neuropsychiatric disorders, compound testing, and lesion models.

Since the initial papers, the maze has been used to study various disease models, including endocrine abnormalities, strokes, Alzheimer’s disease, other neurodegenerative diseases, and their effects on learning and memory (Brandeis et al., 1989).

Hamm et al. used the Morris Water Maze to investigate the generality of cognitive deficits observed after traumatic brain injury (TBI). The participants were subjected to three tests; the Passive avoidance test and constant-start versions of the MWM that did not require hippocampal processing and the standard MWM task that relied on hippocampal processing. In their findings, they were able to observe that fluid percussion TBI did not impair performance in the passive avoidance test and the constant-start tasks of the MWM.

Recent developments

In their investigation, Kishi et al. were able to observe that exercise improved cognitive decline as determined by the Morris Water Maze performance. Cognitive decline is seen as one of the critical organ damage of hypertension and studies have indicated that the decrease in BDNF in hippocampus causes the cognitive decline. Kishi’s investigation tested stroke-prone spontaneously hypertensive rats and was able to conclude that caloric restriction, in addition, to exercise up-regulated BDNF in the hippocampus leading to synergetic protection against cognitive decline.

Hosseini and colleagues (2017) investigated the effects of vitamin C during neonatal and juvenile growth on the learning and memory abilities of rats. The rats treated with 10-500 mg/kg of vitamin C showed reduced latency and travel distance and an increase in time spent in the target quadrant in the MWM task.

Apparatus & Equipment

The escape platforms are generally 8 cm in diameter and are matched to the color of the pool or water (in the case of using colored water). For trials that require the pool water to be made opaque, milk or non-toxic colorant is mixed with the water. Both intra-maze and extra-maze cues may be used to help orient the subject and assist them in remembering the location of the escape platform.

Automated scoring can be performed with the assistance of video and tracking software such as the Noldus Ethovision XT and ANY-Maze.

Modifications

Since its introduction, the Morris Water Maze has seen many variations in protocol and varying pool sizes. The maze has shown great success in a variety of investigatory processes and applications, including the testing of transgenic mice (D’Hooge and De Deyn 2001).

An “on-demand” procedure was described by Buresová et al. in their 1985 paper which replaced rigid platforms with collapsible platforms. The modification prevented a chance finding of the platform by the subject. A computerized system tracked the location of the subject and raised the platform when the subject had remained in the target area for a pre-determined time. The modification was further improved upon by Spooner et al. This modification allowed a highly focused search strategy.

Markowska et al. suggested a variation to the probe test that provided a more sensitive measure of spatial memory and proved more useful for repeated trials. Their modification suggested using a variable interval probe test wherein the platform is made available to the subject during the trial after a set interval. In comparison to the no-platform probe test, the variable-interval probe test proved to be more useful. Steele and Morris suggested another protocol variation in their paper published in 1999. Their suggestion involved moving the escape platform to a new location on each testing day, thus preventing the animal from knowing the location of the platform during the first trial. Eventually, once the animal had located the platform, it learns and remembers the location in one trial. The varying inter-trial interval can also assist in studying spatial memory.

In their 2007 investigation, Clark et al. modified the standard water maze by including spatial beacons in each of the four quadrants of the MWM. This modification causes the subjects to abandon a strict spatial strategy in favor of using the beacons to guide them to the escape platforms.

Other simple modifications include combining the Morris Water Maze with other behavior assessment mazes such as the Radial Arm Maze and T-Maze. RAM inserts add spatial complexity and combine the measures of the dry Radial Arm Maze with the rapid learning and aversive aspect of the Morris Water Maze. Similar in application to the water-based Radial Arm Maze, are the Water Star Maze and Water Plus Maze. Another popular dry behavioral assay that is combined with the Morris Water Maze is the Y-Maze which is modeled on the T-Maze. The Water T-Maze and Water Y-Maze allow the evaluation of spatial memory and learning combined with the fear of drowning.

Further, varying the type of platform, such as using a floating platform (also see Adjustable platforms), can also provide another measure for spatial learning and memory. Another modification of the MWM is using a snowcone insert to create a geometric cue within the arena. The insert is often used in conjunction with a balloon positioned above or near the escape platform to evaluate cue-based navigation preference in rodents. The Morris Water Maze is a highly adaptable behavioral task and is easy to modify.

Data

In the Morris Water Maze task, the data is recorded for the latency to find the platform and the time spent in the target quadrant. With the help of tracking and video recording software, the path traversed by the subject can be mapped, and the velocity of the subject can also be observed.

The data obtained from the Morris Water Maze is generally visualized by graphing the time it takes the animal to locate the escape platform, which is referred to as the latency time. This time is obtained by observing the animals in the maze via a video and tracking software or by analyzing the recorded experiments with a stopwatch.

