Widespread Transduction of Mouse Neocortical Neurons by Subarachnoid Injection of AAV2

Summary

A new technique for widespread delivery of adeno-associated virus that uses subarachnoid virus infusion is described. This method not only ensures widespread transduction of mouse neocortical neurons in superficial layers but also results in selective expression of the transgene in layer five pyramidal neurons, even when using a non-selective promoter.

Abstract

Recombinant adeno-associated viruses are a flexible and powerful tool for the delivery and expression of various genes of interest in many areas of experimental biology, particularly in neuroscience. The most popular method to drive the expression of a desired transgene in a particular brain area is to inject an AAV vector directly into the brain parenchyma. However, this method does not allow widespread neuronal transduction that is required for some in vivo experiments. In this article, we present a new technique for widespread gene expression in the mouse neocortex based on viral infusion into the subarachnoid space of the brain. This neuronal labeling method not only ensures widespread transduction of neurons in adult mouse superficial neocortical layers but also results in expression of the transgene in a large population of layer five pyramidal neurons with high specificity even when using a strong non-selective promoter such as CAG. Moreover, because cell transduction takes place at a significant distance from the injection site, this method can help preserve brain tissue for subsequent optical or electrophysiological recordings of neuronal activity.

Introduction

The mammalian brain consists of many inhibitory, excitatory, and modulatory cells interconnected into circuits by trillions of synapses¹. One of the central challenges of neuroscience is to decode the role of distinct cell types in the organization and function of brain circuits and behavior. Manipulating genetically defined cells within the brain requires methods to introduce and express transgenes. Viral-based gene delivery systems are by far the most effective and simple method for gene delivery into the central nervous system². Viral delivery systems are based on replicating viruses (adenoviruses, adeno-associated viruses (AAVs), lentiviruses, and retroviruses) that have the ability to deliver genetic information into a host cell^2,3.

AAV-based vectors have now become one of the most widely used tools for the delivery of desired transgenes to cells within the brain, both for purposes of basic neuroscience research and to develop gene therapy for neurological diseases. When compared against other viruses, replication-defective AAVs possess many features that make them ideal vectors for these purposes. Most notably, AAV vectors efficiently transduce nondividing (terminally differentiated) cells such as neurons and glial cells, resulting in high levels of transgene expression in vivo². The vectors can be easily produced at a high functional titer suitable for in vivo use^3,4,5. Importantly, adeno-associated virus-mediated gene delivery in vivo does not produce histopathological alterations and vector-related toxicity⁶. Unlike adenoviral vectors, in vivo administration of AAV vectors in animal models usually does not elicit host immune responses against transduced cells, enabling stable transgene expression within the brain parenchyma for extended periods of time^2,7,8.

Another reason for the popularity of AAV vectors is the broad array of AAV serotypes with unique tissue and cell-type tropisms^{9,10,11,12,13,14}. Distinct capsid proteins expressed by different AAV serotypes result in the use of different cell surface receptors for cell entry and, thus, specific tropisms^10,14.

AAV tropism is determined not only by capsid proteins but by many other factors¹⁴. It has been shown that AAV serotypes 1, 2, 6, 7, 8, and 9 transduced both neurons and astrocytes in primary culture^15,16, but exhibited strong neuronal tropism following intraparenchymal brain injection^17,18. The method used for AAV vector preparation can also influence nervous cell tropism, even for the same serotype. For example, CsCl-purified AAV8 possessed strong astroglial tropism following intraparenchymal brain injection, while iodixanol-purified AAV8, injected under identical conditions, transduced only neurons¹⁹. AAV tropism may also be affected by the injected dose and volume¹⁴. For example, high titer rAAV2/1 efficiently transduced both cortical excitatory and inhibitory neurons, but the use of lower titers exposed a strong preference for transduction of cortical inhibitory neurons²⁰.

