Description

Mouse Kit

Rat Kit

Introduction

Microsurgery is performed under magnification employing advanced diploscopes, specialized precision tools, and various operating procedures. The microsurgical methods are primarily used for tissue transplantation and reattachment of the amputated body parts of the rodents.

The introduction of the operating microscope made microsurgical procedures more comfortable, and with it, microsurgery involving tissue transplantation began. In the 1960s, the microsurgical techniques gained popularity as the rabbit’s ear was replanted using a microsurgical procedure, which was a remarkable achievement in the discipline of microsurgery since the vessels anastomosed were small. The success of the procedure during the 1960s further strengthened the microsurgical composite tissue transfer techniques in the 1970s. During the next decade, autologous tissue transplantation was introduced. The success of the procedures over the years made microsurgery a significant procedure in rodent surgery. Shorter breeding cycles, faster regeneration, lower costs, and easier handling make rodents ideal for microsurgery research.

Preoperative Set-Up and Anesthesia Induction

Physically examine the animals before starting the surgical procedure. Animals should be monitored for nutritional status, quality of fur (thinning, dirty), and behavior (movements of the limbs and trunk, abnormal gait, rigid walking, and a flat abdomen as a sign of pain). Also, inspect the natural orifices for discharge from the nose, increased salivation, and impurities around the anus and genitals, and observe the condition of the eyes. It is also essential to monitor the breathing pattern because non-manifesting subclinical pulmonary diseases can lead to severe respiratory failure in general anesthesia with subsequent death of the animal.

Pre-anesthetic medications can be applied to prevent bradycardia and suppress bronchoconstriction. Rat’s liver produces atropine esterase, which may resist the atropine effect; therefore repeated injections might be required. Anesthesia is usually induced in the subject with the help of anesthetic systems using a face mask or an anesthetic chamber. Select the appropriate dose and duration of the anesthetic agent depending on the weight of the animal. Once the anesthesia is induced, assess the depth of anesthesia using the toe pinch test. Monitor the subjects for physiological parameters throughout the surgical procedures to ensure that the anesthesia is effective.

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.

Note: The distance between the vessel ends must be 1mm.

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.

Note: The artery anastomosis is more comfortable than the vein as the fine venous wall, and the lumen may collapse during the suturing procedure.

Interrupted Suture

For the end-to-end interrupted suture technique, clamp the vessels by placing three main sutures and dividing the circumference of the vessels into three equal 120° segments.

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

The Cuff technique is a more straightforward method of microvascular anastomosis. The cuffing technique employs polyethylene cuffs that are inserted into the donor and recipient vessels. The technique is becoming an alternative method to standard vascular anastomosis for many transplantation models in rats in end-to-end vascular anastomosis.

This method is commonly performed for large rat vessels; however, the method can be modified for biliary anastomosis after liver transplantation. Being inexpensive, comfortable, and fast the cuff technique is preferred over other anastomotic procedures.

Polyethylene or the angio-catheter cuff is used for vascular anastomosis. The cuffs are available in various sizes and are chosen depending on the diameter of the cuffed vessels. After inserting the angio-guidewire or other metal rods into the catheter, create a 1.5 mm long flap and cut the 1.5 mm-long body of the cuff. The flap helps to hold the cuff with a clamp during the implantation.

The basic procedure of the cuff technique involves the following steps:

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

Follow the recipient preparation steps mentioned above with the only difference in the inferior vena cava preparation. Do not anastomose the inferior vena cava and do not clean or ligate the branches of the inferior vena cava.

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

Orotracheally intubate and mechanically ventilate all the animals, using the same respirator and ventilation management as for donor animals. With the recipient animal positioned for a left thoracotomy, shave the left lateral thorax and clean it with a 75% alcohol solution.

Triple Axis Stabilizer

An aluminum plate probed with an L-shaped 2 mm steel wire serves as a base for a 15 cm long steel cylinder. To allow vertical movement in the cylinder, tap the cylinder on the side. Mount a mosquito clamp on top and anchor it with an articulated joint. Intraoperatively, attach an aneurysm clip clamping the cuffed vessels and the recipient bronchus during anastomosis to the mosquito clamp. This construction allows for precise longitudinal movements of the clip on a vertical and horizontal axis as well as rotation on a vertical axis.

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.

Remove the transplanted rat from the ventilator and place the animal under a heat lamp until fully awake.

