Contusion Spinal Cord Injury via a Microsurgical Laminectomy in the Regenerative Axolotl

Summary

This manuscript presents protocols for surgically inflicting controlled blunt and sharp spinal cord injuries to a regenerative axolotl (Ambystoma mexicanum).

Abstract

The purpose of this study is to establish a standardized and reproducible regenerative blunt spinal cord injury model in the axolotl (Ambystoma mexicanum). Most clinical spinal cord injuries occur as high energy blunt traumas, inducing contusion injuries. However, most studies in the axolotl spinal cord have been conducted with sharp traumas. Hence, this study aims to produce a more clinically relevant regenerative model. Due to their impressive ability to regenerate almost any tissue, axolotls are widely used as models in regenerative studies and have been used extensively in spinal cord injury (SCI) studies. In this protocol, the axolotls are anesthetized by submersion in a benzocaine solution. Under the microscope, an angular incision is made bilaterally at a level just caudal to the hind limbs. From this incision, it is possible to dissect and expose the spinous processes. Using forceps and scissors, a two-level laminectomy is performed, exposing the spinal cord. A custom trauma device consisting of a falling rod in a cylinder is constructed, and this device is used to induce a contusion injury to the spinal cord. The incisions are then sutured, and the animal recovers from anesthesia. The surgical approach is successful in exposing the spinal cord. The trauma mechanism can produce contusion injuries to the spinal cord, as confirmed by histology, MRI, and neurological examination. Finally, the spinal cord regenerates from the injury. The critical step of the protocol is removing the spinous processes without inflicting damage to the spinal cord. This step requires training to ensure a safe procedure. Furthermore, wound closure is highly dependent on not inflicting unnecessary damage to the skin during incision. The protocol was performed in a randomized study of 12 animals.

Introduction

The overall goal of this study was to establish a controlled and reproducible microsurgical method for inflicting blunt and sharp SCI to the axolotl (Ambystoma mexicanum), producing a regenerative spinal cord injury model.

SCI is a severe condition that, depending on the level and extent, inflicts neurological disability to the extremities along with impaired bladder and bowel control^1,2,3. Most SCI are the result of high energy blunt trauma such as traffic accidents and falls^4,5. Sharp injuries are very rare. Therefore, the most common macroscopic injury type is contusions.

The mammalian central nervous system (CNS) is a non-regenerative tissue, hence no restoration of neurological tissue following SCI is seen^6,7,8. On the other hand, some animals have an intriguing ability to regenerate tissues, including CNS tissue. One of these animals is the axolotl. It is widely used in studies of regenerative biology and is of interest in spinal cord regeneration, because it is a vertebrate^9,10,11,12.

Most SCI studies in the axolotl are performed as either amputation of the entire tail or ablation of a larger part of the spinal cord^9,10,11,12. Recently, a new study was published on blunt injuries¹³ that mimics clinical situations better. Whereas complete appendage amputation in the axolotl results in full regeneration, some non-amputation-based regenerative phenomena are dependent on the critical size defect (CSD)^14,15. This means that injuries exceeding a critical threshold are not regenerated. To develop a regenerative model with a higher clinical translational value, this study investigated whether a 2 mm blunt trauma would exceed the CSD limit.

This method is relevant for researchers working on spinal cord regeneration in small animal models, especially in the axolotl. Furthermore, it may be of more general interest, because it exhibits a way of using standard laboratory equipment to develop a blunt trauma mechanism that is suitable for use in small animals in general.

Protocol

All applicable institutional and governmental regulations concerning the ethical use of animals were followed during this study. The study was conducted under the approval id: 2015-15-0201-0061 by the Danish Animal Experiment Inspectorate. Animals were Mexican axolotls (Ambystoma mexicanum, mean body mass ± STD: 12.12 g ± 1.25 g).

1. Preparation

1. Prepare axolotl for anesthesia.
   1. Use high quality non-chemically treated tap water. If unavailable, use 40% Holtfreter’s solution.
   2. Dissolve 200 mg of ethyl 4-aminobenzoate (benzocaine) in 3 mL of acetone. Dissolve this solution in 1 L of tap water or 40% Holtfreter’s solution.
2. Use a standard Petri dish (100 mm in diameter) placed under a stereo microscope as a surgical table. Place a surgical textile cloth on the Petri dish.
   NOTE: Using a Petri dish as a surgical area enables moving and rotation of the animal without touching it, ensuring spinal stability during surgery.
3. Prepare all sterile microsurgical instruments (i.e., scissors and anatomical forceps).

