Triggering Reactive Gliosis In Vivo by a Forebrain Stab Injury

Summary

This article describes a detailed protocol to produce a forebrain stab injury in adult mice. The stab injury induces severe reactive gliosis and glial scar formation which can be subsequently examined by standard immunohistochemistry methods.

Abstract

Following injury to the CNS, astrocytes undergo a broad range of biochemical, morphological, and molecular changes collectively referred to as reactive astrogliosis. Reactive astrocytes exert both inflammatory and protective effects that inhibit and promote, respectively, neural repair. The mechanisms underlying the diverse functional properties of reactive astrogliosis are not well understood. Achieving a greater understanding of these mechanisms is critical to developing therapeutic strategies to treat the injured CNS. Here we demonstrate a method to trigger reactive astrogliosis in the adult mouse forebrain using a forebrain stab lesion. This lesion model is simple, reliable, and requires only a stereotaxic device and a scalpel blade to produce the injury. The use of stab lesions as an injury model in the forebrain is well established and amenable to studies addressing a broad range of neuropathological outcomes, such as neuronal degeneration, neuroinflammation, and disruptions in the blood brain barrier (BBB). Thus, the forebrain stab injury model serves as a powerful tool that can be applied for a broad range of studies on the CNS response to trauma.

Introduction

A major challenge for developing successful therapies to treat the injured CNS is an incomplete understanding of the complex multicellular events that are triggered by the trauma. Reactive astrocytes are gaining increasing recognition as a promising target for novel therapies¹. Though historically regarded as hostile to neural repair, reactive astrocytes are now recognized as critical components of a complex, multicellular neuroprotective response that includes attenuation of inflammatory processes and limiting secondary damage and neurodegeneration^2-6. Although the neuropathological characteristics of reactive gliosis have long been well defined, the cellular and molecular mechanisms regulating reactive gliosis, and the diverse array of downstream consequences remain poorly understood. Understanding the mechanisms that drive reactive gliosis, as well as the subsequent cellular and molecular events, is an important step towards developing strategies aimed at promoting the neuroprotective properties of reactive gliosis, while attenuating the detrimental effects.

Here we demonstrate a method to induce severe reactive astrogliosis in the forebrain of adult mice using a stab injury. In contrast to other traumatic brain injury (TBI) models, such as controlled cortical impact (CCI) or fluid percussion injury (FPI), which require specialized equipment to produce an injury, the forebrain stab requires only a stereotaxic device to stabilize the head and a No. 11 scalpel blade. Thus the forebrain stab lesion model is more broadly accessible to a wide range of laboratories that do not have access to the specialized devices necessary for creating an FPI or CCI injury. The method described here enables investigators to reliably and reproducibly trigger a robust gliosis response to investigate subsequent cellular and molecular events. Once recovered from surgery, animals that have received a forebrain stab injury can survive for prolonged periods without the need for specialized care and can be returned to the colony for acute, intermediate, or chronic studies. Though less clinically translatable than FPI or CCI models of TBI, a forebrain lesion produced by a stab injury serves as a simple yet useful experimental model to investigate basic biological mechanisms underlying reactive gliosis and other neuropathological events following trauma to the CNS.

Protocol

Adult (3-4 months old) male mice on a mixed C57BL/6 background were used in this protocol. Animals were kept on a 12 hr light/dark cycle, and allowed free access to food and water. All procedures performed in this protocol were conducted according to protocols approved by the Drexel University Institutional Animal Care and Use Committee.

1. Preparing Surgical Area

1. Disinfect surgical table with 70% ethanol, then cover the entire surgical bench with absorbent pads and arrange surgical instruments adjacent to stereotaxic.
2. Set up stereotaxic equipment without manipulator arm. Arrange the heating pad on the stereotaxic and set to 37 °C. Avoid overheating the animal by placing a small piece of paper towel or surgical pad between the animal and the heating pad.
3. Using autoclaved scissors, cut small pieces of autoclaved gelfoam into a sterile Petri dish containing 0.9% sterile saline solution until ready for use.
   Note: Maintain sterile working environment by using autoclaved instruments and sterile surgical supplies. Re-sterilize surgical instruments during the procedure or in between animals by dipping into a bead sterilizer for 10-15 sec, as needed. Maintain clean gloves throughout the procedure by rubbing hands with 70% ethanol, as needed, to disinfect.