Orthotopic Lung Transplantation Method

Animal Preparation

It is recommended to use heavier and bigger animal recipients. Use rats weighing between 250 and 300g as donors and rats weighing between 300 and 350g as recipient animals, respectively. Almost every recipient animal develops a mild pleural effusion within the first postoperative days. The experimental setting should be favorable for animal survival and be giving a gentle pleural effusion place to expand without compressing other major thoracic organs.

Anesthesia

Anesthetize the animals.

Intubation

Intubate the animals using a 14G intravenous catheter and ventilate mechanically with a small animal ventilator. Transillumination of the neck facilitates quick intubation. Perform all transplantations using a binocular surgery microscope. Place donor animals in a supine position under the operating microscope; place the recipient animals in a right lateral position to allow proper access to the complete left hemithorax.

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

Recover the animals on flat paper bedding (sterile paper towels, etc.) rather than standard animal husbandry bedding. Keep the animals warm as warmth may aid in speedy recovery. Place the recovery cage half-on a heating pad so that animals can choose their preferred temperature as they recover from the anesthesia. Do not return the animals or cages to the holding area until all the animals appear healthy. The animals which underwent surgery must have regained the ability to move in the cage freely. It is essential to monitor the animals post-operatively for unexpected signs of illness. The animals lose a small amount of weight after surgery, but proper analgesia and provision of food may help to regain weight quickly.

Monitor the general condition of the animal for five to seven days after surgery. Animals should be bright, alert, and active post-operatively. The animals should generally be interacting with the cage mates, eating and drinking, and able to achieve standard species-specific postures. Depression, anorexia, or sluggishness indicate abnormal behavior. Consider the possibility of untreated pain or infection. Food/fluid intake is also crucial to recovery. There may be some drop-off in consumption after surgery. Easier access to food and water may aid in mitigating food drop-off. Daily provision of wetted food in the bottom of the cage may also encourage animals to eat. Signs of infection include inflammation, redness, swelling, discharge, pain, anxiety, or the opening of the incision. Wound dehiscence should be dealt with by re-suturing of the wound under anesthesia. If repeated re-suturing fails, allow the wound to heal by secondary intention using an antibiotic ointment.

Applications

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

Oncological surgical techniques and radiotherapy are valuable tools to fight cancer. Regional lymph node dissections, widely used procedures, seem to progress as they result in lesser morbidity and better recovery; they are the most common cause of secondary lymphedema in the industrialized world. Lower limb lymphedema, gynecological malignancies, or upper limb lymphedema, secondary cases to breast cancer treatment, varying from mild to severe benefit from lymphedema surgery. Relatively new, microsurgical techniques are becoming the backbone of surgical lymphedema treatment. The model enables anastomosis of lymphatic structures and numerous available small-caliber veins around the neck. The lymphatic venous anastomoses model created by the microsurgical techniques in the rat provided the researchers with great insight into the surgical treatment of lymphedema.

Fallopian Tube Anastomosis (2015)

The microsurgical fallopian tube anastomosis technique is used to restore fertility in women who underwent tubal sterilization or excision of an occluded or diseased portion of the tube. The microsurgical fallopian tube anastomosis procedure restores fertility with excellent results and allows to avoid disadvantages associated with other popular treatment options including in vitro fertilization. The rat uterus possesses unique characteristics making it analogous to the isthmic portion of the human fallopian tube even though the rat’s fallopian tube is highly convoluted and significantly smaller when compared with humans. Female rats possess a duplex uterus consisting of two tube-shaped horns extending upwards toward the kidneys. Each uterine horn has a uniform caliber similar to that of the human oviduct, a thick muscular layer, and mucosa that is not folded abundantly and does not tend to prolapse. The outer serosal layer receives a vascular supply from the broad ligament, which anchors the horn to the dorsal body wall and has a structure highly analogous to the human mesosalpinx. The rat fallopian tube anastomosis provides an excellent model for research and microsurgical procedures.

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

Reconstructive surgery procedures involving free-tissue transfer are predominantly used in plastic surgery. It was researched that the state of microcirculation and its reaction to changeable conditions plays an important role in these procedures. In the field of ongoing research on microcirculation, various in vitro and in vivo experimental animal models are devised to assess microcirculatory structure, pathophysiology, and hemodynamics. Unquestionably, experimental research employing microcirculation models has considerably contributed to advances in reconstructive and plastic surgery and has improved postoperative prognosis.

