Consistent Delivery of Adeno-Associated Virus via Lateral Tail-Vein Injection in Adult Mice

Fady Guirguis; V&#233;ronique Bolduc; Matthew J. Slarve; Haiyan Zhou; Francesco Muntoni; Carsten G. B&#246;nnemann

doi:10.3791/66934

JoVE Journal > Medicine

Medizin

Consistent Delivery of Adeno-Associated Virus via Lateral Tail-Vein Injection in Adult Mice

Published: August 23, 2024

doi:

10.3791/66934

Fady Guirguis^1,2, Véronique Bolduc¹, Matthew J. Slarve³, Haiyan Zhou^4,5, Francesco Muntoni^2,4, Carsten G. Bönnemann¹

¹Neuromuscular and Neurogenetic Disorders of Childhood Section, National Institute of Neurological Disorders and Stroke,National Institutes of Health, ²The Dubowitz Neuromuscular Centre, Molecular Neurosciences Section, Developmental Neurosciences Research and Teaching Department, Great Ormond Street Institute of Child Health,University College London, ³Department of Molecular Microbiology and Immunology,University of Southern California, ⁴NIHR Great Ormond Street Hospital Biomedical Research Centre, ⁵Genetics and Genomic Medicine Research and Teaching Department, Great Ormond Street Institute of Child Health,University College London

Summary

Here we detail an optimized protocol for mouse lateral tail-vein injection to systemically administer adeno-associated virus (AAV) in adult mice. Additionally, we describe protocols of commonly used assays to assess AAV transduction.

Abstract

Many disorders affect multiple organs or involve different regions of the body, so it is critical to deliver therapeutics systemically to target the affected cells located in different sites. Intravenous injection is a widely used systemic delivery route in preclinical studies that assess treatments intended for body-wide administration. In adult mice, it involves the intravenous administration of the therapeutic agent into the mouse's lateral tail veins. When mastered, tail-vein injections are safe and fast, and only require simple and commonly available tools. However, tail-vein injections are technically challenging and require extensive training and continuous practice to ensure the accurate delivery of the intended dose.

Here we describe a detailed, optimized, lateral tail-vein injection protocol that we have developed based on our experience and on recommendations that had been previously reported by other groups. Other than the mouse restrainers and insulin syringes, this protocol requires only reagents and equipment that are readily available in most labs. We found that following this protocol results in consistently successful intravenous delivery of adeno-associated virus (AAV) into the tail veins of unsedated 7-9 week-old mice. Additionally, we describe the optimized protocols for the histological detection of fluorescent reporter proteins and vector genome per diploid genome (vg/dg) quantification used to assess AAV transduction and biodistribution. The goal of this protocol is to aid experimenters in easily performing tail-vein injections successfully and consistently, which can reduce the practice time needed to master the technique.

Introduction

Monogenic disorders make up 80% of rare diseases, which collectively affect 300 million individuals worldwide¹^,². There are currently no approved curative therapies for the majority of these greatly debilitating rare disorders¹^,²^,³. However, monogenic disorders are ideal candidates for gene therapies that can replace, supplement, correct, or silence dysfunctional genes⁴^,⁵. Currently, multiple vectors are being developed and used to deliver gene therapies to specific cell types⁴^,⁶. One of those vectors is adeno-associated virus (AAV). AAV is a non-pathogenic parvovirus that is increasingly being used as a gene therapy vector⁷. Compared to other viral vectors, AAV has lower immunogenicity, lower potential to integrate into the host genome, and the ability to efficiently transduce dividing and non-dividing cells in various tissues⁷^,⁸. Additionally, multiple approaches have been developed to engineer and identify AAVs with desirable characteristics such as specific tissue tropism or further reduced immunogenicity, which greatly enhances AAV's versatility as a viral vector for different indications⁹. These factors have made AAV a widely investigated gene therapy vector and led to the development of multiple FDA-approved AAV-based gene therapies¹⁰.

Mouse models are commonly used to test potential gene therapies in vivo and better understand the pathomechanisms of monogenic disorders. This is due to the mouse models' recapitulation of the pathologies of different conditions, their genome's similarity to the human genome, and the relative ease of mouse handling, maintenance, and generation¹¹^,¹²^,¹³. In vivo testing is particularly important when studying disorders that affect multiple systems or regions of the body, such as muscular dystrophies. For these disorders, in vitro testing might not be sufficient to comprehensively assess the safety, efficacy, pharmacokinetics, and pharmacodynamics of therapeutics intended to reach different body regions after systemic administration¹⁴.

Various systemic administration routes can be used to deliver drugs. Each route has its advantages, drawbacks, and degree of compatibility with the animal model and drug being investigated¹⁵. Intravenous (IV) lateral tail-vein injection is a commonly used route for systemic delivery of AAV in mice¹⁶. Lateral tail-vein injections allow fast and direct administration of the injectate into the mouse bloodstream ensuring high drug bioavailability in systemic circulation¹⁷. They also require relatively simple and commonly available tools to be performed. However, mainly due to the small tail vein diameter and difficulty in locating the vein, lateral tail-vein injections are technically challenging and require a high degree of skill and constant practice to avoid failed injection attempts or incomplete dose delivery¹⁶^,¹⁷^,¹⁸^,¹⁹. These can result in the loss of expensive reagents or inaccurate results, especially if the incomplete injection is not recognized while performing the injection. Our experience summarized here is based on protocols reported in well-documented articles that we have adapted for our use, optimizing various steps of the lateral tail-vein injection procedure to ensure consistently successful injections²⁰^,²¹^,²²^,²³^,²⁴^,²⁵^,²⁶^,²⁷.

Here, we describe this detailed optimized lateral tail-vein injection protocol to deliver AAV into unsedated 7-9-week-old mice using simple and commonly available tools. Additionally, we provide the protocols for methods used to assess AAV delivery and biodistribution. These protocols cover post injection tissue collection, tissue fixation, DNA extraction, and digital polymerase chain reaction (dPCR) vector genome per diploid genome (vg/dg) quantification. The IV injection protocol and pointers provided here aim to enhance the ease of successfully performing lateral tail-vein injections. This will potentially help reduce the time needed to master the injection skills while simultaneously improving the accuracy and consistency of injections.

Protocol

All animal handling and injection procedures were approved by the Animal Care Committee at NINDS. All animal procedures were conducted in compliance with the NINDS animal care and use guidelines.