2. Anesthesia

1. Place the axolotl in a container with benzocaine solution for approximately 45 min to ensure deep and stable anesthesia.
   NOTE: The given concentration of benzocaine will cause anesthesia in all sizes of axolotls.
2. Check for signs of general anesthesia within 30-45 min. These include a complete lack of gill movements, righting reflex, or response to either tactile or painful stimuli (gentle pinching of toe web).
3. To maintain anesthesia, wrap the animals in paper towels wetted in the anesthetic solution. Wet these regularly with this solution during the surgical procedure to ensure that the skin and gills are kept moist.
4. Recover the animal after the surgery by placing it in a container containing fresh tap water. Observe signs of recovery, such as gill movement and regained righting reflex, within 1 h¹⁶.

3. Microsurgical Laminectomy

NOTE: The laminectomy is performed under a stereomicroscope.

1. Place the animal in the prone position on the Petri dish. Wrap it in paper towels so that the tail is exposed.
   NOTE: The paper towels are excellent for ensuring stability throughout the procedure.
2. Identify the hind limbs. Make the first incision just caudal to them.
   1. With a pair of microscissors, perform a vertical incision from the keel until the bony prominence of the spinous processes are felt.
      NOTE: Be very careful when grasping the keel and skin with forceps, because these easily inflict damage to the delicate skin.
   2. Extend the cut laterally, so the incision traverses the entire width of the tail.
   3. Grasp the spinous process with forceps to ensure the right depth.
   4. Extend the vertical incisions 1 mm below the spinous process on both sides.
3. Place the animal on one side to perform ventral and horizontal incisions as stated below.
   1. With a pair of microscissors, starting from the ventral point of the vertical incision, make a horizontal incision of approximately 15 mm for animals 10-20 g in weight. Make the incision longer for larger animals, and shorter for smaller animals.
   2. Using the scissors, dissect medially through the horizontal incision until the vertebral column is felt in the midline.
   3. Repeat steps 3.3, 3.3.1, and 3.3.2 on the other side of the animal.
4. Having dissected in the deep medial plane from both sides, dissect through the midline, thereby connecting the two horizontal incisions.
   1. Move the free piece of tail and keel to one side, exposing the spinous processes (Figure 1).
   2. Fixate the tail piece using wet paper towels.
5. Place the animal in the prone position again with the head facing the surgeon’s non-dominant side.
   1. With a pair of forceps, grasp the spinous processes just caudal to the hind limbs. Apply a gentle lift both up and towards the head of the animal.
   2. Place the blades of a pair of microscissors horizontal around the process and gently cut it. The lift on the process ensures that it is now removed, exposing the spinal cord.
   3. Grasp the spinous process just caudal to the one that was just removed and repeat steps 3.5.1 and 3.5.2.
      NOTE: This should leave an exposed spinal cord corresponding to two vertebral levels. When performing the laminectomy, a white foamy secretion often appears. The spinal cord is easily identified by its distinctive shine, along with a vessel running along the midline.
   4. Depending on the size of the animal, the exposed area may not be wide enough. Using two pairs of forceps, grasp the laminae on both sides of the spinal cord and twist these laterally with a gentle movement.

4. Introducing a Contusion Type Injury (Figure 2)

1. Keep the animal in the prone position.
2. Use the Petri dish to transfer the animal to the trauma unit.
3. Have an assistant shine a flashlight on the spinal cord.
4. Place the contusion trauma unit cylinder above the exposed spinal cord using the microadjusters on the unit. Aim through the cylinder.
5. Lower the cylinder until it is level with the laminae.
6. Attach the falling rod to the electromagnet. Place the desired falling height adjustment cylinder on the trauma unit.
7. Place the falling rod in the cylinder.
   NOTE: For a blinded study, the surgeon should now leave the room without knowing if the animal will be assigned to an injury or a sham surgery group.
8. Turn off the electromagnet. The rod falls to the exposed spinal cord.
9. Use the height adjustment screw to lift the rod from the spinal cord.
10. Confirm the injury by looking at the spinal cord through the microscope. The injured site will appear darker, and bleeding from the midline vessel will be apparent.

5. Introducing a Sharp Injury

NOTE: Perform these steps after 3.5.4.

1. With a pair of microscissors cut the spinal cord in a perfect vertical cut.
2. Repeat the cut 2 mm to the caudal side of the body.
   NOTE: The length of the removed piece of spinal cord can be adjusted as per the study requirement. However, a 2 mm cut will be regenerable¹⁰.
3. Ensure that the cuts are complete. Upon completion, feel the blades of the scissors scraping along the ventral part of the spinal canal.
4. Lift the 2 mm piece of spinal cord from the spinal canal.