The latency to find the platform decreases with repeated trials. The latency time can be easily graphed and compared across the sham control and disease model or intervention groups. Using graphs to compare the latency time between different disease or treatment groups, allows for easy visualization of the effect on spatial memory and learning. Animals in the control groups should show a significant decrease in latency time as they rapidly learn the location of the escape platform. Animals as disease models of neurodegenerative disorders, for example, should show a much slower learning curve with higher latency times, even after several trials. Generally, animal cohorts of 20-30 animals are sufficient to obtain p-values of <0.05 using ANOVA, t-tests, or Bonferroni’s post hoc tests (Harrison et al., 2009).

Traslational Data

The Morris Water Maze is a principal task in behavioral investigations and can also be extended to studies involving the understanding of cerebral functions and in the development of potential treatments.

Laczó et al. validated the translational potential of the Hidden Goal Task in Morris Water Maze in their investigation of disrupting potential scopolamine in rats and humans. Another similar study was conducted by Possin et al. to determine the validity of MWM in translational research for Alzheimer’s disease. Another study by Kishi et al. was able to observe that exercise with the addition of caloric restrictions was able to protect against cognitive decline in stroke-prone spontaneously hypertensive rats.

Virtual applications of the Morris Water Maze have also evolved with evolving technology. The Virtual Morris Water Maze enables testing of human subjects in a virtual reality version of the classic rodent maze. Astur and team were the first to use the Virtual MWM for evaluation of humans in 1998. Since then, investigations using human subjects in both analogous and homologous versions of the Morris Water Maze have occurred. The task can easily evaluate memory and learning performances of participants with different diseases, injuries, and neuropsychiatric disorders. Using a virtual environment is cost-effective and does not endanger the subjects. Since the maze environments are virtual, the possibility of creating environments to suit the needs of any investigation are endless.

Advantages

Double channels displayed on UI enables two animals’ surgeries and experiments independently at the same time.

The UI displays a working progress bar of heating pads, and rectal probes working status: Idle, Warning, Standby, and Working

Simultaneous monitoring Heating pads and Rectal Probes data

A corresponding heat insulating pad is underneath every heating pad to prevent temperature loss.

Strengths and Limitations

The Morris Water Maze has high reliability across a wide range of tank configurations and testing procedures. Further, it serves as an effective method for measuring hippocampal-dependent spatial learning and memory. The navigational task in comparison to other mazes is less laborious and time-consuming, despite the requirement of pre-training.

The task is also able to differentiate between spatial and non-spatial learning by using visible and hidden platforms. The different possible variations in protocols such as Discrimination learning protocol, Cued learning protocol and Latent learning protocol (Vorhees and Williams 2006) allow for measuring the different specificity of spatial learning and memory. Since the task can employ various modifications, it can test the brain function of many brain areas, not only the hippocampus, and this allows the test to evaluate more general cognitive function in addition to specific learning and memory functions.

Despite the presence of an escape platform, this test places a significant amount of stress on the animals. Initially, when the animals are placed in the pool, they are forced into a stressful situation with no obvious escape route. The act of being immersed in water and forced to swim can induce stress that may alter the outcomes of each repeated trial. It is imperative to ensure that the water temperature is appropriate to minimize the stress experienced by the animals. Mazes can be purchased with temperature control to help reduce stress caused by the water being too cold or too hot. It is also essential to understand that there may be variations in performances of different strains and performances may be dependent on the age, gender and other aspects of the subjects used. The ability of the subject to swim is also a crucial factor in obtaining correct results.

• The Morris water Maze was developed by Richard G. Morris in 1981 to prove that the subjects are capable of spatial learning without the need for local cues (visual, olfactory or auditory)
• The task utilizes the averseness of the water in motivating the subject to learn and find the escape platform rapidly.
• The task eliminates the need for choice points as seen with behavioral assays such as the Radial Arm Maze, T-Maze, etc
• The task provides measures of hippocampal-dependent learning, specifically of spatial and long-term spatial memory.
• The pool water can be colored using milk or colorants such as paint to make it opaque.
• The pool should be filled with just enough water such that the animal’s paws do not touch the floor of the pool nor it can climb over the wall.
• The water temperature should be optimally maintained, and the subject should be gently dried and placed under a heat lamp before returning to its housing to minimize the stress on the subject.
• Subjects with cognitive deficits show a decline in performance in finding the platform and tend to have a slower learning curve with higher escape latency.

