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Method Article

AAV Systems and Mouse Models for Investigating Ectopic Expression of Neurod1 in Transduced Cells at Subacute and Chronic Times Post-Ischemic Stroke

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DOI:

10.3791/66965

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November 29th, 2024

 ,  ,  ,  ,  ,  ,  , 
1Department of Surgery, Division of Anatomy, University of Toronto, 2Institute of Medical Science, University of Toronto, 3Institute of Biomedical Engineering, University of Toronto, 4Donnelly Centre for Cellular and Biomedical Research, University of Toronto

* These authors contributed equally

In This Article

Summary

This protocol describes the viral-mediated ectopic expression of Neurod1 following cortical ischemic stroke. Neurod1 is delivered (1) using the Cre-Flex AAV system in wild-type mice during the subacute phase post-stroke (7 days) and (2) using a single AAV vector in conditional reporter mice during the chronic phase post-stroke (21 days).

Abstract

Ectopic expression of neurogenic factors in vivo has emerged as a promising approach for replacing lost neurons in disease models. The use of neural basic helix-loop-helix (bHLH) transcription factors via non-propagating virus-like particle systems, including retrovirus, lentivirus, and adeno-associated virus (AAV), has been extensively reported. For in vivo experiments, AAVs are increasingly used due to their low pathogenicity and potential for translatability. This protocol describes two AAV systems for investigating the ectopic expression of transcription factors in transduced cells post-ischemic stroke. In both systems, Neurod1 expression is controlled by the short GFAP (gfaABC(1)D) promoter, which is upregulated in reactive astrocytes post-stroke as well as in endogenous neurons when combined with neurogenic factor expression. In the ischemic stroke model described, focal ischemia is induced by injecting endothelin-1 (ET-1) into the motor cortex of mice, creating a lesion surrounded by reactive GFAP-expressing astrocytes and surviving neurons. Intracerebral injections of AAV are performed to ectopically induce the expression of Neurod1 in the subacute (7 days) and chronic (21 days) phases post-stroke. Within weeks following AAV injection, a significantly higher number of neurons among transduced cells are identified in mice ectopically expressing Neurod1 compared to mice receiving AAV control viruses. The AAV-based strategies used replicated observed outcomes of increased numbers of neurons expressing the reporter gene in a model of mild-to-moderate cortical stroke. This protocol establishes a standard platform for exploring the effects of ectopic expression of transcription factors delivered with AAV-based systems, contributing to the understanding of neurogenic factor expression in the context of stroke.

Introduction

Stroke is a leading cause of disability worldwide1. A stroke happens when blood flow to the brain is disrupted. This can occur either through a hemorrhagic stroke (~15% of cases), where a blood vessel in the brain bursts, or through an ischemic stroke, where blood flow to the brain is blocked1. Ischemic strokes are most prevalent and account for ~85% of stroke cases1. A stroke reduces the delivery of glucose and oxygen to the brain, leading to swift neuronal cell death and impaired neural function.

Ischemic stroke leads to the loss of cells within minutes in the core of the lesion through mechanisms of excitotoxicity, oxidative and nitrosative stress, calcium dysregulation, cortical spreading depolarization, edema, disruption of the blood-brain barrier (BBB), and inflammation2,3. Cells in the peri-infarct regions undergo apoptotic cell death within hours, and for days, post-stroke4. The peri-infarct environment is exposed to pro- and anti-inflammatory signals that lead to the activation of microglia and astrocytes ("reactive" astrocytes), that play diverse roles after injury5,6,7,8. Reactive astrocytes undergo a number of phenotypic changes that include the upregulation of glial fibrillary acidic protein (GFAP), and the formation of a border around the lesion within days post-stroke that helps to limit the extent of the lesion as well as impacts neuroplasticity within the injured tissue8,9,10.