1. Preinjection preparation

AAV dose preparation
1. Determine the average weight of the mice that will be injected.
2. Calculate the maximum allowed injectate volume according to the institution's animal care guidelines as shown in equation (1). The maximum injectate volume is typically a volume (µL)/mouse weight (g) value (e.g., 10 µL/g.).
  Maximum injectate volume (µL)/mouse= (maximum injectate volume (µL/g)) × (average mouse weight (g)) (1)
  NOTE: Sample Calculation: Maximum injectate volume/mouse= 10 µL/g × 20 g/mouse= 200 µL/mouse
3. Set the AAV vector genome (vg) dose to be delivered per mouse.
  NOTE: This could be the same absolute value across different mice (e.g., all mice receive 1.5 × 10¹² vg regardless of how much each mouse weighs). Or the dose could be in vg/kg, so the total vg to be injected per mouse must be calculated for each mouse according to that mouse's weight on the injection day.
  1. If the dose is in vg/kg, weigh each mouse on the injection day prior to dose preparation.
  2. Calculate the vector genomes to be delivered for each mouse according to its weight using equation (2):
    Vector genomes to be delivered in a specific mouse (vg) = Prespecified vg/kg value (vg/kg) × (2)
    NOTE: Using vg/kg as the dose unit instead of vg/mouse might be more appropriate in certain preclinical studies to ensure valid comparisons between injected doses. This is due to the weight differences between male and female mice of the same age or possibly between mice of the same sex.
  3. Use the maximum injectate volume and the AAV (vg) dose to calculate the volumes of stock AAV and sterile phosphate-buffered saline (PBS) that are needed to prepare the required dose (see equations (3-6)). Ensure that the volume to be injected is equal to or less than the allowed maximum injectate volume. Always prepare an injectate volume that is at least 15 µL greater than the volume that will be injected to account for pipetting errors and the syringe dead space.
    Concentration of injectate (vg/µL) = (3)
    Total AAV vector genomes to be added to prepare the injectate (vg) = Concentration of injectate (vg/µL) × volume to be prepared (µL) (4)
    Volume of AAV stock to be added to prepare the injectate) (µL) = (5)
    Volume of PBS to be added to prepare the injectate (µL) = Volume to be prepared (µL) – Volume of AAV stock to be added to prepare the injectate (µL) (6)
    NOTE: Sample calculation:
    6 × 10¹³ vg/kg (the dose) will be delivered in 200 µL/mouse (the volume to be injected) in a mouse that weighs 25 g. AAV stock titer is 3.0 × 10¹³ (vg/mL)
    Vector genomes to be delivered in this specific mouse (vg) = 6 × 10¹³ (vg/kg) × = 1.5 × 10¹² vg for this mouse
    Concentration of injectate= = 7.5 × 10⁹ (vg/µL)
    Total AAV vector genomes to be added to prepare the injectate = 7.5 × 10⁹ (vg/µL) × (200 (µL) + 15 (µL)) = 1.6125 × 10¹² (vg)
    Volume of AAV stock to be added to prepare the injectate = = 53.75 (µL)
    Volume of PBS to be added to prepare the injectate = 215 (µL) – 53.75 (µL) = 161.25 (µL)
4. AAV dose preparation procedure
  NOTE: Following the institution's biosafety and PPE guidelines for handling AAV, in an autoclaved sterile RNase-free and DNase-free 1.7 mL microcentrifuge tube, prepare the AAV injectate using the AAV stock and sterile PBS according to the calculations in step 1.1.3.3. Always keep the stock AAV and AAV injectate on ice. Use clean micropipettes and new micropipette tip boxes to ensure sterility. Discard the AAV-contaminated micropipette tips according to the institution's waste management guidelines.
  1. Thaw the stock AAV on ice.
    NOTE: Avoid thawing and refreezing the AAV stock. Either order or prepare the AAV stock in aliquots of 100-200 µL to avoid having excess AAV after dose preparation that will need to be refrozen.
Injection station preparation
1. Cleaning
  1. Clean the working area with 70% ethanol (EtOH).
  2. Disinfect the working area using a bactericidal, fungicidal, and virucidal reagent(s).
  3. Clean the mouse tube restrainer with water and soap.
2. Station tools setup
  1. Place a clean empty 15 mL conical tube in a tube holder/rack.
  2. Set up an elevated platform on which the mouse tube restrainer will be placed (Figure 1A,B).
  3. Place the clean mouse tube restrainer in the working area.
  4. If injecting mice that are much smaller than the available mouse tube restrainer, use a plastic rodent restrainer cone to make a restraining sleeve. See steps 2.1.3-4.
  5. Place the syringes that will be used for injections in the working area. Use 0.3 mL insulin syringes with 29 G needles.
  6. Place the waste container as close as possible to the injection station to allow immediate disposal of AAV-contaminated tools.
  7. Pull and push the plunger of each syringe multiple times to make sure the plunger moves smoothly so there is no syringe-caused resistance during the injection. If the plunger does not move smoothly, discard this syringe and replace it with a new one.
  8. Have a mouse scale and mini centrifuge available next to the injection area.
  9. Have the prepared AAV on ice ready in the injection area.
  10. Prepare warm water (38-40 °C). Take care that the water temperature does not exceed 40 °C to avoid causing burns to the mouse tail.