Summary

• Developed by Richard G. Morris in 1981. Designed to demonstrate spatial learning without relying on local cues.
• Aversiveness of water used to motivate subjects to learn and find escape platform. Assess spatial learning, similar to the Radial Arm Maze (RAM), but under different conditions.
• Eliminates choice-point decisions seen in other mazes. Forces animals to use their spatial localization system.
• 1.3 m diameter pool, 0.6 m height, filled with opaque water. Two escape platforms: visible (painted black) and invisible (painted silvery-white).
• Popular in behavioral neuroscience for studying spatial learning and memory. Various modifications and hybrid mazes (Water T-Maze, Water RAM, etc.).
• Used in disease models (strokes, Alzheimer’s) and interventions. Different protocols and variations for improved sensitivity and memory assessment. Latency to find platform and time spent in target quadrant recorded.
• Path traversed and velocity observed using video and tracking software. High reliability, effective for measuring hippocampal-dependent spatial learning. Differentiates between spatial and non-spatial learning.
• Less laborious than other mazes, despite stress on animals. Stress reduction considerations: water temperature control, drying, and heat lamps. Performance variations based on strains, age, gender, and swimming ability of subjects.

References

Brandeis, R., Brandys, Y., & Yehuda, S. (1989). The use of the Morris Water Maze in the study of memory and learning. The International journal of neuroscience, 48(1-2), 29–69. https://doi.org/10.3109/00207458909002151

Buresová, O., Krekule, I., Zahálka, A., & Bures, J. (1985). On-demand platform improves accuracy of the Morris water maze procedure. Journal of neuroscience methods, 15(1), 63–72. https://doi.org/10.1016/0165-0270(85)90062-7

D'Hooge, R., & De Deyn, P. P. (2001). Applications of the Morris water maze in the study of learning and memory. Brain research. Brain research reviews, 36(1), 60–90. https://doi.org/10.1016/s0165-0173(01)00067-4

Hamm, R. J., Lyeth, B. G., Jenkins, L. W., O'Dell, D. M., & Pike, B. R. (1993). Selective cognitive impairment following traumatic brain injury in rats. Behavioural brain research, 59(1-2), 169–173. https://doi.org/10.1016/0166-4328(93)90164-l

Kishi, T., & Sunagawa, K. (2012). Exercise training plus calorie restriction causes synergistic protection against cognitive decline via up-regulation of BDNF in hippocampus of stroke-prone hypertensive rats. Annual International Conference of the IEEE Engineering in Medicine and Biology Society. IEEE Engineering in Medicine and Biology Society. Annual International Conference, 2012, 6764–6767. https://doi.org/10.1109/EMBC.2012.6347547

Markowska, A. L., Long, J. M., Johnson, C. T., & Olton, D. S. (1993). Variable-interval probe test as a tool for repeated measurements of spatial memory in the water maze. Behavioral neuroscience, 107(4), 627–632. https://doi.org/10.1037//0735-7044.107.4.627

Morris, R. G., Garrud, P., Rawlins, J. N., & O'Keefe, J. (1982). Place navigation impaired in rats with hippocampal lesions. Nature, 297(5868), 681–683. https://doi.org/10.1038/297681a0

Morris R. (1984). Developments of a water-maze procedure for studying spatial learning in the rat. Journal of neuroscience methods, 11(1), 47–60. https://doi.org/10.1016/0165-0270(84)90007-4

Specifications

Modifications

Goal Box

Available for all arms

Mouse: Cost $250

W 9cm x W 9cm x H 10cm

Rat: Cost $250

W 9cm x W 9cm x H 10cm

Light Cues

For Mice and Rats

Single Push buttom Manual Light: used for light cues & win experiments.

6 Arm Radial Maze

Available with same modifications: Doors, Goal boxes, Light cues, backlights

Waterproofing

For Mice and Rats

Plus Maze Insert

Goal Box

Y Maze Insert

Made to fit your RAM. Turns de 8arm radial into a Y

T Maze Insert

Made to Fit

Elevated Stand

Leg Stand

Made to fit.

Elevates your RAM.

3D Radial Arm Maze

Escape Hole RAM

Arm Lenght Variants

n Arm variants

Introduction

See our citation list

History

Origin

Developments

Recent Developments

Apparatus & Equipment

Radial Arm Maze - RAM

Advantages

Training Protocol

Modifications

Traslational Research

Strengths and Limitations

Summary

• The Radial Arm Maze is extensively used to test spatial learning and memory.
• RAM task asks animals to retrieve food rewards at the end of each of eight arms without visiting an arm more than once.
• Different groups have adapted this maze in order to collect data regarding working or reference memory, and there are many modifications available, both regarding experimental protocol and data analysis.
• Animals in control groups show rapid learning as they remember the location of the arms from which they have retrieved the food reward, while in comparison, animals with neurological lesions show a much slower learning curve with lower memory scores.

References

Cell options

Accesories

*Cell covers available for all Vertical and Horizontal Diffusion Tank Options.