Thus, it is not possible to achieve robust cell-type specificity based solely on the capsid serotype. Cell-type specific promoters can be used to overcome the broad natural tropism of the AAV capsid. For example, human synapsin I is used for targeting neurons²¹, the CaMKII promoter can drive transgene expression in glutamatergic excitatory neurons with high specificity²⁰, the ppHcrt promoter targets hypocretin (HCRT)-expressing neurons in the lateral hypothalamus²², the PRSx8 promoter targets noradrenergic and adrenergic neurons that express dopamine beta-hydroxylase²³, and the GFAP promoter can drive astrocyte-specific expression²⁴. However, some cell-specific promoters have weak transcriptional activity and cannot drive sufficient levels of transgene expression²⁵. Furthermore, the short promoters that fit in AAV viral vectors often do not retain cell-type specificity^1,26. For example, it has been shown that a CaMKII construct also transduced inhibitory neurons¹².

Besides cell-type specificity (tropism), another significant feature of AAVs is transduction efficiency. The various AAV serotypes have different diffusional properties. AAV2 and four viral vectors diffuse less readily through the brain parenchyma and, therefore, mediate transduction over a smaller area^17,27. The most widespread neuronal transduction is observed with AAV serotypes 1, 9, and rh.10^{11,17,18,19,28}.

The most popular method to drive the expression of a desired transgene in a particular brain area is to inject the AAV vector directly into the brain region of interest (parenchyma)³. Following intra-parenchymal injection, even AAV serotypes with more effective diffusion through the brain transduce typically only a local area around the injection site ¹². Moreover, intraparenchymal injection is an invasive procedure and leads to tissue damage adjacent to the region of interest. Thus, this method of virus injection is unsuitable for some experimental tasks. For example, extensive labeling of cells is highly desirable in experiments aimed at studying cortical neuron functions in freely moving animals, including with the use of one- or two-photon microscopy^29,30,31,32.

Here, we describe a new adeno-associated virus injection technique that uses subarachnoid virus infusion to provide widespread transduction of neocortical neurons in adult mice and preserve brain tissue for subsequent optical or electrophysiological recordings of neuronal activity. This method not only ensured widespread transduction of neurons in superficial neocortical layers but resulted in expression of the transgene in a large population of layer five pyramidal neurons with high specificity even when using a strong non-selective promoter such as CAG.

Protocol

Experiments were performed on adult C57Black/6 mice, 2-4 months of age, of both sexes (Pushchino Breeding Center, Branch of the Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry of RAS). Mice were housed in a temperature-controlled vivarium (22 ˚C ± 2 ˚C, 12 h light/dark cycle, lights on at 08.00 h) with food and water ad libitum. All experimental procedures were conducted in accordance with the ARRIVE guidelines and Directive 2010/63/EU for animal experiments. The study protocol was approved by the Ethics Committee of the IHNA RAS (protocol N1 from 01.02.2022). Every effort was made to minimize animal suffering and to ensure the reliability of the results.

1. Preparation for surgery

1. Sterilize all surgical instruments prior to the start of surgery. Clean the surgical area using 70% ethanol.
2. Check the isoflurane level in the anesthesia system and fill it if needed. Place a clean paper towel on the bottom of the induction chamber.
3. Place a heating pad on the stereotaxic frame. Cover the pad with a clean paper towel.
4. Prepare a bottle with 2% bleach to collect micropipette tips, tubes, cotton swabs, and other items that come into contact with the virus.
5. Remove an aliquot of AAV from the -80 °C freezer and place it on ice to thaw.

2. Syringe preparation

1. Clean a 5 µL Hamilton microsyringe with a 33G blunt RN needle. Aspirate and then dispense 70% ethanol. Repeat 3x with fresh ethanol. Rinse the syringe with distilled water to remove the excess ethanol. Repeat 3x.
2. Take the plunger out and fill the barrel with Vaseline oil through the flange using an insulin syringe. Make sure that there are no air bubbles in the microsyringe.
   NOTE: Trapped air is compressible and affects syringe accuracy and precision.
3. Put the plunger back in the microsyringe and dispense a drop of oil. Place the syringe into the stereotaxic injector so that the scale is visible for monitoring the volume of solution dispensed.