2. Prepping Mouse for Surgery

1. Remove mouse from home cage and weigh (g).
2. Place mouse into isoflurane induction chamber and set oxygen to 2 L/min and isoflurane vaporizer to 5 to induce a surgical plane of anesthesia, about 3-5 min. Monitor for slowed breathing and immobilization. Check that the mouse is fully sedated using the toe pinch reflex.
3. When mouse is fully sedated, place in stereotaxic frame, secure the nose in the nose cone, which is attached with tubing to the isoflurane. Insert ear bars into ear canal and tighten, ensuring the head is stable.
4. Shave the head from ear to ear, and from between the eyes to behind the ears.
5. Sterilize the skin with alternating wipes of isopropyl alcohol and betadine iodine solution, 3 times each.
6. Apply artificial tears to both eyes to prevent them from drying out during the surgical procedure.

3. Surgical Procedure

1. Monitor the depth of anesthesia by pinching the toe or tail. The mouse is in the appropriate surgical plane when there is no response, and the respiration is slow and even.
2. Make a parasagittal skin incision from just behind the eyes to almost between the ears in one single, firm motion using a No. 11 scalpel blade. Move skin aside and clip right side with hemostat.
3. Clear skull of overlying membrane using the dull side of the No. 11 scalpel and cotton tipped applicators. Optionally, wipe the skull with cotton tipped applicator dipped in 0.9% saline solution. Allow to dry completely.
4. Using a small ruler, mark the anterior border of the craniotomy at 1 mm caudal to the coronal suture, and the left edge of the craniotomy at 1 mm lateral to the sagittal suture (Figure 1), with a permanent marker. Then mark the right and caudal borders of the craniotomy at 4 mm from the sagittal and coronal sutures, respectively (Figure 1).
5. Using a 0.5 mm drill bit, begin to make craniotomy by drilling slowly following the permanent marker outline. Be sure not to break through the skull completely. Press gently on the isolated piece of parietal bone with No. 5 forceps, areas of weakness will give way to the pressure. When the thinned bone is sufficiently weak throughout the perimeter, the bone piece is ready for removal.
   NOTE: If the investigator experiences difficulty removing the bone in one piece, this suggests that bone was not sufficiently thinned during drilling. Consider drilling the skull further in subsequent animals to facilitate easy removal of the bone piece.
6. Using a 10 ml syringe fitted with a 23 G needle, apply a small amount of 0.9% saline to soak the isolated bone and drilled area.
7. Attach the manipulator arm to the stereotaxic equipment. Attach a new No. 11 scalpel blade to the probe holder with the sharp side of the blade facing rostrally.
   NOTE: Although the rostral and caudal tissues experience the sharp and blunt edges of the blade, respectively, the mechanical damage induced by the penetrating injury is comparable throughout the extent of the lesion. We observe no appreciable differences in major features of reactive gliosis including upregulation of GFAP expression or proliferation, between rostral and caudal sections.
8. Keeping the manipulator arm out of the way, carefully lift the isolated bone using 5/45 angled forceps. Insert the tip of the forceps into the side of the isolated bone and lift, using leverage to pull off the piece of bone left behind in one full movement.
   NOTE: Be careful not to stab the brain or disturb the dura underneath the skull.
9. Take a small piece of the soaked absorbable gel foam and place on the uncovered brain to prevent it from drying out and soak up any blood that might be present.
10. Once the gel foam is in place, swing the manipulator arm into place and adjust blade to center of craniotomy over the gel foam. Remove the gel foam and lower the blade until the tip touches the dura without puncturing the dura. Mark dorsal/ventral coordinates using the vernier scale on the vertical arm of the stereotaxic.
11. Using the manipulator arm, slowly lower the blade precisely 3 mm into the brain. This is achieved by using the vernier scale markings on the manipulator arm. Allow blade to stay in place for 5-10 sec. Move the stereotaxic arm with blade attachment rostral to caudal three times allowing the blade to reach the rostral and caudal boundaries of the craniotomy before moving to the opposite end.
    NOTE: The dura is not removed prior to inserting the blade. In contrast to the rat dura, which is ~80 μm in thickness⁷, the mouse dura is considerably thinner (only a few cell layers thick) and does not produce an appreciable resistance to the scalpel blade during insertion. Use a new scalpel blade for each mouse to ensure that each animal receives a consistent injury.
12. Slowly raise the stereotaxic arm, removing the blade from the brain. After removal of the blade, immediately place another piece of gelfoam on the brain surface to soak up any excess blood or fluid.
13. Meanwhile, remove the stereotaxic arm and dispose of the No. 11 scalpel blade. Once bleeding has stopped, remove the gelfoam.
14. Close the wound by suturing the skin with non-absorbable suture, such as ethilon or prolene. Sutures should be removed 9-10 days following surgery.
15. Return the mouse to its home cage, and allow the mouse to recover slowly on a heating pad and monitor for any signs of distress. Recovery from isofluorane-induced anesthesia typically occurs within 2-5 min after removal from the isofluorane. Do not leave the animal unattended until it has regained sternal recumbancy.
16. Administer 0.5-1 ml of lactated Ringer’s solution subcutaneously to ensure hydration.