Direct cellular reprogramming allows for the conversion of one mature cell type into a different mature cell type without going through a pluripotent state11. Turning resident brain cells into new neurons to substitute those lost to injury presents an appealing avenue for brain restoration. Stroke is an ideal target for cell-based treatments, given the necessity for therapeutic methods to replace lost neurons, broaden the therapeutic timeframe, and ultimately facilitate functional recovery. To this end, there have been many papers demonstrating astrocyte to neuron (AtN) reprogramming in vitro and in vivo, using a variety of basic- helix-loop-helix (bHLH) neurogenic transcription factors, including Ascl1, Neurog2, Neurod1 and more recently, mutated versions of Ascl1 (Ascl1-SA6) which demonstrated improved reprogramming efficacy11,12,13,14. Important to consider are the many experimental design factors that can impact outcomes, interpretation, and comparisons between studies. Ghazale et al. demonstrated that AtN reprogramming efficiency varied between different mouse strains, AAV serotypes, and promoters when using the same bHLH transcription factor14. Other considerations, such as the injury model, brain region, viral dosing, timing of gene delivery, the reprogramming factor, and the mode of delivery, will impact the outcomes. Recent reports have demonstrated that pre-existing transduced neurons have been misinterpreted as reprogrammed neurons15,16. While the reports do not necessarily negate that AtN conversion occurs, these studies have highlighted the importance of well-controlled experiments, and several papers have put forth recommendations for the next steps in the field15,17,18,19. What is apparent from the literature is that the ectopic expression of bHLH transcription factors can improve functional outcomes in a variety of animal models of neurodegenerative disease/injury13,20,21,22,23,24.

Herein, this protocol Neurod1, a transcription factor of interest, is ectopically expressed in transduced cells13,22. The well-established endothelin-1 (ET-1) model of ischemic stroke to the sensorimotor cortex in adult mice is used. Injection of ET-1 leads to local vasoconstriction mimicking the pathophysiology of ischemic stroke, including functional impairment22,23,25. The commonly employed short-GFAP promoter (gfaABC(1)D) is used to drive the ectopic expression of Neurod1 at the injury site and perilesional region using two viral strategies: (1) AAV5-CAG-Flex::GFP+AAV5-GFAP-Neurod1-Cre delivery in wild-type mice and (2) AAV5-GFAP-Neurod1-Cre delivery in tdTom-Cre reporter mice. To further understand the potential for ectopic expression of Neurod1 in the stroke-injured brain, AAVs were delivered in the subacute phase (7 days), or in the chronic phase (21 days), post-stroke. In both paradigms, significantly more labeled neurons were found within weeks following ectopic Neurod1 expression.

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Protocol

This protocol was approved by the Animal Care Committee at the University of Toronto and adheres to the Guide to the Care and Use of Experimental Animals (2nd Edition, Canadian Council on Animal Care, 2017). For this study, wild-type (C57BL/6J) and transgenic tdTom-Cre reporter (B6.Cg-Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J)22 mouse strains were used. Mice were 7-9 weeks old and included both males and females. Details of the reagents and equipment used are listed in the Table of Materials.

1. Endothelin-1 stroke surgery

NOTE: All surgical instruments and materials are sterilized by autoclave prior to surgery. The stereotaxic frame is sterilized with PREempt. Syringes are sterilized with PREempt, 70% alcohol, and primed with sterile PBS prior to surgery. Endothelin-1 (ET-1) is stored on ice throughout the surgery.