2. Injection procedure

Mouse restraining
1. Weigh each mouse to calculate the dose in vg/kg if needed.
2. Ensure that the mouse is fully restrained and not able to move.
  NOTE: If the mouse is not fully restrained, it might move during the IV administration resulting in the displacement of the needle. This might cause the needle to get out of the vein and/or injure the mouse.
3. For tube restrainer:
  1. Wash the restrainer with warm water and soap between mice to clean and warm up the holder.
  2. Hold the mouse by its tail. Insert the mouse's tail into the tube's top opening; then, slowly pull the mouse into the tube restrainer. If the tube restrainer's size is suitable for the mouse's size, place the plug ahead of the mouse to prevent the mouse from escaping.
    NOTE: The plug should be close enough to the mouse to prevent the mouse from moving or rotating inside the tube, but the plug should not obstruct the mouse's nose to allow the mouse to breathe freely. If the mouse is smaller than the tube restrainer size, use a flexible disposable restrainer cone in addition to the tube restrainer as described below in step 2.1.4.
4. For the flexible disposable restrainer cones if needed (to make a restraining sleeve for smaller mice):
  1. Make a cut at the nose end of the cone to make sure the mouse's nose is not obstructed and the mouse has enough room to breathe.
  2. Cut the back of the cone so that the cone is almost the same length as the mouse (so the cone will fit inside the tube) (Figure 1A,B).
  3. Place the mouse in the tube restrainer as described above in steps 2.1.3. While holding the tail of the mouse, insert the restrainer cone with the wider opening side first into the tube.
  4. While holding the tail of the mouse, let the mouse walk into the cone; then, slide the rest of the cone into the tube. Ensure that the tail of the mouse is all the way out of the back of the tube, and the mouse has room to breathe inside the cone.
  5. Secure the tube plug immediately in front of the cone's nose opening while making sure that the mouse is fully restrained and has enough room to breathe.
Injecting the mouse
1. Fill the 15 mL conical tube with warm water.
2. Place the tube restrainer with the mouse inside it on the elevated platform (Figure 1B).
3. Dip as much as possible of the restrained mouse's tail in the warm water for at least 1 min until the lateral veins are clearly dilated and visible (Figure 1B).
4. During the tail-warming step, load the AAV dose into the syringe.
  1. Place the uncapped AAV-containing 1.7 mL microcentrifuge tube in a tube rack. Insert the needle vertically into the tube with the dominant hand. Once the needle is inside the tube, hold the tube with the non-dominant hand.
    NOTE: The vertical insertion of the needle prevents damage to the needle that might be caused by touching the tube wall. Other alternative syringe-loading methods can be used, but the safety of the experimenter and injected mouse should be ensured. Holding the tube with the non-dominant hand protects against accidental needle puncture injuries if the needle is inserted while holding the tube.
  2. With the needle inside the tube, raise the tube and the syringe simultaneously to eye level while making sure the needle does not touch the tube wall. Rest both arms on the table to stabilize them. Slowly pull the dose into the syringe.
    NOTE: Slow aspiration keeps fine air bubbles from attaching to the syringe barrel sides.
  3. Expel air bubbles from the syringe. If injecting AAV, make sure the air bubbles are expelled from the syringe over a disposable absorbent pad that will be disposed of in a biohazard box.
  4. Hold the injectate for at least 40 s to warm it up.
    NOTE: Always ensure that the injectate is warm before injection. If cold injectate is administered, the injection might not flow through the vein except the first few µL.
  5. Check the tail veins every minute.
    NOTE: The veins must be VERY visible all the way to the injection site (add tail-warming time and replace with fresh warm water as needed until the vein is clearly visible but not to exceed the mouse restraining time allowed by the institution's animal handling protocol).
  6. After ensuring that the veins are clearly visible, remove the tube restrainer from the top of the elevated platform and place the tube restrainer directly on the table. Position the mouse in the restrainer with its feet downwards and not to the side for easy handling of the tail.
    NOTE: The mouse should not be on its side or back inside the restrainer. The mouse should be fully restrained and unable to move or rotate inside the restrainer or move/pull its tail.
  7. Quickly wipe the tail with a gauze to dry the tail; wipe the tail with an alcohol swab and then, wipe it dry with a dry gauze.
    NOTE: Use dry gauze to make the tail dry enough to allow secure gripping of the tail but not completely dry. It can be more difficult to see the vein when the tail is completely dry.
  8. Rotate the tail approximately 90° to the left or the right so one of the two lateral veins is facing upwards. Locate a suitable injection site within the middle third of the tail. Start the initial injection distally (closer to the tip of the tail) and move proximally if additional injections are required due to failed attempts or necessitated by the experimental design.
    NOTE: Do not attempt to inject distal to a previous injection site since the injectate might leak out of that previous injection site. Optional: use the non-dominant hand's thumb and index finger to apply pressure proximal (upstream/closer to the mouse body) to the injection site for 10 s. The fingers act as tourniquets to further dilate the vein at the injection site. Immediately after the 10 s tourniquet, release the tourniquet fingers and ensure that one of the two lateral veins is facing upwards and is clearly visible.
  9. Hold the tail with the non-dominant hand using the thumb and index finger immediately distal to the injection site. Fold the tail over the index finger so the injection site lies flat on the index finger. Pull back on the tail so the tail is stretched and the injection site is completely horizontal (at 0°) (parallel to the horizontal table) (Figure 1C).
  10. Hold the syringe using the index and middle fingers of the dominant hand on either side of the syringe's barrel flange while keeping the thumb ready at the plunger.
    NOTE: This will make it easier not to move the thumb or the needle once the needle is inside the vein (Figure 1C).
  11. Rest both hands on the table to stabilize them and place the needle directly on and parallel to the tail and the vein with the bevel facing upwards. Keep the injection site close to the index finger holding the tail to improve control and stability of the injection site.
    NOTE: The tube restrainer should be deep enough in the table so both hands are supported on the table.
  12. While keeping the needle parallel to the tail vein and applying downward pressure on the needle, slide the needle forward into the vein.
    NOTE: The downward pressure should be enough to insert the needle at the correct angle into the vein. The vein is extremely shallow, so the needle should be as flat as possible when trying to get into the vein.
  13. Slowly inject the solution into the vein. After administering the dose, slowly withdraw the needle and immediately apply pressure with a gauze at the injection site for at least 10 s to stop bleeding.
    NOTE: Apply the pressure as long as needed until the bleeding has completely stopped to avoid the potential loss of the injected reagent. A blood drop usually appears after withdrawing the needle indicating that the needle has penetrated the vein. Occasionally, the blood drop does not appear even with a successful injection. The blood drop does not indicate successful injection; it only indicates that the needle penetrated the vein. The reliable successful injection indicator is the complete absence of plunger resistance during the injection. If the needle is inside the vein, there should be no resistance at the needle plunger during the injection of the injectate, and the vein proximal to the injection site will momentarily appear slightly lighter in color (blanches) (the vein blanching might not be very clear in some mouse strains). If there is resistance and/or a bulge starts to appear at the injection site, then the needle is not correctly placed inside the vein. If this occurs, completely remove the needle from the tail and attempt to inject the vein at a new injection site proximal to the failed injection site (closer to the mouse body).
  14. Discard the AAV-contaminated syringes and tubes according to the institution's waste management guidelines.
  15. Free the mouse from the restrainers and place it back in a new cage separate from uninjected mice. Monitor the mice for 10 min to ensure normal activity levels post injection.
    NOTE: This avoids potential transmission of injected agents to uninjected mice if transmissible agents are administered.
  16. Disinfect the working area using bactericidal, fungicidal, and virucidal reagent(s) and 70% EtOH. Clean the mouse tube restrainer with water and soap.