3. Preparation of mice for surgery

1. Weigh a mouse. Place the mouse into the anesthesia induction chamber connected to the isoflurane. Turn on the isoflurane vaporizer set to 5% and adjust the flow rate to 250 mL/min. An adequate depth of anesthesia is achieved within 5 to 7 min.
2. To verify the surgical level of anesthesia, check for the absence of whisking and withdrawal reflex during a hind paw painful pinch, and the absence of blinking upon eye contact.
3. Switch the vaporizer outflow valve from the induction chamber to the stereotaxic mask. Reduce the isoflurane to 1.8%-2.0% and set the flow rate according to the mouse weight (70-90 mL/min for mice weighing between 25 to 35 g).
4. Remove the mouse from the induction chamber and place it in the stereotactic apparatus on top of a heating pad (37 °C) to prevent hypothermia during surgical anesthesia. Place the front teeth in the tooth bar and then mount the animal mask.
5. Carefully position the animal on the ear bars. The correct head position in the stereotactic apparatus allows vertical but not lateral movement of the head. Ensure the animal's positioning does not cause distress. Monitor the depth of anesthesia, animal breathing, and body temperature carefully throughout the operation.
6. Shave the head from the eyes to behind the ears. Clean the shaved surface of the head by swabbing with 70% ethanol, followed by swabbing with a 5% alcohol solution of iodine.
7. Apply ophthalmic gel to prevent eye drying. Apply 4% lidocaine solution topically and dexamethasone (0.02 mL at 4 mg/mL) subcutaneously to prevent surgery-related pain and reduce the possible inflammatory response.
8. With a sterile scalpel blade and scissors, make a 4-5 mm long incision along the midline of the head to open the scalp. Start with a small incision between the ears and then expand it using scissors to avoid damage to the skull.
9. Swab the surface of the skull with a small amount of 3% hydrogen peroxide to visualize the stereotaxic landmarks: bregma and lambda. Stop the reaction with 0.9% NaCl saline immediately. Scrape away tissues on top of the skull with a bone scraper.
10. Mount the pre-prepared motorized injector with the Hamilton syringe on the stereotaxic arm. Direct a surgical light source onto the exposed skull and focus the microscope onto the bregma.
11. Looking through the microscope, manipulate the arm of the stereotactic apparatus to center the top of the needle directly over the bregma.
12. Using the tip of the needle, align the bregma and lambda horizontally and then move the arm back to bregma and record the coordinates. Use these bregma coordinates and the atlas coordinates of the region of interest to calculate the relative coordinates of the targeted area.
13. Move the needle to the targeted area. Lower the needle at the new coordinates and mark this position. Select the site for viral microinjection in close proximity to the target area while considering the spread of the virus.
    NOTE: This will avoid damaging the tissue in the region of interest. We used the following coordinates for the region of interest: AP-3.4, ML -2.0, and for viral microinjection: AP-2.0, ML -1.4. The above coordinates are optimal if primary visual cortex neurons are to be infected.
14. If possible, avoid regions with large blood vessels. Under a surgical microscope, bright, cold light illumination, and 0.9% NaCl saline immersion, large blood vessels in the skull and on the brain surface should become apparent.