Apparatus & Equipment

Conduct Science's diffusion cell apparatus offers six stirring with six transdermeal cups at 15ml cell capacity. It comes with a digital RPM indicator and speed controller, a water level indicator, and a digital temperature controller. The cell holders contain diffusion cells with stirring bars. The apparatus provides constant stirring to the solutions. It comprises a water heater that helps in attaining the desired temperature. The water circulation system is also provided with a water level sensor that protects the heating element from damage. A magnetic induction knob aids the heavy-duty stirrers to offer constant and non-stop stirring. The stirring range lies between 200 and 800rpm. Moreover, the stainless-steel body of the cell apparatus makes it corrosion-free. Sheet membrane is required to be cut to size for each diffusion cell.

FAQs

How many cell tank positions are included in the system? The standard model includes 6 pcs vertical cells (15ml).

Is it possible to choose a smaller cell tank for the set-up of the system? Yes. There are 10 ml vertical and 5m Horizontal cell tank options

Vertical and Horizontal Cell tanks can be used simultaneously in the system?

10 ml vertical diffusion tanks can be used with 15 ml vertical diffusion tanks together. The horizontal diffusion tank can't be used with the vertical diffusion tank at the same time.

Does the system include jacketed or unjacketed cell tanks? All, Vertical (10ml, 15ml) and horizontal (5ml) cell tank options are unjacked. The system includes a water circulation whet-slate.

How to load the polymeric membrane and tissue?
The cell cap is connected to the cell body through a cell clamp. This way the cell clamp is secured with a steel clip.

How to prepare the samples (polymeric membrane and tissue)?

The size of the sample should be larger than the outer diameter of the bottle mouth. The thickness should not exceed 2mm

Does the system include membranes? No. Clients can obtain them according to the experimental protocols, either locally or from other suppliers.

Six Stirrings

Water Heater

Digital RPM Indicator

Speed Controler

Water Level Indicator

Six Transdermal Cups

Cell Holders: Diffusion Cells + Stirring Bars

Constant Stirring of Solutions

Stainless-steel body of cell apparatus

Digital Temperature Controller

Introduction

The diffusion cell apparatus is used to measure in vitro release of drugs from creams, ointments, oils, and gels. Diffusion is the random movement of molecules across the concentration gradient, i.e., from high concentration to low concentration. In vitro diffusion is the passive diffusion of a solution molecular species from a donor chamber through a membrane into a ‘receptor fluid’ present in the recipient compartment.

The molecular species passing/diffusing through the membrane is called ‘permeant.' 'Permeation’ is defined as "the movement of the permeant through the tissue/membrane that incorporates the first membrane and then diffuses through the membrane."

Flux is defined as the amount of permeant crossing through per unit area of the membrane into the circulating system per unit time. In in vitro diffusion, the circulating system is referred to as the 'system' and expressed in mass, area, and time.

The amount of molecules passing through the membrane in the given time is known as accumulation and is defined in mass and area.

Diffusivity is a characteristic of permeant that measures its ease of penetration through a particular membrane and is expressed in area/time.

Permeability coefficient is described as the 'rate of penetration of the permeant per unit concentration,' expressed in units of distance/time.

Any permeant that permeates through a membrane diffuses into the receptor fluid and reaches a steady state of diffusion takes some time. This time is the ‘lag time.' The permeation rate across the membrane rises in the lag time. Once the sample reaches a steady state, the movement of permeant through the membrane becomes consistent. The attainment of steady-state depends on various factors, including membrane/tissue permeability, compound properties, and the receptor fluid flow rate.

The permeability barrier is a lipid barrier for human cells that depends on lipid type and organization. It should be noted that the permeability of tissue is not related to its thickness. For instance, different permeants exhibit different permeability over different sites on human skin.

Principle

The working of diffusion cell apparatus or Franz cell is based on 'in vitro diffusion.' It consists of two chambers, a donor compartment and a receptor compartment, separated by a membrane. The product to be tested is introduced through the donor compartment (the top chamber). The bottom chamber or the receptor compartment contains fluids from which samples to be analyzed are extracted at regular intervals. The sample collected determines the amount of "active" that permeates through the membrane at point time. The cell temperature constantly remains at 37oC. The receptor compartment of a diffusion cell or Franz cell has a fixed volume, and it allows the stirring of both receptor and donor chambers (Kelin et al., 2018).

A diffusion cell apparatus helps to evaluate compound uptake into the membrane and determines tissue/membrane permeability, concentration in the receptor compartment, and flux. When working with a highly permeable compound and a large receptor compartment, the reduction in the gradient builds up the compound in the receptor chamber. On the other hand, if the receptor chamber is small, the concentration gradient is reduced due to compound buildup. The reduction in concentration gradient ultimately slows down the compound flux resulting in non-sink conditions. However, if the compound has low permeability, the detection of the compound in a large volume receptor chamber can be problematic.