  1. Before starting the surgery, set up the recovery area by placing a clean cage on a heating pad set to 37 °C. Alternatively, if a heating lamp is being used, place only half of the cage under the heat.
    1. To anesthetize mice using isoflurane, follow the protocol approved by the Animal Care Committee. Place the mouse in the anesthetic induction chamber and turn on the oxygen to 1-1.5 L/min, then adjust the isoflurane vaporizer to 3%-5% to initiate anesthesia.
  2. Remove the animal from the chamber once it lies on its side and can no longer stand, and place it onto a clean surface. Ensure the animal is adequately anesthetized by confirming the absence of a toe pinch reflex before and throughout the surgery.
    1. Secure the nose cone extended from the anesthetic machine on the animal and adjust isoflurane to 1%-2% for maintenance. Minimize the exposure to isoflurane (5 min at 3%-5% for induction and the remaining time at 1%-2%), as the procedure can take up to 40 min to complete.
  3. Use animal hair trimmers to remove fur from the neck/behind ears to the nose of the animal to expose the skin on the skull for surgery.
  4. Weigh the animal and administer the appropriate dose of pre-operative analgesic subcutaneously (Meloxicam (2.0 mg/kg)) based on the weight.
  5. Apply eye lubricant ointment on each eye of the mouse to prevent corneal desiccation during anesthesia.
  6. Sterilize the fur-clipped skin area (step 1.3) with 70% ethanol, followed by Chlorhexidine alcohol twice to form a surgical field.
  7. Transfer the mouse onto a mouse stereotaxic frame by first placing its nose in the stereotaxic nose cone, then stabilize the skull with ear bars. Keep the mouse on a heating pad set to 37 °C for thermal support throughout the surgery procedure. Maintain isoflurane at 1%-2% so that the mouse displays a respiratory rate of 1 breath/s.
  8. Repeat application of eye lubrication to prevent corneal desiccation during the procedure if needed. Apply a sterile surgical drape on the body of the mouse, below the ears, to extend the size of the sterile field.
  9. Make a vertical incision parallel to the sagittal plane of the animal from behind the mid-point of the eyes to the back of the head (parietal bone) with a #10 scalpel blade. Expose the skull by gently retracting the skin towards the sides.
  10. Use a Q-tip to disrupt the fascia on the skull and allow the skull to dry to expose bregma.
  11. Set up manipulator arms with drill holder carrying high-speed stereotaxic drill with a .018" drill bit.
  12. Drill 3 burr holes (.018" diameter) with a high-speed stereotaxic drill at the coordinates in the right sensorimotor cortex at 2.25 mm lateral to bregma and +0.0 mm rostral, +1.0 mm rostral and -1.0 mm caudal to bregma.
  13. Replace the drill with a 26 G syringe with a 0.375″ long needle.
  14. Load 1.5 µL of 400 µM ET-1 dissolved in sterile PBS into the syringe. To ensure good flow, release a small volume of ET-1 until a drop can be observed at the tip of the needle. Use a sterile Q-tip to wipe off the excess before performing an injection.
  15. Lower the tip of the needle past the skull in the middle burr hole (in the right sensorimotor cortex at + 0.0 anterior and 2.25 mm lateral to bregma) without puncturing the dura mater. Once aligned, lower the needle 1 mm into the brain.
  16. Inject 1 µL of ET-1 by dispensing 0.1 µL at a time. Push the plunger of the Hamilton syringe forward to dispense 0.1 µL and wait for a minute before injecting the next 0.1 µL, inject until 1 µL has been dispensed.
    1. After the final 0.1 µL has been dispensed (0.1 µL x 10 injections = 1 µL total), keep the needle in place for 10 min to prevent backflow and then slowly withdraw upward to remove the syringe.
  17. To ensure the needle does not become blocked during injection, release a small volume of the ET-1 until a drop can be observed at the tip of the needle. Use a sterile Q-tip to wipe off the excess before performing an injection. Sterilize syringes before injecting the next animal with PREempt, followed by 70% alcohol, and then sterile PBS.
  18. To close the wound, suture the overlying skin on the skull together using a 4-0 polysorb polyester sterile suture.
  19. Administer antibiotic ointment using a cotton swab on the sutured area to prevent post-operative infection.
  20. Remove the mouse from the stereotaxic frame and transfer it to a clean, pre-heated animal housing cage in the recovery area. Turn off the isoflurane vaporizer and oxygen.
  21. Keep the cage on top of a heating pad for the duration of recovery from anesthesia. Alternatively, if using a heating lamp place the mouse on the side of the cage under the lamp so that the mouse can move to the cool side of the cage as it wakes up from anesthesia to prevent overheating. Monitor the mouse as it comes out of anesthesia in the next 5-10 min.
  22. Monitor mouse over the next 3 days (weight monitoring), provide post-operative care (moist mashed chow to ensure hydration), and analgesics (2 mg/kg meloxicam, 2x daily subcutaneously, for 2-3 days) in accordance with regulations set out by local animal care committee.

2. Adeno-Associated Virus (AAV) delivery (7 days - subacute model or 21 days - chronic model) post-stroke

CAUTION: Refer to institutional biosafety guidelines for working with AAVs. As per the guidelines at the University of Toronto, this surgery is performed in a Containment Level 2 Biosafety Cabinet.