4. Virus injection

1. Take a sterile dental bur (0.5 - 0.8 mm in diameter). Viewing the surface of the skull through the surgical microscope, drill a small craniotomy manually (by hand) or using a microdrill. Take care not to put excessive pressure on the skull.
2. Move the arm out of the way to prevent damage to the microsyringe needle during the drilling of the craniotomy (hole). Apply 0.9% NaCl saline immersion and pause intermittently to avoid heating the bone and damaging the dura mater. Use pressurized air to blow away bone dust.
   NOTE: The following types of carbide dental burs are suitable: pear-shaped (preferably), rounded cylinder, and round.
   1. When thinning the bone with the bur, ensure the thinning is uniform across the entire circumference. This will make it easier to remove the bone without damaging the dura mater. To do this, use a bur with a round tip initially and then a bur with a flat tip. Keep the bur perpendicular to the bone; otherwise, one side will be thinner than the other.
      NOTE: Additionally, the smaller the size of the bur and the smaller the craniotomy performed, the more complex the manipulation will be. It is recommended that a craniotomy of smaller diameter (0.5-0.6 mm) be performed when further using the animal for in vivo work. If planning to use the animal's brain for ex vivo work, a larger craniotomy diameter is acceptable to simplify the procedure.
3. When the thinned bone is soft and transparent, stop drilling. When a sufficiently large indentation forms in the bone, frequently stop the rotation and monitor the thickness of the bone. The appearance of cracks in the bone indicates that thinning is sufficient.
4. Bathe the hole with sterile saline and then remove the excess saline with a cotton swab. Remove the remaining layer of bone using a 27G needle with a hook-shaped tip and/or using fine-tipped tweezers. Avoid damaging the dura.
5. Cover the surface of the skull with a sterile piece of paper towel moistened with saline. Place a piece of clean, transparent film on the mouse skull surface above the paper.
6. Move the stereotaxic arm back and position the microsyringe needle in place directly above the film. Dispense the excess oil until a volume of 2 µL is reached.
   NOTE: The final volume of the oil may vary according to the volume of the micro syringe. We used a 5 µL 75-RN Hamilton micro syringe.
7. Pipette a volume of virus equal to the injected volume + 2 µL onto a piece of transparent film.
8. Looking through the surgical microscope, lower the arm down until the tip of the needle is in the center of the drop of the virus. Load the virus into the microsyringe using a motorized injector. Dispose of the tip of the micropipette and transparent film into the bottle with 2% bleach.
9. Remove the paper moistened with saline from the surface of the skull and dry the skull with a cotton swab. Use the stereotaxic arm to position the needle above the insertion site.
10. Dispense a drop of the virus to make sure that the needle is not clogged. Make a small slit in the dura using a 30 G needle with a hook-shaped tip.
    ​NOTE: It is important to make the smallest possible slit in the dura mater, allowing the needle to enter without leaving a gap between the needle and the dura mater, from which the virus could leak.
11. Lower the needle of the syringe down to the dura and make the appropriate calculations for depth. Estimate the cortical depth of the insertion relative to the point at which the needle first touched the surface of the cortex.
12. Slowly insert the needle tip into the cortex to a depth of 300 µm. Wait 2-3 min to allow the dura to adhere to the needle, and then slowly retract the needle to a depth of 200 µm. Wait another 2-3 min to allow the brain tissue to settle. This results in the needle pulling the dura up, creating a subdural space for the virus to spread.
    NOTE: To minimize cortical damage, use a 33 G needle or lower G needles. Keep in mind that the smaller the tip's inner diameter is, the higher the chance of tissue backflow clogging it. It is important to use a blunt needle instead of beveled because a blunt needle expels a more controlled drop of viral solution.
    1. If the needle does not pass through the dura mater when lowered to 150-200 µm and instead bends it, stop lowering. Lift the needle and make a slightly larger incision in the dura mater, then try lowering the needle again.
    2. If cerebrospinal fluid starts leaking actively from the incision while lowering the needle, wait until it stops leaking; otherwise, it is hard to control the needle passing through the dura. Blot the excess cerebrospinal fluid with a tissue tip. Once the fluid stops flowing, lift the needle and try again.
13. Begin injecting the viral suspension at a rate of 0.06 µL/min while monitoring the volume dispensed. Inject 1 µL of virus. Inject the virus at a single depth in order not to break the seal between the needle and cortex tissue.
    1. Troubleshooting: At the beginning of the injection, virus backflow to the brain surface may occur. Stop injecting, wait 2-3 min, and then continue the virus injection. Repeat these actions (steps) until the virus backflow stops. Virus and cerebrospinal fluids will stick to the needle and prevent backflow.
    2. Another way to stop virus backflow, put a small amount of agarose on the surface of the cortex. It will suppress the pulsation of the cortex and will also help to seal the needle. But make sure that the pipette is not clogged by the agarose.
    3. Since the needle may not completely pass through the dura mater when lowered, but only partially, one side of the needle may not fit tightly against the dura, which can cause backflow of the virus. In this case, slowly withdraw the needle from the tissue and try lowering it again.
14. Once the infusion is complete, keep the needle at the target location for an additional 10 min. This allows the virus to disperse away from the injection site. Retract the needle out of the brain slowly to prevent the virus from flooding back out of the needle tract.
15. Once the needle withdrawal is completed, check that it is not clogged by dispensing a small drop of virus.Close the skin incision using 5-0 absorbable or non-absorbable sutures.

5. Post-operative care

1. After closing the incision, apply antibacterial ointment topically. Inject a mix of saline (5 mL/kg) and 5% glucose (5 mL/kg) subcutaneously to prevent dehydration and facilitate recovery after anesthesia. Inject ketoprofen intramuscularly (2.5 mg/kg) to reduce pain.
2. Place the animal in a fresh cage on top of a heating pad and monitor until it recovers from anesthesia. After the mouse is reactive, place the animal in their home cage.