Membrane Types

Various types of membranes can be used for diffusion experiments. Some of these are given below:

• Human membranes: Healthy human tissues can be used for permeability experiments on a large scale. Some laboratories also use human cadavers' skin obtained from accredited tissue banks. However, there are some ethical issues with using healthy human tissues.

• Animals: Small and large animal tissues are used for diffusion experiments. Small animals like rodents are used for permeability experiments as they are inexpensive and easy to keep; however, they are morphologically different from human tissues and are relatively thin. Large animals like dogs, monkeys, pigs, etc., might also be used, as their tissue morphology is like that of humans; however, they are expensive.
• Synthetic/Polymeric Membranes:  Polymeric membranes are frequently used for in vitro release testing. They are synthetic, inert, and commercially available as cellulose, polysulfone, etc. Hydrophilic membranes with 0.45µm pore size are usually used for In vitro release tests (IVRT). Once set in the developing phase, the same membrane should be used for the entire experiment.
• Human Skin Equivalents (HSEs): These are synthetically engineered 3D human skin tissues prepared from a combination of human skin cells and extracellular matrix components under controlled conditions. Their main advantage is that they remain viable. However, they have higher permeability as compared to the tissues in vivo. Artificial membranes are beneficial as one can use them to predict the in vivo effect of compounds and formulations.

While selecting a suitable membrane, one must consider its source, integrity, time of excision, and shelf life. If it is obtained from an animal source, the scientist must know the health and age of the animal, and if the source is human, the researcher must consider the race, age, and gender of the human. If the permeability of the membrane/tissue is altered, it may affect the results of your experiment. Therefore, the experimenter must know beforehand whether the membrane was exposed to any chemicals. Skin tissue, however, may be refrigerated and used later without a change in permeability.

Donor Compounds

The donor compound can be prepared as liquid, gel, suspension, ointment, lotion, powder, or adhesive patches. One can determine permeant behavior and characteristics based on concentration and permeability.

1. Permeant Concentration

The permeant concentration depends on the aim of the study. An infinite dose is when the permeant does not deplete from the donor compound throughout the experiment and is used to test fundamental permeation behavior in the presence of permeability enhancers. On the contrary, a finite dose relates to the permeant amount that would be practical in actual usage, for example, a specific amount of drug to be administered to a patient. It aims to determine the amount of permeant required to be incorporated in a vehicle that will deliver the required amount of drug to the patient. Alternatively, the application of a compound that will temporarily remain on the tissue can be mimicked. In this case, the permeant is applied to the membrane for a short time and then removed. The researcher should then collect the samples for a given length of time following permeation.

2. Vehicle Formulation

The vehicle is the medium in which the permeant of interest is incorporated. Vehicle formulation is the “dose to be administered without the drug or active pharmaceutical agent (API).” Aqueous solutions and phosphate buffers serve as excellent vehicles for getting basic permeation data. Lipophilic compounds, however, have low solubility in aqueous solutions. Therefore, vehicles like water, alcohol, or propylene glycol should be used while working with them. Inert materials may alter the flux of compounds.

Moreover, the potency of enhancers may be altered by the nature of the vehicle. A selection buffer controls the degree of ionization; therefore, adjusting pH can completely dissociate the permeant into ions. Furthermore, vehicle components may also be modified by thermodynamic activity.

3. Permeant Detection

Spectroscopy and chromatography techniques are used for permeant detection.

4. Other Considerations

A group of five to six replicate doses should be administered to avoid errors resulting from the biological variability of skin. A minimum of three parallel cells should be used for testing each formulation.

Receptor Media

The receptor solution is selected based on the nature of the permeant. An ideal receptor compound should mimic in vivo situations. The accumulation in the receptor compartment is minimized by using flow-thru systems, and aqueous receptor fluid is sufficient for this purpose. On the other hand, it raises a concern for static cells where the permeant is not continually cleared. The solubility of the compound is a major concern in this regard. It should be such that it is in its desired form in the donor media and delivered in a certain amount to the receptor compound. Hydrophilic and lipophilic permeants show the best results with the aqueous receptors.

Similarly, PBS is used for ionizable permeants. If the permeant is un-ionizable, a solubilizer should be added to the receptor compound. Moreover, permeants with low aqueous solubility, like lipophilic permeants, require additional solubilizing agents such as surfactants or organic solvents. However, these agents might alter the membrane permeability and damage or alter the tissue/membrane. Therefore, one should use the minimum possible amount of these agents.