3. Dissection and tissue collection and fixation²⁷

Tissue collection station preparation
1. Clean the working station with a DNA degradation reagent according to the manufacturer's instructions to degrade contaminating DNA that might be present in the working area.
2. Place methylbutane in a metal container. Place the methylbutane metal container inside a Styrofoam box; then, surround the metal container with dry ice so the level of the dry ice surrounding the container is higher than the methylbutane level inside the container.
3. Label and place empty 2 mL microcentrifuge tissue storage tubes on the dry ice. Leave the methylbutane and tissue storage tubes to cool on dry ice for at least 20 min before beginning to freeze the tissues. Place tissue transfer forceps on dry ice.
4. Label and fill another set of 2 mL microcentrifuge tissue storage tubes with fresh 4% paraformaldehyde (PFA) and keep them at room temperature. Add enough 4% PFA to each tube to completely submerge the tissues that will be placed in the tube.
Tissue collection and fixation
1. Euthanize the mouse according to the institution's animal care guidelines.
  NOTE: Here, the mice were euthanized using cervical dislocation.
2. Fully spray the mouse with 70% EtOH.
3. Collect the required tissues.
  NOTE: The collection and fixation protocol described here was tested on skeletal muscles and the liver.
4. For tissues that will be used for DNA extraction:
  1. Drop the tissue in methylbutane prechilled on dry ice and leave the tissue in methylbutane for at least 1 min. Use the prechilled transfer forceps to transfer the frozen tissues from the methylbutane into the prechilled empty 2 mL microcentrifuge tissue storage tubes. Store the tissue at -80 ˚C.
    NOTE: Optional: The tissue can be cut into 20 mg pieces before dropping in methylbutane to be ready to be used in step 4.1.4.
5. For tissues that will be used for histological analysis and for preserving reporter proteins' fluorescence:
  1. Using a PFA-designated forceps kept at room temperature, drop the tissues in their respective microcentrifuge tube that contains 4% PFA (kept at room temperature) while making sure the tissue is completely submerged in the 4% PFA solution.
    NOTE: PFA contamination can negatively affect different downstream molecular assays. Only use PFA-designated forceps when handling PFA to prevent PFA contamination of other tissues or tools.
  2. Place the microcentrifuge tubes on a rack and cover the rack with foil paper to keep the tubes in the dark. Incubate the covered rack at 4 ˚C on a shaker with gentle shaking overnight.
  3. After overnight incubation, prepare 5% sucrose (% w/v) in 1x PBS by dissolving 5.0 g. of sucrose in 70 mL of 1x PBS by vigorous shaking. Add enough 1x PBS to a final total volume of 100 mL to achieve a 5% sucrose solution (% w/v).
  4. Sterilize the 5% sucrose solution using the 0.22 µm syringe filter. Label and fill 2.0 mL microcentrifuge tubes with freshly prepared 5% sucrose.
  5. Transfer the tissues from 4% PFA to their respective microcentrifuge tube that contains 5% sucrose (kept at room temperature) while making sure the tissue is completely submerged in the 5% sucrose solution.
  6. Place the microcentrifuge tubes on a rack and cover the rack with foil paper to keep the tubes in the dark. Incubate the covered rack at 4 ˚C on a shaker with gentle shaking overnight.
  7. After overnight incubation, prepare 20% sucrose (% w/v) in 1x PBS by dissolving 20.0 g. of sucrose in 70 mL of 1x PBS by vigorous shaking. Add enough 1x PBS to a final total volume of 100 mL to achieve a 20% sucrose solution (% w/v).
  8. Sterilize the 20% sucrose solution using a 0.22 µm syringe filter. Label and fill 2.0 mL microcentrifuge tubes with freshly prepared 20% sucrose.
  9. Transfer the tissues from 5% sucrose to their respective microcentrifuge tube that contains 20% sucrose (kept at room temperature) while making sure the tissue is completely submerged in the 20% sucrose solution.
  10. Place the microcentrifuge tubes on a rack and cover the rack with foil paper to keep the tubes in the dark. Incubate the covered rack at 4 ˚C on a shaker with gentle shaking overnight.
  11. After overnight incubation, place methylbutane in a metal container and place the methylbutane metal container inside a Styrofoam box. Surround the metal container with dry ice so the level of the dry ice surrounding the container is higher than the methylbutane level inside the container.
  12. Label and place empty 2 mL microcentrifuge tissue storage tubes on dry ice. Leave the methylbutane and tissue storage tubes to cool on dry ice for at least 20 min before beginning to freeze the tissues. Place transfer forceps on dry ice.
  13. Quickly blot the tissues using precision wipes to remove any excess 20% sucrose. Drop the tissue in methylbutane prechilled on dry ice. Leave the tissue in methylbutane for at least 1 min.
  14. Use the prechilled transfer forceps to transfer the frozen tissues from the methylbutane into the prechilled empty 2 mL microcentrifuge tissue storage tubes. Store the tissue at -80 ˚C.

