Induction of Complete Transection-Type Spinal Cord Injury in Mice

Podsumowanie

This protocol describes how to create a precise laminectomy for induction of stable transection-type spinal cord injury in the mouse model, with minimal collateral damage for spinal cord injury research.

Streszczenie

Spinal cord injury (SCI) largely leads to irreversible and permanent loss of function, most commonly as a result of trauma. Several treatment options, such as cell transplantation methods, are being researched to overcome the debilitating disabilities arising from SCI. Most pre-clinical animal trials are conducted in rodent models of SCI. While rat models of SCI have been widely used, mouse models have received less attention, even though mouse models can have significant advantages over rat models. The small size of mice equates to lower animal maintenance costs than for rats, and the availability of numerous transgenic mouse models is advantageous for many types of studies. Inducing repeatable and precise injury in the animals is the primary challenge for SCI research, which in small rodents requires high-precision surgery. The transection-type injury model has been a commonly used injury model over the last decade for transplantation-based therapeutic research, however a standardized method for inducing a complete transection-type injury in mice does not exist. We have developed a surgical protocol for inducing a complete transection type injury in C57BL/6 mice at thoracic vertebral level 10 (T10). The procedure uses a small tip drill instead of rongeurs to precisely remove the lamina, after which a thin blade with rounded cutting edge is used to induce the spinal cord transection. This method leads to reproducible transection-type injury in small rodents with minimal collateral muscle and bone damage and therefore minimizes confounding factors, specifically where behavioral functional outcomes are analyzed.

Wprowadzenie

Spinal cord injury (SCI) is a complex medical problem that results in drastic changes in health and lifestyle. There is no cure for SCI, and the pathophysiology of SCI is not thoroughly understood. Animal SCI models, in particular rodent models, offer an invaluable tool for trialing new treatments, and have been used to explore SCI for decades. To date, over 72% of pre-clinical SCI studies have employed rat models, as compared to a mere 16% that have used mice¹. Although rats, due to their larger size and tendency to form cavities akin to human SCIs, have traditionally been the preferred model animals for studying novel therapeutic approaches, mice (including many transgenic mouse models) are now being used more frequently to study cellular and molecular mechanisms of the SCI². The mouse model offers additional benefits in terms of easier handling, faster reproductive rates and lower costs than rats; mice also exhibit a high degree of genomic similarity with humans^1,2,3. The major disadvantage of the mouse model has been identified as the significantly smaller size that creates challenges for surgical interventions for creating and treating spinal cord injuries^4,5.

There is a gap in the existing literature that highlights the need for a robust and reproducible surgical protocol to induce stable SCI in the mouse model. Therefore, we provide a novel and precise surgical approach in this protocol to overcome these limitations. This protocol provides in-depth guidelines to induce a transection-type injury in mice, as this injury type has been recognized to be the most appropriate to study regenerative and degenerative changes following an injury⁶, as well as neuroplasticity, neural circuitry and tissue engineering approaches⁷. We have chosen to induce the injury in the lower thoracic region, since thoracic level SCI is used most commonly in the literature¹.

Protokół

All procedures were carried out with the approval of the Griffith University Animal Ethics Committee (ESK/04/16 AEC and MSC/04/18 AEC) under the guidelines of the National Health and Medical Research Council of Australia.

1. Animal set-up procedure for the surgery

1. Anesthetize and stabilize the animal.
   1. Use 8–10-week old female C57BL/6 mice. Use 5% isoflurane in 1 L/min oxygen for induction of anesthesia. For maintenance of anesthesia, use 1.5–2% isoflurane in 1 L/min oxygen. Confirm appropriate anaesthetization by establishing a lack of pain reflex in tail and hind paws.
2.  Administer buprenorphine (0.03 mg/kg body weight) for analgesia and enrofloxacin (10 mg/kg body weight) for antibiotic cover, subcutaneously according to body weight. Meloxicam (2 mg/kg body weight) can be given for long term analgesia if required. 
3. Keep the animal’s body temperature steady at 37 °C with a heating pad
   1. Shave the back fur to expose the surgical area over the dorsal spine. Remove the shaved fur from surgical area before sterilizing the incision site. Sterilize the shaved area with sterile cotton swabs soaked in povidone iodine antiseptic liquid and surgical spirit.
4. Tape the paws of the mouse to the surgical area to stabilize the animal (Figure 1A). Place an oval window drape over mouse (Figure 1B).