Sampling

The method of sampling depends on your research question. One can measure the amount of compound passing through the membrane at short intervals or the total amount that passes over a long-time duration. One can also determine flux, permeability constant (Kp), accumulation, etc., using a diffusion cell apparatus.

The researchers can use radiolabeled or unlabeled compounds for measuring flux and permeability. The results are reported as one of the two types of accumulation:

1. The total amount of the compound in the collected sample per time interval (irrespective of volume).
2. The compound concentration.

Radiolabeled compounds can be detected in very small amounts in large volumes of receptor media; however, the results are accurate and robust for such compounds. On the other hand, non-radiolabeled compounds used in permeability research can be detected using HPLC and ELISA methods.

Calculating flux and Permeability Constant

Flux is the amount of permeant passing through the membrane in unit time. It is measured in 'Joules' (J), and its formula is given by: J=Q/ (A x t). In the given formula, Q is the amount of permeant crossing the membrane in time t, and A is the membrane area usually given in cm2.

Steady State Flux is the amount of permeant crossing the membrane at a constant rate. This state is achieved after the lag phase as the amount of permeant continues to increase. When you measure the amount at regular short intervals at this stage, no significant difference in the values is observed. The units of steady-state flux (Jss) are quantity/(cm2*h). the formula for calculating steady-state flux is J = dQ/dt*A, where it is the permeation time, and A is the membrane area in cm2.

Permeability constant (Kp) can be calculated by using the formula Kp= Q/ [A*t*(Co-Ci)]. It is measured when an infinite dose of permeant is applied to the membrane. In the given formula, Q is the quantity of the permeant passing through the membrane in time t, A is the area of the membrane (cm2), Co is the concentration of the compound on the outer side (donor chamber), and Ci is the compound concentration on the inner side (receptor chamber).

The apparent permeability of the solution can be calculated by normalizing ‘J’ over drug concentration in the donor compartment Co, i.e., Papp = J/ Co.

Applications

In-vitro Release Testing

In vitro release testing (IVRT) employs diffusion cell apparatus to evaluate topical dosage form’s performance and determine the physiochemical properties of a product. Kelin et al. (2018) examined the critical parameters of diffusion cell apparatus and developed methods using this apparatus. IVRT has been used for the past 50 years to understand the release rates of products like hydrocortisone and betamethasone dipropionate. It allows the optimization of a drug formulation to make it ideal for in vivo use. This method also helps establish novel drugs' sameness by making minor changes in drug formulations during clinical testing. These minor changes ensure the sameness of the drug between clinical development stages. They stated that VDC (vertical diffusion cell) method could be validated by assessing attributes like drug solubility and stability in receptor media, binding the drug to the membrane, release rates precision, discrimination sensitivity, selectivity, and specificity, the sturdiness concerning critical parameters, and many others. Moreover, several parameters (like temperature, diffusion cell dimensions, sampling intervals, receptor fluid, membrane, and the amount of the permeant applied to the membrane) are tightly controlled to optimize the permeation experiments using VDC (Klein et al., 2018).

Pre-formulation Characterization of Semi-Solid Dosage Forms

Salamanca et al. (2018) evaluated the ketoprofen (KTP) releasing profiles of two drug formulations, i.e., a gel and a suspension. Ketoprofen is a widely used drug for treating rheumatoid arthritis, spondylitis, osteoarthritis, and gout. The researchers prepared KTP suspension by mixing 12.5g of ketoprofen with 2 ml of glycerin and 500ml of ultra-pure water and used 2.5g of sodium alginate as a suspensor agent. On the contrary, they similarly prepared KTP gel with an additional 2.5g of calcium chloride. The experiment was conducted in three individual Franz cells with 125ml volume of the acceptor compartment and membrane area of 2.5cm2. The amount of permeant in the donor chamber was set at 5g. the samples were stirred at a speed of 480rpm for 24 hours. They used triplicates of each sample and maintained the temperature throughout the experiment at 37oC. Samples were evaluated at regular short intervals, and the scientists measured their releasing efficiency in terms of mass flux. Two aqueous media were used as acceptor phases imitating the physiological conditions corresponding to buffers with pH 5.6 and 7.14 at 0.15M.

Furthermore, two types of membranes were used, i.e., regenerated cellulose membrane and a transdermal diffusion test. Mass flux was calculated using the formula, and kinetic study of ketoprofen permeation was done using statistics. The researchers concluded that the gel formulation had higher permeation efficiencies as compared to the suspension.  They also reported that evaluating the performance of pharmaceutical products at the pre-formulation stage becomes easier if one uses artificially engineered membranes.