4. dPCR for vg/dg quantification

DNA extraction from tissues and initial RNA digestion
NOTE: The handbook of the DNA extraction kit listed in the Table of Materials was used to derive this DNA extraction protocol. Always keep the tubes containing the frozen tissue pieces on dry ice.
1. Prepare a bucket of ice.
2. For each DNA sample, label one 1.5 mL lysis bead tube and two empty RNase-free and DNase-free 1.7 mL microcentrifuge tubes.
3. Add 180 µL of the first DNA extraction kit buffer to each bead tube. Tare the first bead tube containing buffer.
  NOTE: If the tissues were not precut in step 3.2.4.1, use a razor blade prechilled on dry ice to cut the tissue into 20 mg. pieces. This step must be done inside a clean cryostat kept at -20 ˚C or colder.
4. Add one piece of tissue into the tube; weigh and record the weight of the tissue (to be ~20 mg.).
5. Immediately place the lysis bead tube with the tissue in it on ice. The buffer might crystallize.
6. Repeat the previous steps for each tissue sample.
7. Transfer the tubes to the lysis bead tube blender and run for 1 min at maximum speed (speed 10) at 4 ˚C.
8. Place the samples on ice to transfer them to the centrifuge. Centrifuge for 1 min at 20,000 × g at 4 ˚C.
9. During the centrifugation step, add 20 µL of proteinase K to the first series of 1.7 mL microcentrifuge tubes. After the centrifugation step, transfer the supernatant of the homogenates to the 1.7 mL tubes containing the proteinase K and mix well. Incubate at 56 ˚C for 15 min, with 500 RPM mixing.
10. Collect the drops from the walls and lid of the tube by centrifuging the tube for 1-2 s using a mini centrifuge. Incubate at room temperature for 2 min.
11. Add 4 µL of RNase A and mix by brief pulse vortexing. Incubate at room temperature for 2 min. Pulse-vortex for 15 s.
12. Collect the drops from the walls and lid of the tube by centrifuging the tube for 1-2 s using a mini centrifuge. Add 200 µL of the second DNA extraction kit buffer. Pulse-vortex for 15 s.
13. Collect the drops from the walls and lid of the tube by centrifuging the tube for 1-2 s using a mini centrifuge. Add 200 µL of 100% EtOH. Pulse-vortex for 15 s.
14. Collect the drops from the walls and lid of the tube by centrifuging the tube for 1-2 s using a mini centrifuge. Transfer the lysates to the DNA extraction spin column. Spin at 6,000 × g for 1 min.
15. Place the spin column in a new collection tube. Add 500 µL of the third DNA extraction kit buffer to the spin column. Spin at 6,000 × g for 1 min.
16. Place the spin column in a new collection tube. Add 500 µL of the fourth DNA extraction kit buffer to the spin column. Spin at 20,000 × g for 3 min.
17. Place the spin column in a new 1.7 mL microcentrifuge tube. Add 100 µL of molecular grade water to the spin column. Incubate at room temperature for 1 min. Spin at 6,000 × g for 1 min at room temperature.
18. Measure the DNA concentration if needed. Store at 4 ˚C for short-term storage or -20 ˚C for long-term storage.
DNA extraction from FACS-sorted cells
NOTE: The handbook of the DNA extraction kit listed in the Table of Materials was used to derive this DNA extraction protocol.
1. After sorting the cells, centrifuge the samples at 300 × g for 5 s to collect all drops on the sides and lid. Ensure all drops are collected.
  NOTE: If the sample volume is less than 1.5 mL, proceed directly to the next step. If the sample volume is greater than 1.5 mL, carefully remove and discard the top portion of the supernatant using a micropipette leaving 1-1.5 mL of the sample.
2. Mix the sample by pipetting up and down multiple times and transfer the sample to a 1.7 mL microcentrifuge tube. Centrifuge at 515 × g for 1 min at room temperature.
3. Discard the supernatant except for the last 50 µL. Resuspend the pellet in 50 µL of the first DNA extraction kit buffer for a final volume of 100 µL.
4. Follow the manufacturer's protocol for the isolation of genomic DNA from small volumes of blood (see Table of Materials).
  1. Add 10 µL of proteinase K and 100 µL of second DNA extraction kit buffer; mix by pulse-vortexing for 15 s. Incubate the samples at 56 °C for 10 min with 300 RPM mixing. Mix the samples twice by gentle inversion during this incubation period.
  2. Collect the drops from the walls and lid of the tube by centrifuging the tube for 1-2 s using a mini centrifuge. Add 50 µL of 100% EtOH and mix by pulse-vortexing for 15 s. Incubate the samples at room temperature for 5 min.
  3. Collect the drops from the walls and lid of the tube by centrifuging the tube for 1-2 s using a mini centrifuge. Transfer the samples to the DNA extraction column (the column is in a 2 mL collection tube) without wetting the rim. Centrifuge at 6,000 × g for 1 min.
  4. After placing the column in a clean 2 mL collection tube, discard the collection tube containing the flowthrough. Add 500 µL of the third DNA extraction kit buffer to the column without wetting the rim and centrifuge at 6,000 × g for 1 min.
  5. Again, after placing the column in a clean 2 mL collection tube, discard the collection tube containing the flowthrough. Add 500 µL of the fourth DNA extraction kit buffer to the column without wetting the rim and centrifuge at 6,000 × g for 1 min.
  6. Place the column in a clean 2 mL collection tube and discard the flowthrough-containing collection tube. Centrifuge at 20,000 × g for 3 min.
  7. Place the column in a clean 1.7 mL microcentrifuge tube and discard the flowthrough-containing collection tube. Add 20 µL of molecular grade water to the center of the column membrane for elution; close the lid and incubate the samples with the molecular grade water at room temperature for 5 min.
  8. Centrifuge at 20,000 × g for 1 min. Store the eluted DNA at 4 ˚C for short-term storage or -20 °C for long-term storage.
RNA digestion and cleanup
NOTE: The handbook of the DNA extraction kit listed in the Table of Materials was used to derive this DNA cleanup protocol. Depending on dPCR conditions, reagents, and primer and probe designs, it might be necessary to ensure the complete absence of RNA in the DNA sample before proceeding to the dPCR vg/dg quantification. RNA contamination might result in various degrees of inaccurate vg/dg values under certain dPCR conditions.
1. In a 0.2 mL PCR tube or a 1.7 mL microcentrifuge tube, add at most 20 µL of the extracted DNA sample and 1.5 µL of the DNase-free RNase to each DNA sample. If the DNA/RNase mix volume is less than 21.5 µL, add enough molecular grade water to a final volume of 21.5 µL and mix 25x by inverting the tubes. Incubate at 37 ˚C for 30 min with periodic mixing every 10 min by inverting the tubes.
  NOTE: The total amount of nucleic acids added to the tube should be between 175 ng and 700 ng. Modifications might be needed if the DNA samples contain volumes or nucleic acid amounts outside of this range or if DNA samples were isolated differently.
2. Place on ice for 2 min. Add enough molecular grade water to each DNA/RNase mix to a final volume of 100 µL.
  NOTE: The RNase listed here is recommended since it digests the contaminating RNA without negatively affecting the target DNA or downstream PCR assays.
3. Follow the manufacturer's protocol for cleaning up of genomic DNA (see Table of Materials).
  1. Add 10 µL of the first DNA extraction kit buffer and 250 µL of the second DNA extraction kit buffer. Mix by pulse-vortexing for 10 s.
  2. Transfer the samples to the DNA extraction column in a 2 mL collection tube without wetting the rim. Centrifuge at 6,000 × g for 1 min.
  3. After placing the column in a clean 2 mL collection tube, discard the collection tube containing the flowthrough. Add 500 µL of the second DNA extraction kit buffer to the column without wetting the rim. Centrifuge at 6,000 × g for 1 min.
  4. Place the column in a clean 2 mL collection tube and discard the flowthrough-containing collection tube. Centrifuge at 20,000 × g for 6 min.
  5. Place the column in a clean 1.7 mL microcentrifuge tube and discard the collection tube containing the flowthrough. Add 20 µL of molecular grade water to the center of the column membrane for elution, close the lid, and incubate the samples with the molecular grade water at room temperature for 5 min.
  6. Centrifuge at 20,000 × g for 1 min. Store at 4 ˚C for short-term storage or -20 °C for long-term storage.
  7. Confirm the absence of RNA contamination in the RNase-digested sample with endpoint PCR or dPCR or quantitative PCR (qPCR) using a PCR primer pair that would amplify an mRNA region that spans multiple exons and not only a single exon.
    NOTE: The mRNA target should be of a gene highly expressed in the target tissue/cell type to ensure that the mRNA contamination will be correctly identified if present. Amplicons that span multiple exons differentiate between bands resulting from contaminating mRNA versus genomic DNA. Always have a non-RNase digested DNA sample as a positive control for the PCR reaction to ensure that mRNA contamination will be detected if present.
Digital PCR (dPCR)
1. End-point PCR to check primers' specificity and optimal PCR conditions (optional)
  1. Design dPCR primer pairs and probes for the vector genome and dPCR primer pairs and probes for the mouse reference gene that will be used to quantify the diploid genomes in the sample.
    NOTE: Aim for amplicon size between 60 bp and 150 bp. The mouse reference gene should be a gene that has a constant gene copy number per diploid genome. For the calculations listed here, the reference gene (Polr2a) has two copies per diploid genome.
  2. For 10 µL end point PCR reaction, prepare the PCR mix using the reagents and final concentrations to be used later for the dPCR reaction. Add the RNase-digested template DNA sample (nucleic acids amount range 56-223 ng) to a final concentration of 1x dPCR master mix that contains the DNA polymerase and dNTPs, 0.8 µM of each forward primer, 0.8 µM of each reverse primer, 0.4 µM of each probe, and 0.025 U/µL of restriction enzyme (final concentration of restriction enzyme depends on the restriction enzyme and brand used). Add molecular grade water to reach a final volume of 10 µL.
    NOTE: There should be at least two primer pairs and two probes in the PCR mix: one primer pair and one probe to detect the vector genome and one primer pair and one probe to detect the mouse genome.
  3. PCR thermal cycling conditions: Initial heat activation step at 95 °C for 2 min, followed by 35-45 cycles of a denaturation step at 95 °C for 25 s and a combined annealing/extension step at 58-62 °C for 1 min.
    NOTE: The optimal annealing temperature must be determined for each amplicon and primer pair. The number of cycles could be adjusted according to the amount of template DNA in the sample.
  4. Visualize the PCR product on an agarose gel using gel electrophoresis to determine the presence of the target amplicon bands and any possible non-specific amplification bands.
  5. Proceed to the next dPCR step after confirming that the primer pairs and cycling conditions result in specific amplification of the target sequences.
2. dPCR reaction
  1. For a 40 µL dPCR reaction, add up to 4 µL of the RNase-digested template DNA sample (nucleic acids amount range 50-330 ng) to a final concentration of 1x dPCR master mix that contains the DNA polymerase and dNTPs, 0.8 µM of each forward primer, 0.8 µM of each reverse primer, 0.4 µM of each probe, and 0.025 U/µL of restriction enzyme (final concentration of restriction enzyme depends on the restriction enzyme and brand used). Add molecular grade water to reach a final volume of 40 µL.
  2. dPCR thermal cycling conditions: Initial heat activation step at 95 °C for 2 min, followed by 40-50 cycles of a denaturation step at 95 °C for 25 s, and a combined annealing/extension step at 58-62 °C for 1 min.
    NOTE: The optimal annealing temperature must be determined for each amplicon and primer pair. The number of cycles could be adjusted according to the amount of the template DNA in the sample. The volumes, concentrations, and conditions listed here are optimized for dPCR plates, reagents, and device listed in the Table of Materials. These conditions reduce the effect of any potential dPCR inhibitors that may reduce the accuracy of the reaction.
  3. After running the dPCR reaction and obtaining the absolute values for the vector genomes and the mouse reference gene, calculate vg/dg in the sample using equations (7-8).
    For reference genes with two gene copies/diploid genome:
    Absolute value of diploid genomes (dg) = (7)
    vg/dg = (8)
  4. Check the 1D scatterplot of the dPCR reaction to confirm the validity of the assay and quantification (Figure 3A,C). For the assay to be valid, confirm that the 1D scatterplot meets all the following criteria: the presence of positive and negative partitions; clear separation between the positive and negative partitions to allow accurate threshold determination; and the presence of no-to-few droplets between the positive and negative partitions (also known as rain), which can reduce the accuracy of the dPCR quantification.