4. Post-surgical Care

1. Closely monitor animals post-operatively until recovery from anesthesia before returning to the colony.
   1. To reduce pain and discomfort post-operatively, administer 0.05-0.1 mg/kg buprenorphine by i.p. injection immediately following the procedure.
   2. Observe animals for 2-3 days post-operatively for severe signs of distress such as restricted movements, lack of grooming, or weight loss. Euthanize animals demonstrating any of these signs of distress and remove from the study.
2. To examine histopathology of injured tissues, euthanize animals by standard intracardial perfusion.
   1. Briefly, anesthetize animals with an overdose of ketamine/xylazine, then intracardially perfused with 15-20 ml of 0.9% NaCl, or until the liver is cleared of blood, followed by 60 ml of 4% paraformaldehyde at the conclusion of the experiment.
3. Dissect brains and post-fix for 2-4 hr in 4% paraformaldehyde before transferring to 30% sucrose solution. Section brains on a cryostat at 40-60 µm, and process by standard histological or immunohistochemical procedures, or as described in Garcia^8,9.

Results

Because animals undergoing this procedure do not require specialized post-operative care, short or long-term time survival periods are easily incorporated into the study, depending on the need to investigate acute or chronic pathology following injury. Principal features of reactive gliosis, such as upregulation of GFAP and hypertrophy of soma, can be observed as early as 2-3 days following injury. The peak phase of proliferation for reactive astrocytes is during days 3-5 following injury¹⁰. The representative...

Discussion

It is critical that the skull or underlying dura are not damaged during the drilling. Use light pressure while drilling to ensure the skull is not punctured. In addition, care should be taken while lifting the skull piece to ensure the dura is not lifted off with the bone.

The forebrain stab injury described here models a penetrating injury to the CNS. Though less clinically translatable than TBI models such as FPI or CCI, the forebrain stab lesion model serves as a useful tool for a broad ran...

Materials

Name	Company	Catalog Number	Comments
Stereotax	Harvard Apparatus	726049
High speed micro drill	Harvard Apparatus	724950
stainless steel scalpel blade, #11	MedVet	JOR581S
5/45 angled forceps	Fine Science Tools	11251-35
Gelfoam sponge 12 cm x 7 mm	Fisher	NC9841478
Rb anti-GFAP	DAKO	Z033429-2	Dilution - 1:20,000 (bright-field); 1:1,000 (fluorescence)
Shp anti-BrdU	Abcam	ab1893	Dilution - 1:20,000 (bright-field); 1:500 (fluorescence)
Biotinylated goat anti-rabbit	Vector Laboratories	BA-1000	Dilution - 1:400 (bright-field)
Biotinylated rabbit anti-sheep	Vector Laboratories	BA-6000	Dilution - 1:400 (bright-field)
Alexafluor 488 goat anti-rabbit	Life Technologies	A-11008	Dilution - 1:400 (bright-field)
Alexafluor 568 donkey anti-sheep	Life Technologies	A-21099	Dilution - 1:1,000 (fluorescence)
DAPI Nucleic Acid Stain	Life Technologies	D3571	Dilution - 1:1,000 (fluorescence)
Cresyl Violet Acetate	Sigma Aldrich	C5042-10G	Dilution - 1% (bright-field)