Evaluation of cinnamon oil and chlorhexidine permeation through C. Albicans biofilm

Satthanakul et al. (2020) presented an in vitro method to analyze the permeation of cinnamon oil and chlorhexidine (CHX) through Candida albicans biofilm using diffusion cell apparatus. C. Albicans biofilms contain extracellular polymeric substances (EPS) that act as a barrier to antifungal agents like the mentioned products and prevent them from reaching the target site of the yeast cells. They prepared cinnamon oil and CHX solutions and grew C. Albicans biofilms on membrane filters by inoculating 50µL of C. Albicans on the UV-exposed sterilized filters. They allowed different combinations of CHX and cinnamon oil to pass through the biofilm. To achieve this, they filled the Franz cell with 5.3ml of the sample solution. The sample solution contained 0.5% w/v CHX and 8µL/ml cinnamon oil or 0.2% w/v CHX and 8µl/ml of cinnamon oil. The researchers maintained the volume by adding the solution to the sample port. The sample was stirred at 50rpm for homogenization, and the temperature was maintained at 37oC to mimic the oral cavity environment. The C. Albicans biofilm cultivated at 25mm membrane filters was mounted into a sample holder, and smaller-sized filters were placed on the fixed biofilm. The paper disks moistened with 10µl distilled water were placed on the membrane filters to prevent capillary action. The sample holder was covered with the cap, and the Franz cell was assembled. The experimenters set the temperature at 37oC via a temperature controller and collected paper disks at regular intervals. The disks were then placed on Sabouraud dextrose plates containing C. Albicans. Zones of inhibition were measured after 48 hours of incubation at 37oC. They compared these inhibition zones with the zones formed by cellulose membrane filters lacking biofilms. The sessile biofilms were tested for viability, and the results were subjected to statistical analysis. They concluded that sessile C. Albicans biofilms tested with 0.5% w/v CHX and 8µl/ml of cinnamon oil resulted in a “4 hours log reduction”.

Biomimetic Artificial Membrane Permeability Assay

Permeability of oral drugs through the lipid bilayer of the intestinal epithelial cells is an important parameter to measure the extent of drug absorption. Teixeira et al. (2020) validated a biomimetic artificial membrane permeability assay to assess the permeation profile of six drugs, namely "acyclovir, cimetidine, diclofenac, ibuprofen, piroxicam, and trimethoprim," along with eight model drugs from the Biopharmaceutic Classification System (BCS) using a diffusion cell apparatus. They impregnated the membrane supported with a solution containing a mixture of phospholipids and weighed it on a microanalytical scale. These impregnated membranes were then refrigerated to protect them. They positioned the impregnated membranes between the donor and the recipient compartments for permeation studies, each containing PBS, and stirred the receptor media at 500rpm.

Moreover, they maintained the temperature at 37oC and added triplicates of each drug in the donor compartment. The donor compartment contained a fixed concentration of the fluid, i.e., 10mg/ml and 1ml of saturated drug solution, and was capped to avoid evaporation. They ran the experiment under infinite dose conditions except for these drugs: caffeine, atenolol, metoprolol, naproxen, propranolol, and ranitidine. Drug solubility for each compound was measured. Then the researchers did permeability calculations using appropriate formulas. They classified 10 out of 17 drugs as low permeability drugs, and out of these 10, the permeability of only three drugs was underestimated as per BCS.

Diffusion Cell Apparatus

Strengths and Limitations

Ease of Reproducibility. Limited tissue handling. Minimal Sample Collection Requirements.

Low Drug Sample Requirement. Versatility in Permeation Studies:is well-suited for various permeation studies, allowing researchers to investigate the diffusion of substances across membranes under controlled conditions.

Limited Sensitivity with Radiolabeled Compounds. Challenges in Handling Radiolabeled Substances.

Precautions

• If you are not a pro at permeation experiments, run some simple initial experiments to avoid personal shortcomings.
• While preparing the membrane/tissue, do not overstretch the membrane or damage it by perforation.
• The specimen must be large enough that an edge of the tissue can be safely fixed in the diffusion cell around the orifice.
• The tissue should not protrude into the receptor compartment as it could block the receptor fluid flow.
• Freeze the membrane at -80oC if it is to be used later.
• Be precise with sample preparation, pipetting, and sampling time. The addition of solubilizers or pH changes can alter the membrane permeability, which can affect the results.
• Ensure that the room temperature and skin temperature remain consistent throughout the experiments as temperature variations may disrupt the experimental results. Report the temperature while writing methodologies.