2. Laminectomy

1. Make a vertical midline incision at the T10 vertebral level using a surgical scalpel.
   1. Locate the spinous process of T10 vertebra to determine the site of laminectomy. The body of the vertebra lies slightly cranial to the tip of the spinous process⁸ (see Figure 2). The tip rests approximately at the midpoint of the T11 vertebra⁸.
   2. Make the incision ~2.5 cm long, so that the spine of T10 vertebra is approximately at the middle of the length of the incision.
2. Reflect the skin and retract with retractors.
   1. Use the straight forceps to lift the skin from the underlying fascia. This will create space for the retractors to be placed; these will keep the surgical field open.
3. Perform blunt dissection of subcutaneous tissue and fascia to expose the spinous processes.
   1. Use the blunt edge of the scalpel to make a small midline incision in the subcutaneous tissue and underlying fascia to expose the spines of T9–T11 vertebrae. Use the fine tip forceps (non-sharp) to perform blunt dissection and reflect the fascia.
4. Split and separate the para-spinous muscles to expose the laminae.
   1. Use the blunt tip of the scalpel to split the dorsal trunk muscles and para-spinous muscles along the spines of the T9–T11 vertebrae. Use the blunt fine tip forceps to perform blunt dissection of the muscles in layers and expose the laminae of the vertebrae. This should minimize any bleeding.
      NOTE: If there is any bleeding, use warmed saline (37 °C) irrigation and cotton swabs to control it and clear blood from the surgical field.
   2. Use the same forceps to make small pockets around the transverse processes of T10 vertebra. Use the curved forceps to stabilize the T10 vertebral body by hooking its prongs under the transverse processes, in the pockets created (Figure 1C).
   3. Thoroughly rinse the T10 laminae with warm saline and gently wipe clean with cotton swabs to clearly visualize the bony surface. Make sure that no muscle/ligament attachments remain along the surface bilaterally.
5. Use a drill with a fine tip (0.55 mm diameter, 7 mm length) to break the laminae bilaterally.
   1. Use the tip of the drill to trace a vertical path from T9–T10 intervertebral space to T10–T11 intervertebral space along both the T10 laminae, without turning the drill on. This is to ensure that the drill bit does not catch any tissue (Figure 1D).
   2. Now turning the drill on, slowly and carefully dig a vertical trench on the right lamina of the T10 vertebra. This part of the laminectomy should create a precise surgical defect throughout the thickness of the bone in a straight vertical line. Maintain the grip with the curved forceps to keep the vertebra stable.
6. Make sure that the tip of the drill does not penetrate through the bone and injure the spinal cord. Repeat the same process on the left side of the lamina, keeping the vertebra stable with the curved forceps. Irrigate with warm saline to wash away any remaining bone fragments.
7. Lift and remove the posterior part of the neural arch (Figure 1F).
   1. Use the angled fine tip forceps to grip the spinous process and remove the whole dorsal segment of the laminae separated by the bilateral drilling. Irrigate and swab again if there is any bleeding, to clearly visualize the spinal cord exposed under the laminectomy window (Figure 1E).

3. Transection

1. Induce the spinal cord injury by transection of the exposed cord with a single slice of the blade.
   1. Use the narrow, round cutting edged blade to slice the cord at the center of the laminectomy window. Ensure to sweep the lateral recesses of the spinal column to induce a complete transection injury (Figure 1G).
2. Confirm completeness of the transection injury using the blunt fine tip forceps and remove any remaining connections at the transection site.
3. Control the bleeding if any, before closing the surgical layers.
4. Use the warm saline to irrigate and clear any bleeding that occurs from the transected cord stumps. Use a cotton swab to apply gentle pressure to achieve hemostasis if needed. Take care not to compress the spinal cord.

4. Closure and immediate post-operative care

1. Bring the muscles together and suture in layer.
   1. Once hemostasis is achieved at the transection site, release the curved forceps grip on the T10 vertebrae. Bring the edges of dissected muscles together along the midline to achieve good apposition.
   2. Suture the muscles in a layer using 5-0 polyglactin 910 absorbable sutures. Make sure that the natural curvature of the spine does not cause any tension at the suture line or open up the sutures, exposing laminectomy site.
2. Close subcutaneous tissue and skin.
   1. Use 5-0 non-absorbable silk sutures to close the skin incision. Make sure that there is no bleeding, clots or debris remaining under the skin before closure. A final irrigation with warm saline may be necessary at this step.
3. Stop the anesthesia. Observe the animal for 10–30 min until recovery. The animal should remain on the heating pad for this duration. Provide water gel and hydrated food in the cage.
4. For post-operative care, include buprenorphine (twice daily), enrofloxacin (once daily) for the first two days prophylactically. Additionally, manually empty the bladder at least twice daily, adhering to the animal ethics committee’s guidelines.
   NOTE: The animals in this experiment were assessed twice daily for their general health and wellbeing, which included checking for persisting pain (to give additional doses of buprenorphine) or infection (additional enrofloxacin), their nutrition and hydration status (give injectable fluids if dehydrated) and any signs of autotomy (provide wound care if minor autotomy). It is strongly recommended that these aspects of post-operative care, including the euthanasia decisions must be determined with the guidance of the institutional animal ethics committee.