NOTE: All surgical instruments and materials are sterilized by autoclave prior to surgery. The stereotaxic frame is sterilized with PREempt. Syringes are sterilized with PREempt, 70% alcohol, and primed with sterile PBS. AAVs are stored on ice throughout the surgery.

  1. Before starting the surgery, set up the recovery area by placing a clean cage on a heating pad set to 37 °C. Alternatively, if a heating lamp is being used, place only half of the cage under the heat.
    1. To anesthetize mice using isoflurane, place the mouse in the anesthetic induction chamber and turn on the oxygen to 1-1.5 L/min, then adjust the isoflurane vaporizer to 3%-5% to initiate anesthesia (following the instutionally approved protocols).
  2. Remove the animal from the chamber once it lies on its side and can no longer stand on a clean surface. Ensure the animal is adequately anesthetized by confirming the absence of a toe pinch reflex before and throughout the surgery.
    1. Secure the nose cone extended from the anesthetic machine on the animal and adjust isoflurane to 1%-2% for maintenance. Minimize the exposure to isoflurane (5 min at 3%-5% for induction and the remaining time at 1%-2%), as the procedure can take up to 40 min to complete.
  3. Weigh the animal and administer the appropriate dose of pre-operative analgesic subcutaneously (Meloxicam (2.0 mg/kg)) based on the weight.
  4. Apply eye lubricant ointment on each eye of the mouse to prevent corneal desiccation from being under anesthesia.
  5. Sterilize the fur-clipped skin area with 70% ethanol, followed by Chlorhexidine alcohol twice to form a surgical field.
  6. Transfer the mouse onto a mouse stereotaxic frame by first placing its nose in the stereotaxic nose cone, then stabilize the skull with ear bars. Keep the mouse on a heating pad set to 37 °C for thermal support throughout the surgery procedure. Maintain isoflurane at 1%-2% so that the mouse displays the respiratory rate of 1 breath/s.
  7. Repeat application of eye lubrication to prevent corneal desiccation during the procedure if needed. Apply a sterile surgical drape on the body of the mouse, below the ears, to extend the size of sterile field.
  8. If still present, cut sutures with a pair of surgical scissors and remove the scab with a pair of forceps. Retract the skin to expose the skull.
    NOTE: For the chronic model it may be necessary to excise the skin with #10 scalpel blade.
  9. Use a sterile Q-tip to dry the skull and locate burr holes.
  10. Set up manipulator arms of the stereotaxic apparatus with a needle holder carrying a 26 G Syringe with a 0.375″ long needle.
  11. Load 3.4 µL of AAV (1 x 109 Genome Copies(GC)/µL for acute study; 1 x 1010 GC/µL for chronic study) into the syringe. To ensure that the flow is good, release a small volume of AAV until a drop can be observed at the tip of the needle. Use a sterile Q-tip to wipe off the excess before performing an injection.
  12. Lower the tip of the needle past the skull in the first burr hole used for ET-1 (in the right sensorimotor cortex at + 0.0 anterior and 2.25 mm lateral to bregma) without puncturing the dura mater. Once aligned, lower the needle 1 mm into the brain. (0 AP, +2.25 ML, −1.0 DV from bregma).
  13. Inject 1 µL of AAV (1 x 109 GC/µL for acute study; 1 x 1010 GC/µL for chronic study) by dispensing 0.1 µL at a time. Push the plunger of the Hamilton syringe forward to dispense 0.1 µL and wait for a minute before injecting the next 0.1 µL; inject until 1 µL has been dispensed. After the final 0.1 µL has been dispensed (0.1 µL x 10 injections = 1 µL total), keep the needle in place for 10 min to prevent backflow and then slowly withdraw upward to remove the syringe.
  14. To ensure the needle does not become blocked during injection, release a small volume of the AAV until a drop can be observed at the tip of the needle. Use a sterile Q-tip to wipe off the excess before performing an injection.
  15. Repeat steps 2.11-2.14 for the AAV injections into the two remaining burr holes (in the right sensorimotor cortex at +1 AP, +2.25 ML, −1.0 DV; and −1 AP, +2.25 ML, −1.0 DV mm from bregma. Sterilize syringes before injecting the next animal with PREempt, followed by 70% alcohol, and finally sterile PBS.
  16. To close the wound, suture the overlying skin on the skull together using a 4-0 polysorb polyester sterile suture.
  17. Administer antibiotic ointment by using a cotton swab on the sutured area to prevent post-operative infection.
  18. Remove the mouse from the stereotaxic frame and transfer it to a clean, pre-heated animal housing cage in the recovery area. Turn off the isoflurane vaporizer and oxygen.
  19. Keep the mouse inside the cage and sterilize the outside of the cage with PREempt and 70% alcohol before placing it on top of a heating pad for recovery from anesthesia.
    1. Alternatively, if using a heating lamp, place the mouse on the side of the cage under the lamp so that the mouse can move to the cool side of the cage as it wakes up from anesthesia to prevent overheating. Monitor the mouse as it comes out of anesthesia in the next 5-10 min.
  20. Monitor the mouse over the next 3 days (weight monitoring), provide post-operative care (moist mashed chow to ensure hydration), and analgesics (2 mg/kg meloxicam, 2x daily subcutaneously, for 2-3 days) in accordance with regulations set out by local animal care committee.