Summary

• The diffusion cell apparatus is used to measure in vitro release of drugs from creams, ointments, oils, and gels.
• Permeant is the compound passing through a membrane, and permeation is the movement of the permeant through the tissue/membrane that incorporates the first membrane and then diffuses through the membrane.
• Flux is the amount of permeant crossing through per unit area of the membrane into the circulating system per unit time.
• Permeability coefficient is described as the 'rate of penetration of the permeant per unit concentration.'
• Human skin tissues, small and large animals, and artificially engineered 3D Human Skin Equivalents (HSEs) can be used as membranes in permeation studies.
• The donor compound can be prepared as liquid, gel, suspension, ointment, lotion, powder, or adhesive patches. A vehicle is sometimes used to incorporate the permeant of interest.
• Receptor media are selected based on the nature of the permeant.
• Diffusion cell apparatus is used for in vitro release testing (IVRT), pre-formulation characterization of semi-solid dosage forms, evaluation of permeation of drugs through biofilms, and biomimetic artificial membrane permeability assay.

References

Klein, R. R., Heckart, J. L., & Thakker, K. D. (2018). In vitro release testing methodology and variability with the vertical diffusion cell (VDC). Dissolution Technol, 25(3), 52-61.

Salamanca, C. H., Barrera-Ocampo, A., Lasso, J. C., Camacho, N., & Yarce, C. J. (2018). Franz diffusion cell approach for pre-formulation characterisation of ketoprofen semi-solid dosage forms.  Pharmaceutics, 10(3), 148.

Teixeira, L. de Souza., Vila Chagas, T., Alonso, A., Gonzalez-Alvarez, I., Bermejo, M., Polli, J., & Rezende, K. R. (2020). Biomimetic Artificial Membrane Permeability Assay over Franz Cell Apparatus Using BCS Model Drugs.  Pharmaceutics, 12(10), 988.

Satthanakul, P., Taweechaisupapong, S., Luengpailin, S., & Khunkitti, W. (2020). In vitro method for studying the penetration of cinnamon oil and chlorhexidine through Candida albicans biofilm using Franz diffusion apparatus.  Songklanakarin Journal of Science & Technology, 42(2).1

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Introduction

This 3-chambered device is a fantastic contraption for researchers studying socialization who require an apparatus in which variables may be altered to change the premises of the experiment. The design permits socialization but disallows aggravated socialization so that auspicious and accurate data may be collected. Measurable factors include transitions between chambers, time spent in direct contact, and unique behavioral variables such as jumping and grooming. Accouterments for this product include floor cues, stainless-steel grids or perforated stainless-steel, to forge an aversive stimulus, and removable doors to establish biased and unbiased conditioned place preference testing.

• The Three-Chamber Sociability apparatus is a novel apparatus developed for studying social deficits as seen in neuropsychiatric disorders and neurodevelopment disorders.
• The apparatus is divided into three chambers using dividers that have sliding doors.
• Social deficits are observed in the apparatus by allowing the test subject to choose between chambers containing a novel live stimulus vs. empty chamber and familiar live stimulus vs. unfamiliar live stimulus in the choice chambers.
• Photoelectric sensor beams in an automated Three-Chamber Sociability apparatus allow tracking entries and exits to and from the chambers.
• The acrylic cage used to hold the live stimulus allows for sensory interactions while limiting aggressive behaviors.
• The Three-Chamber Sociability test can be coupled with other tests such as Partition tests, Resident-Intruder tests, Reciprocal Interactions test, etc. to understand the social behavior of the test subject further.

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Read More about this maze

• The Active Place Avoidance Arena is used to assess spatial navigation and memory in rodents.
• The rotating arena used in the Active Place Avoidance task eliminates the use of simple associations and forces the subject to keep moving to avoid the shock area.
• The shock sector of the arena is defined by distal room cues.
• The dry-arena makes Active Place Avoidance task suitable for evaluating spatial navigation performances of weanling rodents.
• Active Place Avoidance can be used as an effective tool for investigating the effects of brain lesions, pharmacological manipulations, and diseases on spatial learning and memory.

Introduction and Principle 

• Properties of the optogenetic tool like ionic current, nature of transported ions, current amplitude, and much more.
• Ability to specifically target desired cell type and deliver light to the area of interest in a precise manner (Ferenczi et al., 2019).

Apparatus and Equipment

Applications

• Activation and Inhibition of Neural Activity

• Neural Circuit Interrogation

• Bidirectional Neural Activity Modulation

Strengths and Limitations

Summary

• Optogenetic stimulation kit combines genetic and optical techniques for neuromodulation experiments in animal species via photostimulation.
• Edward Boyden and Karl Deisseroth developed the technique of optogenetics in 2005.
• The kit works on the principle that when a specific wavelength of light is incident on neural cells, it stimulates light-sensitive proteins in these cells, called opsins. These opsins hyperpolarize or depolarize the cell membrane, thereby exciting or inhibiting cell responses.
• The equipment is frequently employed for bidirectional neuronal activity modulation, neural activity interrogation, and exciting or inhibiting neurons within a millisecond.
• Optogenetic stimulation is more specific than any other neural stimulation method.

References 

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