5. Evaluation of the injury model

1. Establishing the loss of motor function.
   1. Assess motor behavior on the injured mice 2, 7, 14, 21 and 28 days after the injury in an open field using the Basso Mouse Scale (BMS) to determine the loss of function⁹ (Figure 3C–E).
2. Histological confirmation of injury
   1. Harvest the injured spinal cords with the vertebral columns after euthanasia (done with carbon dioxide in this experiment, as approved by the animal ethics committee).
   2. Fix the tissue by overnight submersion in 4% paraformaldehyde and decalcify the bones by treatment with 20% ethylene diamine tetraacetic acid (EDTA) over 3 weeks, replacing fresh EDTA every 48 h.
   3. Prepare the decalcified spines for cryo-sectioning and cut them into 30 µm thick sections.
   4. Mount the sections on gelatin-coated glass slides for immuno-staining with anti-GFAP antibodies and Hoechst 33342.
   5. Image the slides on a fluorescent microscope (Figure 3A) and perform measurements of the injury size using image analysis software (e.g., Nikon analysis software - NIS Elements [Figure 3B]).

Wyniki

The resulting method as depicted in Figure 1, involves adequate stabilization of the mouse (Figure 1A) and good visualization of the spine and paraspinous tissue (Figure 1B). Spinous process and laminae can be clearly visualized with minimal muscle dissection and blood loss (Figure 1C, highlighted zone). The fine tip drilling is performed as shown in Figure 1D, to create a ...

Dyskusje

This method induces a complete transection type injury at the T10 vertebral level in mice, which results in complete paraplegia of the animal, below the level of injury. Overall, this method results in minimal bleeding, negligible collateral damage and a stable, reproducible injury. As compared to previously published methods of transection without laminectomy¹⁰, this method offers the benefits in terms of direct visualization without manipulating the curvature of the spine, better control over co...

Materiały

Name	Company	Catalog Number	Comments
Baytril injectable 50 mg/mL, 50 mL	Provet	BAYT I	Post-operative care drug
Betadine 500 mL	Provet	BETA AS	Consumable
Castroviejo needle holder, locking	ProSciTech	T149C	Reusable
Ceramic zirconia blade, round with sharp sides, single edge, angled	ProSciTech	TXD101A-X	Reusable
Cotton swabs (5pcs)	Multigate	21-893	Consumable
Dremel Micro	DREMEL	8050-N/18	Cordless rotary tool
Dressing forceps fine	Multigate	06-306	Single use disposable
Drill bits	Kemmer Präzision	SM 32 M 0550 070	Reusable
Dumont #7b forceps	Fine Science Tools	11270-20	Reusable
Dumont tweezers, style 5	ProSciTech	T05-822	Reusable
Fur trimmer	WAHL	WA9884-312	Zero Overlap Hair Trimmer
Iris scissors, Ti, sharp tips, straight, 90mm	ProSciTech	TY-3032	Reusable
Isoflurane isothesia NXT 250	Provet	ISOF 00 HS	Anaesthetic agent
Colibri Retractor - 4cm	Fine Science Tools	17000-04	Reusable
Scalpel handle	ProSciTech	T133	Reusable
Signature latex surgical gloves size 7.5	Medline	MSG5475	Consumable
Sodium Chloride 0.9%	STS	PHA19042005	Consumable
Sterile Dressing Pack	Multigate	08-709	Single use disposable
Sterile Fluid Impervious Drape 60x60 cm	Multigate	29-220	Single use disposable
Surgical spirit 100 mL	Provet	# SURG SP	Consumable
Suture Material - SILK BLK 45CM 5/0 FS-2	Johnson & Johnson Medical	682G	Silk Suture
Suture Material - Vicryl 70CM 5-0 S/A FS-2	Johnson & Johnson Medical	VCP421H	Vicryl Suture
Temgesic 0.3 mg in 1 mL, x 5 ampoules (class S8 drug)	Provet	TEMG I	Post-operative care drug