6. Histology

1. No earlier than 21 days after virus injection deeply anesthetize mice with isoflurane as described in step 3.
2. Make a midline incision (10 -15 mm) along the thoracic region using narrow scissors and expose the chest cavity. Carefully separate the diaphragm and open the chest using scissors.
3. Insert a 22G 1 1/2 needle into the left ventricle and make an incision in the right atrium with scissors. Perfuse with 50 mL of pre-chilled 100 mM Phosphate Buffered Saline (PBS), followed by 100 mL of pre-chilled 10% buffered formalin.
4. Carefully extract the entire mouse brain. To do this, make a sagittal incision along the center of the scalp using scissors. Next, remove the cranial bone piece by piece using bone pliers with flat tips starting at the intersection of the sagittal and lambdoid sutures and working forward to the nasal bone. When the brain is exposed, cut through the olfactory bulbs, ease the brain out with a curved spatula, and drop it into a 50 mL tube containing 10% buffered formalin.
5. Fix the brain overnight. Mount the brain tissue on a metal platform of a Vibratome using tissue adhesive. Cut frontal sections at a thickness of 50 µm using carbon steel blades.

7. Immunostaining

1. Day 1
   1. Wash the brain slices 3x for 5 min each with 1x PBS containing 0.3% Triton X-100 (PBS-T). Incubate the sections in PBS-T for 20 min at room temperature (RT).
   2. Block nonspecific binding for 1 h at RT using a blocking buffer composed of 5% Normal Goat Serum (NGS) and 0.3% Triton X-100 in 1x PBS.
   3. Incubate the sections overnight at 4 °C with primary antibodies against parvalbumin or calbindin diluted 1:500 in the blocking buffer (5% NGS and 0.3% Triton X-100 in 1x PBS).
2. Day 2
   1. Wash the sections 3x for 10 min in PBS-T. Incubate in the dark for 2 h at RT with secondary antibodies (Goat anti-Rabbit IgG (H+L) Cross-Adsorbed Secondary Antibody, Alexa Fluor 546) diluted 1:500 in the blocking buffer (5% NGS and 0.3% Triton X-100 in 1x PBS).
   2. Wash the sections 3x for 5 min each in 1x PBS. Transfer brain slices onto glass slides using a soft brush.
   3. Immediately add 0.1 mL of antifade mounting medium to each section. Cover the slices with a 22 mm x 50 mm coverslip.
   4. Configure the confocal microscope to a magnification of 20x or 60x (A/1.4, oil). Use 488 nm and 594 nm wavelength lasers to acquire multi-channel images of the brain regions of interest.

Results

In a pilot series of experiments, we used the traditional intracortical injection method to transduce layer five pyramidal neurons in the mouse neocortex by AAV2 carrying the fast channelrhodopsin (oChIEF) gene fused with EGFP fluorescent protein under the CaMKII promoter. Consistent with the characteristic feature of AAV2¹², we obtained a relatively small area of infection, not exceeding 1 mm in width (Figure 1A). However, in some experiments, we observed un...

Discussion

We have developed a new method for transducing mouse neocortical neurons by injecting a suspension of AAV2 viral particles into the subarachnoid space of the brain. This provides widespread virus distribution, almost four-fold greater than the tissue volume infected when the same amount of virus is injected directly into the brain parenchyma.

Injection of virus vectors directly into the cerebrospinal fluid (CSF) via different routes (e.g., intracerebroventricular, intrathecal, or intracisterna...