3. Tissue preparation and dissection

  1. To euthanize mouse, intraperitoneally inject the mouse with Tribromoethanol (Avertin) (conc. 12.5 mg/mL, 250 mg/BW(kg)) and allow 2-3 min for complete induction. Ensure the animal is adequately anesthetized by removing the mouse from the cage and confirming the absence of a toe pinch reflex.
  2. In a fume hood, perform transcardial perfusion26 with saline followed by 4% paraformaldehyde. Spray the head of the euthanized animal with 70% ethanol until soaked.
  3. Cut off the head at the base of the skull by using a pair of large scissors. Cut open the skin along the midline of the entire skull using straight 4-1/2" iris scissors. Retract the skin to expose the skull.
  4. From the opening of the spinal cavity at the neck, insert the iris scissors and cut through the skull along the sagittal and inter-frontal suture.
  5. Then insert scissors into the two eye sockets and cut through the frontal-nasal suture.
  6. Take particular care not to damage the brain. Pry open the skull plates to the left and right until the brain is exposed.
  7. Transfer the brain into a scintillation vial with 4% paraformaldehyde. Keep the brain in 4% paraformaldehyde at 4 °C for 4 h.
  8. The brain is then cryoprotected by replacing 4% paraformaldehyde with 30% sucrose. Keep the brain in 30% sucrose at 4 °C until the brain sinks to the bottom of the vial (~12 h).

4. Sectioning frozen brains

  1. Set the temperature of the cryostat chamber between -20 °C to -26 °C.
  2. Make a coronal cut across the cerebellum and attach the flat, caudal part of the brain onto a cryostat chuck with optimal cutting temperature compound (OCT).
  3. Place the chuck with the brain in the disk holder and align the olfactory bulb with the knife blade.
  4. Section the brain into coronal slices with a thickness of 20 µm. To avoid streaks or scratch lines on sectioned brain slices, place a piece of tape on the sectioning platform to create a small gap between the anti-rolling plate and the platform. A paintbrush can be used to adjust the orientation of the section before capturing it on a positively charged glass slide.
  5. Store slides containing brain sections at -20 °C.