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

Small bowel rat transplantation (SBT) is a cumbersome, time-consuming, and technically demanding procedure with high postoperative mortality in the first seven postoperative days. The small intestine transplantation procedure is either heterotopic or orthotopic with the portal or systematic venous drainage. The crucial factor for animal survival is the time of vascular anastomosis (manipulation time). A threshold of manipulation time of less than 45 minutes is recommended. Microsurgical procedure for small intestine transplantation has significantly reduced the manipulation time thereby increasing the animal survival post-operatively. Microsurgery has provided researchers with a straightforward and more comfortable small intestine transplantation procedure.

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

The microsurgical groin skin flap model in the rat is accepted widely, as the experimental model offers an inexpensive, practical, and valid instrument and techniques to practice microvascular anastomosis as well as to investigate numerous research questions. The modified rat groin flap employing the inguinal fat pad as an obstacle to minimize the effects of the bed on the skin of the groin flap has further developed the method of the groin skin flap. The standard model of the groin flap and its modifications describe the experimenters about flap design and vascular anatomy. The microsurgical groin skin flap model ensures 100% animal survival for rats post-operatively. The model is not only reliable and reproducible for practicing end-to-end and end-to-side microvascular anastomosis but also a time-saving, less technically demanding, economical, and useful tool to explore numerous research questions that extend from the underlying mechanisms of flap survival to the development of a further surgical design.

Precautions

As small rodents possess high metabolic activity, do not exceed pre-anesthetic fasting beyond 2 hours. Extended periods of food deprivation can lead to disturbances in balance, metabolic acidosis, and hypoglycemia. During prolonged fasting, essential intestinal flora dies, which may result in the resorption of endotoxin. Water must never be restricted during the pre-anesthetic period. For more comfortable handling the organs in the abdominal cavity, liquid food can be given in place of solid food approximately 8–12 hours before surgery.

Animal handling should be calm and gentle to avoid the intense release of stress which may cause tachyarrhythmia with subsequent cardiac arrest during the general anesthesia. Animal strain and breed must be selected depending on the requirements of the investigation since the subject’s strain affect the experimental results. Also, consider the age and gender of the subject. For pre-operative preparations, the surgical area should be separate from the main surgical site. Thoroughly clean all the equipment and the instruments before starting the surgical procedures. Ensure that the animal is properly anesthetized before beginning the surgery. Avoid damaging the surrounding tissues and muscles during surgical operations.

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

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.
Gurunluoglu, R., & Siemionow, M. Z. (2015). The Microsurgical Groin Skin Flap Rat Model. In Plastic and reconstructive surgery (pp. 53-62). Chicago: Springer.
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.
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.
Kwiecien, G. J. (2015). Fallopian Tube Anastomosis. In M. Z. Seimionow, Plastic and reconstructive surgery (pp. 39-43). Chicago: Springer.
Yazici, I., & Siemionow, M. Z. (2015). Lymphaticovenous Anastomosis Model in Rat. In Plastic and reconstructive surgery (pp. 33-38). Chicago: Springer.
Yu, H., Sagi, A., Ferder, M., & Strauch, B. (1986). A simplified technique for end-to-end microanastomosis. J Reconstr Microsurg, 2, 191–194.

CLASSIC

SPOTLIGHT

AUTOMATED

OPERANT

PROTOCOLS

ACTIVITY

PAIN & HEAT

ACCESORIES

DISCOVER

VIDEO TRACKING

SPECIES

Microsurgery Kit

You might also like

Frequently bought together

Microsurgery Kit

Description

Mouse Kit

Rat Kit

Introduction

Preoperative Set-Up and Anesthesia Induction

Microsurgery Protocols

Preparation of the Vessel

End-to-End Anastomosis

Continuous Suture

Interrupted Suture

End-to-Side Anastomosis

Continuous Suture

Interrupted Suture

Cuff Technique for Vascular Anastomosis

Organ Transplantation

Kidney Graft Procurement Protocol

Kidney Graft Transplantation Protocol

Microsurgical Implantation of the Rodent’s Heart Protocol

Pre-surgery Preparations

Recipient’s Preparation

Heart Graft Procurement Protocol

Heart Transplantation Method

Heterotopic Abdominal Heart and Lung Transplantation Protocol

Recipient Preparation

Heart and Lung Grafts Procurement

Heart and Lung Transplantation Protocol

Orthotopic Lung Transplantation Procedure

Triple Axis Stabilizer

Operating Procedure

Orthotopic Lung Transplantation Method

Animal Preparation

Anesthesia

Intubation

Lung Graft Procurement

Lung Dissection

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

References

Additional information

Reviews

Our Location

Conduct Science

Resources

Support

Customer service