Representative Results

Seven to nine weeks-old male mice were injected with AAV via lateral tail-vein injection at 1.5 × 10¹² vg/mouse delivered in 150-200 µL of injectate volume. The ssDNA AAV used here delivered Cre recombinase transgene driven by CMV promoter. The injected mice were homozygous for the Cre reporter Ai14 allele. When exposed to Cre recombinase, Ai14 allele-containing cells express the fluorescent tdTomato protein. Since tdTomato expression is caused by Cre-induced genomic recombination, tdTomato-expressing cells indicate cells that were either directly transduced by the AAV or were progeny cells of transduced cells. The data shown here are of mice injected with AAV9-CMV-Cre at 1.5 × 10¹² vg/mouse delivered in 160 µL (5.8-5.9 × 10¹³ vg/kg). The mice were sacrificed 28 days post injection, and the tissues were collected as described above. A few skeletal muscles and liver lobes were digested, and their cells were collected using FACS. A few liver lobes were frozen immediately using prechilled methylbutane for nucleic acid extraction. A few skeletal muscles and liver lobes were fixed-frozen for histological imaging of fluorescent tdTomato. tdTomato was expressed diffusely throughout the liver (Figure 2A) and quadriceps (Figure 2C) indicating that AAV9 broadly reached and transduced different regions of both tissues.

DNA extracted from fresh-frozen liver and FACS-sorted cells was used to quantify vg/dg using dPCR. Vg/dg quantification can be used to assess injection consistency and the transduction efficiency of AAV in the analyzed sample. The 1D droplet scatterplots from the fresh-frozen liver tissue sample and FACS-sorted cells were used to ensure the validity of the assay (Figure 3A,C). The scatterplot showed the presence of positive and negative partitions, clear separation between the positive and negative partitions that allows accurate determination of the detection threshold, and the presence of no-to-few droplets between the positive and negative partitions, which can reduce the accuracy of the dPCR assay. Meeting all these criteria indicated that the dPCR assay results were valid. The number of Polr2a gene copies in each sample was quantified to determine the number of mouse diploid genomes (2 Polr2a gene copies/mouse diploid genome), and primers/probe against the Cre recombinase transgene sequence were used to quantify the viral genome (1 transgene copy/viral genome, Table 1). The vg/dg value was quantified for the fresh-frozen liver tissue sample and FACS-sorted cells and showed the presence of 187.7 vg/dg and 4.7 vg/dg in each sample, respectively (Figure 3B,D). Samples from PBS-injected mice and non-template controls containing no nucleic acids were used as negative controls.