Materials

Name	Company	Catalog Number	Comments
Equipment
10 µL Gastight Syringe Model 1701 RN (5 uL 75 RN Hamilton microsyringe)	Hamilton Company	Part/REF # 7634-01, Hamilton or cat no. HAM7634-01, Merck
33 G RN needle, point style 3	Hamilton Company	Part/REF # 7803-05, Hamilton
Binocular Microscope	Nikon or Micromed	Model MC-4 ZOOM
Cerna-based laser scanning confocal microscope	ThorLabs
Cold light source	RWD	Model 76312
Leica VT1000 S Vibrating blade microtome	Leica Biosystems	76001-014
Low-Flow Anesthesia System with starter kit	Kent Scientific Corporation	13-005-111 (Model SomnoSuite)
Mechanical Pipette 0.1 – 2.5 µL Eppendorf Research plus	Eppendorf	3123000012
Mechanical Pipette 10 – 100 µL Eppendorf Research plus	Eppendorf	3123000047
Mice Shaver	RWD	Model CP-5200
Microdrill with drill bits (0.5 mm, round)	RWD	78001, 78040
or Desctop Digital Stereotaxic Instrument, Mouse anesthesia Mask, Mouse ear bars (60 Deg)	RWD	Models 68027, 68665, 68306
Pressurized air	KUDO
Single Channel Manual Pipette 0.5-10 µL	RAINN	17008649
Small Animal Stereotaxic Instrument	KOPF	Model 962
Stereotaxic Injector	Stoelting	10-000-004
Surgical Instruments (Tools)
30 G dental needle (Ni-pro)	Biodent Co. Ltd.		To slit the dura
Bone scraper	Fine Science Tools	10075-16
Dental bur	DRENDEL + ZWEILING		For craniotomy; Shape: pear shaped/round end cylinder/round; Tip Diameter: 0.55-0.8 mm diameter
Needle holder (Halsey Micro Needle Holder)	Fine Science Tools	12500-12
Polypropylene Surgical Suture or Surgical Suture Vicryl (5-0, absorbable)	Walter Products (Ethicon)	S139044 (W9442)
Scalpel handle (#3) with scalpel blades (#11)	Fine Science Tools	10003-12, 10011-00
Scissors (Extra Narrow Scissors)	Fine Science Tools	14088-10	to cut the skin
Scissors (Fine Scissors)	Fine Science Tools	14094-11	to cut suture
Surgical suture PROLENE (Polyproptlene)	Ethicon (Johnson & Johnson)
Tweezers (Forceps #5)	Fine Science Tools	11252-20
Tweezers (Polished Inox Forceps)	Fine Science Tools	11210-20
Disposables
1 mL insulin syringe	SITEKMED		To load vaseline oil into a microsyringe, to administer drugs
Cell Culture Plate	SPL Life Science
Cotton swabs
Cover Glasses	Fisher Scientific	12-545E
Insulin syringe needle (27 G)	SITEKMED		To remove debries from a hole (craniotomy)
Lint-free wipes CLEANWIPER	NetLink
Microscope Slides	Fisher Scientific	12-550-15
Paper towels	Luscan
Parafilm	StatLab	STLPM996
Sterile Surgical Gloves	Dermagrip
Drugs/Chemicals (Reagents)
10% buffered formalin or 4% paraformaldehyde	Thermo Scientific Chemicals	J61899.AK
Alcohol solution of iodine (5%))	Renewal
Antibiotic ointment Baneocin (bacitracin + neomycin)	Sandoz		Antibacterial agent for external use
Aqua Polymount	Poly-sciences	18606-20
Carbomer Eye Gel Vidisic (Ophthalmic gel)	BAUSCH+LOMB (Santen)
Carboxylate-Modified FluoSphere Microspheres (red)	Thermo Fisher Scientific	F-8801
Dexamethasone (4 mg/mL)	Ellara (KRKA)		Synthetic glucocorticoid
Distilled H2O
Ethanol (70%)
Flexoprofen 2.5% (Ketoprofen)	VIC		Nonsteroidal Anti-Inflammatory Drugs (NSAIDs)
Glucose solution 5%	Solopharm
Goat anti-Rabbit IgG (H+L) Cross-Adsorbed, Alexa Fluor 546	Thermo Fisher Scientific	A-11010
Isoflurane	Karizoo
lidocaine solution (2 % / 4%)	Solopharm
Normal Goat Serum (NGS)	Abcam	ab7481
Phosphate Buffered Saline (PBS)	Eco-servis
Rabbit Anti-Parvalbumin Antibody	Merck Millipore	AB15736
Rabbit Recombinant Monoclonal anti-Calbindin antibody	Abcam	ab108404
Saline (0.9% NaCl in H2O)	Solopharm
Triton X-100	Sigma-Aldrich	50-178-1844
Vaseline oil	Genel