6. Closing the Surgical Wound

1. Return the animal to the surgical table. In a blinded study, reposition the keel so the spinal cord is not visible to the surgeon.
2. Keep the animal in the prone position.
   1. Begin placing 10.0 nylon sutures from the most caudal part of the horizontal incision. Close the wounds in one layer.
      NOTE: Do not grasp the skin too tight, because it will inflict necrosis.
   2. Work towards the vertical part of the incision.
   3. When reaching the angle, turn the Petri dish and suture the other horizontal incision.
   4. Set sutures on the vertical incisions.
   5. Do not place sutures in the uppermost part of the keel, because the skin here will not be able to hold.

7. Returning the Animal to the Anesthetic-free Solution

1. Lift the Petri dish with the animal and submerge both very gently into fresh water only 5 cm deep and let the animal slide off.
   NOTE: The shallow water depth ensures that the animal will not attempt to swim to the surface to breathe.
2. Do not change the water during the first week.
3. When feeding the animals, ensure that the food is placed near the animal’s head.
   NOTE: The purpose of these measures is to avoid as much movement as possible during the first week.

8. Postoperative Ultrasound

1. Prior to the termination of anesthesia, use a high frequency ultrasound system to acquire images of the injury that can be used for the construction of three-dimensional images of the SCI site.
2. Attach the transducer to a micromanipulator preferably governed by a remote joystick.
3. Submerge the anesthetized animal in the prone position into a small container filled with anesthetic solution.
   NOTE: Fix the animal with miniature sandbags or other equipment to avoid movement during the scanning sequence.
4. Align the tip of the transducer with the animal’s length axis and submerge it into the benzocaine solution until it is only a few millimeters above the keel behind the hind limbs of the animal.
5. Identify the SCI site.
   NOTE: The injury site is easily recognizable due to the missing spinous processes directly above the SCI.
6. Optimize the image by adjusting the ultrasound settings. Ensure that the SCI site is in the center of the image. Adjust the field of view (i.e., image depth, depth offset, and image width) to cover the SCI site and adjacent healthy tissue. Adjust the two-dimensional gain to optimize the image contrast.
7. By sweeping the ultrasound transducer across the SCI site with an electronically operated micromanipulator, acquire B-mode images covering the SCI site at multiple sagittal cross-sectional slice locations, with consecutive slices with an interslice interval of 50 µm. Acquire cine-images containing 500 frames with a frame rate of ~50 frames/s and a transducer frequency of 40 MHz.
   NOTE: This setup requires an electronic micromanipulator governed by a remote joystick (step 8.2).
8. After finishing the scanning sequence return to step 7.

Results

The purpose of the protocol is to produce an SCI that will paralyze the motor and sensory functions caudal to the injury. Because the axolotl is regeneration-competent it restores function within weeks, allowing researchers to study CNS regeneration during a short time span.

Anesthesia was provided for 45 min to all animals, and no episodes of preterm recovery were experienced. All animals recovered within an hour and showed no signs of damage from anesthesia in the following weeks

Discussion

Because risk of injury to the spinal cord is significant, the critical steps of the protocol are removing the spinous processes and widening of the bony access to the spinal canal if needed. As mentioned in the protocol, removing the most cranial process first is highly recommended. This will mean that the more caudal processes protect the spinal cord from being hit by the scissors. It is recommended to ensure enough surgical access, meaning to not make too small a primary incision. Also, when grasping anything with forc...

Materials

Name	Company	Catalog Number	Comments
25 g custom falling rod	custom home made
30 mm PVC pipe	custom home made
Acetone	Sigma-Aldrich	67-64-1	Propanone
Axolotl (Ambystoma mexicanum)	Exoterra GmbH	N/A	12-22 cm and 10 g - 80 g, All strains (wildtype, melanoid, white, albino, transgenic white with GFP)
Benzocain	Sigma-Aldrich	94-09-7	ethyl 4-aminobenzoate
Electromaget	custom home made
Excel 2010	Microsoft	N/A	Excel 2010 or newer
ImageJ	National Institutes of Health		ImageJ 1.5e or newer. Rasband, W.S., ImageJ, U. S. National Institutes of Health, Bethesda, Maryland, USA, https://imagej.nih.gov/ij/, 1997-2016.
Kimwipes
Microsurgical instruments	N/A	N/A	Forceps and scissors
MS550s	Fujifilm, Visualsonics	MS550s	40 MHz center frequency, transducer
MS700	Fujifilm, Visualsonics	MS700	50 MHz center frequency, transducer
Petri dish	any maker
Soft cloth	N/A	N/A	Any piece of soft cloth measuring approximately 70 x 55 cm2 e.g. a dish towel
Stereo microscope
Vevo 2100	Fujifilm, Visualsonics	Vevo 2100	High frequency ultrasound system