Figure 1: Intravenous injection station overview. (A) Tools needed to perform IV injection. Shown here is the (1) timer, (2) mouse tube restrainer, (3) uncut and (4) cut plastic restrainer cones, (5) alcohol swab, (6) empty pipette tips box used as a platform to elevate the mouse tube restrainer, (7) disposable absorbent pads, (8) 15 mL conical tube with warm water, (9) 15 mL tube holder, (10) gauze, and (11) insulin syringe. (B) The mouse is first placed inside the tube restrainer. Then, the cut restrainer cone is inserted to create a restraining sleeve around the mouse, if the mouse is too small to be restrained by the tube restrainer only. Ensure that the mouse's breathing is not obstructed by the restrainers. The tube restrainer is placed on top of the elevated platform to allow the placement of the mouse tail in warm water. (C) Mouse tail positioning and needle holding angle immediately before performing the injection. Pull back on the tail so the tail is stretched, and the injection site is completely horizontal. The needle is parallel to the tail and the vein, and the bevel is facing upwards. Please click here to view a larger version of this figure.

Figure 2: Detection of fluorescent reporter protein post IV injection. Seven to nine weeks-old male mice harboring the Cre reporter Ai14 allele were IV injected with either AAV9-CMV-Cre at 1.5 × 10¹² vg/mouse delivered in 160 µL (5.8-5.9 × 10¹³ vg/kg) or PBS. Representative fluorescence images of mouse (A) liver or (C) quadriceps sections post AAV9 delivering Cre IV injection. (B) Liver or (D) quadriceps sections from PBS-injected mice were imaged to serve as negative controls. The tissues were collected and fixed-frozen 28 days post IV injection. Post-Cre exposure, fluorescent tdTomato protein is expressed in transduced cells and progeny cells of transduced cells. 10 µm-thick sections were imaged at 10x magnification. Scale bars = 100 µm. Please click here to view a larger version of this figure.

Figure 3: Vector genome per diploid genome (vg/dg) quantification. 1D scatterplot of dPCR vector genomes quantification in (A) liver tissue or (C) FACS-sorted cells collected from mice injected with either AAV9-CMV-Cre or PBS. The scatterplots show the positive and negative dPCR partitions, as well as the detection threshold indicated by the horizontal line across the samples. (B,D) vg/dg quantification after quantifying the mouse diploid genomes and vector genomes in the (B) liver tissue or (D) FACS-sorted cell samples. Results shown here are from a single AAV9-injected mouse and a single PBS-injected mouse with a technical dPCR duplicate for each mouse. Error bars indicate the 95% confidence interval for each sample. Abbreviations: NTC= Non-template control; dPCR = digital PCR; FACS = fluorescence-activated cell sorting. Please click here to view a larger version of this figure.

Primer	Sequence
Cre forward primer	CTGACGGTGGGAGAATGTTAAT
Cre reverse primer	CATCGCTCGACCAGTTTAGTT
Cre probe	/56-FAM/CGCAGGTGT/ZEN/AGAGAAGGCACTTAGC/3IABkFQ/
Polr2a forward primer	GACTCCTTCACTCACTGTCTTC
Polr2a reverse primer	TCTTGCTAGGCAGTCCATTATC
Polr2a probe	/5HEX/ACGAGATGC/ZEN/TGAAAGAGCCAAGGT/3IABkFQ/

Table 1: Sequences of primers and probes used for vg/dg quantification. Cre primers and probe were used to quantify the vector genome. Polr2a primers and probe were used to quantify the mouse diploid genome.

Discussion

AAV-based therapies hold great potential for monogenic disorders due to the versatility of AAV as a gene therapy vector, which makes it possible to customize AAVs to meet the various delivery needs of different disorders⁴^,⁵^,⁷^,⁹. AAVs are commonly administered via IV injection in preclinical mouse models to test the safety and efficacy of potential therapeutics¹⁶. As different injected AAV doses can result in marked differences in the experimental outcomes, it is critical for experimenters to be able to consistently inject the intended AAV dose to ensure the validity and robustness of the generated in vivo data²⁸. IV injections are widely used, but they are technically challenging requiring extensive training and continuous practice to develop and maintain a skill level that ensures consistently successful injections¹⁶^,¹⁷^,¹⁸^,¹⁹. In addition to correctly injecting AAV, it is usually desired to use assays to assess the injected AAV's biodistribution and delivery efficiency to the target tissues or cells²⁹^,³⁰.

This protocol aims to assist experimenters to easily perform IV injections successfully and consistently by thoroughly describing the details of an optimized IV injection protocol to administer AAV in 7-9 week-old, unsedated mice. It is important to note that mice that are markedly smaller or larger than wild type mice in the age range used here may present a greater challenge due to a reduced visibility of the veins or incompatibility with the restrainers used in this method. It has been previously reported that tail IV injections are not appropriate for administering reagents intravenously in mice younger than 6 weeks old due to the small vessel size³¹. Although possible, it might be difficult to consistently inject mice weighing less than 22.0 g. successfully. Investigators using mice of atypical size may need to make adaptations to the procedure. This protocol also outlines several assays that can be used to assess AAV biodistribution and transduction efficiency.

Some critical points need to be kept in mind while following this protocol. During injection, 29 G needles provide greater resistance if the needle is not inside the vein. This reduces the volume lost from accidental perivascular injection of the solution during failed injection attempts. Insulin syringes have smaller dead volumes than regular syringes. If using a different syringe and/or needle than the ones listed here, additional injectate volume might need to be prepared in protocol steps 1.1.3.3 to account for larger dead space volume (e.g., add 30 µL to the intended dose instead of 15 µL).

If fine aspiration-caused air bubbles are formed on the syringe sides while aspirating the AAV dose into the syringe, slowly pull the injectate further up the syringe. This will remove most small air bubbles. Load at least an additional 10-15 µL of AAV to the intended volume to be injected. This additional volume is to account for any volume that might be lost during the expelling of air bubbles or potential failed injection attempts. (e.g., if the target volume to be injected is 150 µL, load 165 µL into the syringe (halfway between the 160 µL and 170 µL marks on the syringe scale). If the needle is correctly placed inside the vein, and the volume in the syringe is at 165 µL immediately before the successful injection attempt, deliver the reagent until 15 µL are left in the syringe (halfway between the 10 µL and 20 µL marks), thus delivering 150 µL (165 µL – 150 µL= 15 µL)). Aligning the bevel lumen (bevel facing upwards) with the syringe scale allows tracking the delivered volume during the injection.

Some experimenters might prefer to place the mouse on its side so that one of its veins is straight and easily accessible compared to a mouse on its feet. However, the tail of a mouse on its side will be slanted at different angles depending on the mouse size requiring injection-angle adjustment when injecting mice of different sizes. This might negatively impact the consistency of success of the procedure. During initial practice attempts, experimenters can try both mouse restraining orientations to determine their preferred approach. Having the mouse on its feet allows quick and easy access to both lateral tail veins. This reduces the restraining time when access to both veins is needed in the case of multiple failed injection attempts.