5. Immunohistochemistry and quantification

  1. Process the sections for immunohistochemistry using a primary antibody against NeuN, a pan-neuronal marker, according to standard protocols22,26.
  2. Thaw cryopreserved brain sections for 5 min at room temperature. Wash the sections three times with 1x phosphate-buffered saline (PBS) for 5 min each.
  3. Incubate the sections in 0.3% Triton X-100 in PBS for 20 min at room temperature to permeabilize.
  4. Prepare the blocking solution (5% normal goat serum and 0.3% Triton X-100 in PBS) and incubate the sections in the blocking solution for 1 h at room temperature.
  5. Dilute the primary antibody, rabbit anti-NeuN, at a 1:500 ratio in the blocking solution. Incubate the sections with the primary antibody solution overnight at 4 °C.
  6. Wash the sections three times with 0.3% Triton X-100 in PBS, for 5 min each wash.
  7. Dilute the secondary antibody, goat anti-rabbit 568 or 488, at a 1:1000 ratio in PBS. Incubate the sections with the secondary antibody solution for 1 h at room temperature, protected from light.
  8. Perform three washes with 0.3% Triton X-100 in PBS, for 5 min each. Incubate the sections with DAPI (1:10,000 in PBS) for 5 min at room temperature.
  9. Wash the sections three more times with PBS, for 5 min each. Mount the stained sections on a glass slide with a fluorescence mounting medium and a glass coverslip. Allow mounted slides to dry overnight.
  10. Image stained slides using a 20x objective with a confocal microscope to obtain views of Reporter+ cells around the stroke lesion in 3 different coronal sections (between the anatomical regions of the crossing of the corpus callosum anteriorly and the crossing of the anterior commissure posteriorly) for each animal.
    NOTE: The distance of the AAV spread will vary within the hemisphere. However, based on our experience and post-tissue processing analysis of Reporter+ cells, the AAV spread generally spans approximately +3 mm to -3 mm from bregma. Reporter+ cells around the stroke lesion (as identified by the absence/sparse staining of NeuN) in the area between where the corpus callosum crosses the midline and the anterior commissure connects should be quantified.
  11. For quantification of transduced neurons, perform colocalization analysis. Manually count the number of double-positive cells (Reporter+NeuN+) and the total number of transduced (Reporter+) cells in brain slices. For results herein, >3 fields of view were counted surrounding the lesion (medial, lateral, and inferior) from 3 coronal sections per mouse.

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Results

To examine the cellular outcomes of short GFAP (gfaABC(1)D) promoter driven, ectopic Neurod1 expression in wild-type (C57BL/6J) and in transgenic tdTom-Cre reporter strains (specifically, B6.Cg-Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J)22 two AAV5-based systems were utilized in ET-1 stroke injured mice (Figure 1A-C). In a subacute model of stroke, Neurod1 was packaged in the Cre-Flex AAV system and delivered to...

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Discussion

This protocol details two AAV systems and mouse models for investigating the ectopic expression of Neurod1 in the context of a mild-moderate ET-1 cortical stroke model. A number of critical steps relating to the ET-1 stroke are important to consider for the reproducibility and consistency of the injury. Burr holes must be carefully drilled without puncturing the dura mater to prevent unintentional surgical injury. It is important to use consistent landmarks, in this case bregma, to ensure a similar region of int...

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Disclosures

The authors declare no competing interests.

Acknowledgements

This work was supported by the Heart and Stroke Foundation, Ontario Institute of Regenerative Medicine, Canada First Research Excellence Fund (Medicine by Design, MbD), and Connaught Innovation Award (University of Toronto).

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
#77 Drill Bit (.018”)David Kopf Instruments8177
AAV5-GFAP(0.7)-mNeurod1-2A-iCreVector Biolabs
Absorbable Suture with Needle Polysorb™ Polyester CV-15 3/8 Circle Taper Point Needle Size 4 - 0 BraidedCovidienGL-881
Anti-NeuN Antibody (rabbit) Millipore SigmaABN78 
C57BL/6 MiceCharles River027
Chlorhexidine SolutionPartnarPCH-020
Contec™ PREempt™ RTU Disinfectant SolutionFisher Scientific29-636-6212
CryostatThermo Scientific HM525 NX 
Endothelin 1Millipore Sigma05-23-3800-0.5MG
Feather Safety Razor Microtome BladesFeather12-631P
Fisherbrand Cover Glasses: RectanglesFisher Scientific12-545M 
Fluorescence Mounting MediumAgilent TechnologiesS3023 
Hamilton SyringeHamilton Company7634-01
High Speed Stereotaxic DrillDavid Kopf Instruments1474
Metacam Solution (Meloxicam)Boehringer Ingelheim
O.C.T CompoundFisher Scientific23-730-571
pAAV2/5-GFAP-iCreVector BuilderP190924-1001suq
Polyderm Ointment USPTARO2181908
SOMNI Scientific™ The Animal Temperature Heating PadFisher Scientific04-777-177
Stereotaxic InstrumentsDavid Kopf InstrumentsModel 902
Superfrost Plus Microscope Slides, whiteFisherbrand12-550-15
tdTomato Reporter Mice The Jackson Laboratory007914
V-1 Tabletop Laboratory Animal Anesthesia SystemVetEquip901806

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