If injecting the lateral vein close to the tail base (closer to the mouse body) (especially for mice weighing >30 g.), adjust the injection angle from parallel to the vein to 5°-10° to the vein since the vein at the tail base is slightly deeper than it is distally.

The RNase digestion and RNA contamination check protocols listed here were verified on DNA samples isolated from fresh-frozen liver tissues containing a total of 175-700 ng of nucleic acids in 20 µL. The RNase digestion protocol was also tested on DNA samples isolated from fresh-frozen liver tissues and FACS-sorted cells to confirm the presence of the vector genome and the mouse genome after RNase digestion. The results were visualized using agarose gel electrophoresis of endpoint PCR amplification of the target amplicons.

Following the described methodology can reduce the training and practice time needed to master IV injections and result in a higher successful injection rate, which would save reagents. This protocol utilizes simple and commonly used tools without the need for advanced equipment or setups that might not be readily available. Furthermore, the IV injection steps listed here can be applied to a wide range of injectates that need to be administered intravenously, such as antisense oligonucleotides (ASOs), with the appropriate modifications made to the injectate preparation steps depending on the injectate.

Offenlegungen

The authors have nothing to disclose.

Acknowledgements

The authors would like to thank the NINDS animal care facility staff for their support. This work was supported by the Division of Intramural Research of the NIH, NINDS (Annual Report Number 1ZIANS003129). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

Materials

0.22 µm syringe filter	Millipore	SLGVM33RS
0.3 mL insulin syringes with 29G needle	BD Biosciences	324702
1.7 mL microcentrifuge tube	Crystalgen	23-2051
10 mL syringe	BD Biosciences	302995
100% EtOH	The Warner Graham Company	201096
10x phosphate-buffered saline (PBS)	Corning	46-013-CM	Used to prepare 1x PBS for tissue fixation
15 mL conical tube	Corning	430766
15 mL conical tube holder	Multiple sources	N/A
190 proof ethyl alcohol	The Warner Graham Company	6810-01-113-7320	Used to prepare 70% ethanol
1x sterile PBS	Gibco	10010023
2 mL microcentrifuge tissue storage tubes	Eppendorf	022363344
4% paraformaldehyde (PFA)	Electron Microscopy Sciences	157-4
Adeno-associated virus (AAV)	Charles River	N/A	Single-stranded DNA (ssDNA) AAV was packaged to deliver Cre recombinase as the transgene driven by CMV promoter
Alcohol swab	BD Biosciences	326895
Bead lysis tube	Next Advance	GREENE5
BsuRI (HaeIII) restriction enzyme	Thermo Fisher Scientific	ER0151
Bullet blender	Next Advance	BBX24B
Ai14-derived mice from JAX 007914 strain (genetic background: C57BL/6J)	N/A	N/A	Mice containing Ai14 Cre-reporter allele were purchased from JAX (catalog number: 007914)
Disposable absorbent pads	Fisherbrand	1420662
Dissection forceps	Fine Science Tools (F.S.T)	11251-35
Dissection scissors	Fine Science Tools (F.S.T)	14085-08
DNA degradation reagent (DNAZap)	Invitrogen	AM9890
DNA-Extraction RNase A	Qiagen	19101	For RNA digestion during nucleic acid extraction
DNase-free RNase for DNA cleanup	F. Hoffmann-La Roche	11119915001	For RNA digestion after nucleic acid extraction
dPCR Probe PCR Kit	Qiagen	250102
dPCR software	Qiagen	N/A	QIAcuity Software Suite
Elevated platform	Multiple sources	N/A	An empty pipette tips box was used to elevate the mouse restrainer during tail warming up
Fluorescence microscope	Multiple sources	N/A	Model used here: Nikon Eclipse Ti
Fluorescence microscope software	Multiple sources	N/A	Software used here: NIS-Elements
Gauze	Covidien	9022
Heat block	Eppendorf	Thermomixer 5350
High-speed centrifuge	Eppendorf	22620689
Metal container	Vollrath	80125
Methylbutane	J.T. Baker	Q223-08
Molecular grade water	Quality Biological	351-029-131
Mouse tube restrainer	Braintree Scientific	TV-RED-150-STD
Myfuge mini centrifuge	Benchmark Scientific	C1012
Polymerase chain reaction thermal cycler	Bio-Rad Laboratories	1851148	Model: C1000 Touch
Precision wipes	Kimberly-Clark Professional	7552
Proteinase K	Qiagen	19131
QIAcuity dPCR Nanoplate 26k 24-well	Qiagen	250001
QIAcuity One dPCR system	Qiagen	911020
Qiagen DNeasy Blood & Tissue Kit	Qiagen	69504	Used for DNA extraction from tissues
Qiagen QIAamp DNA Micro Kit	Qiagen	56304	Used for cleanup of genomic DNA, and the isolation of DNA from small volumes of blood prtocotol was used for DNA extraction from FACS-sorted cells
Rodent restrainer cone	Braintree Scientific	MDC-200
Scale	Ohaus	72212663
Styrofoam box	Multiple sources	N/A
Sucrose	Sigma-Aldrich	S9378-1kg
Surface cleaner and disinfectant	Peroxigard	29101
Timer	Multiple sources	N/A
Transfer forceps	Fine Science Tools (F.S.T)	91113-10
Vortex	Daigger & Company	22220A	Model: Daigger Vortex Genie 2

Referenzen

Tisdale, A., et al. The ideas initiative: Pilot study to assess the impact of rare diseases on patients and healthcare systems. Orphanet J Rare Dis. 16 (1), 429 (2021).
Condo, I. Rare monogenic diseases: Molecular pathophysiology and novel therapies. Int J Mol Sci. 23 (12), 6525 (2022).
Vaisitti, T., et al. The frequency of rare and monogenic diseases in pediatric organ transplant recipients in italy. Orphanet J Rare Dis. 16 (1), 374 (2021).
Bulcha, J. T., Wang, Y., Ma, H., Tai, P. W. L., Gao, G. Viral vector platforms within the gene therapy landscape. Signal Transduct Target Ther. 6 (1), 53 (2021).
Lundstrom, K. Viral vectors in gene therapy: Where do we stand in 2023. Viruses. 15 (3), 698 (2023).
Wang, C., et al. Emerging non-viral vectors for gene delivery. J Nanobiotechnology. 21 (1), 272 (2023).
Naso, M. F., Tomkowicz, B., Perry, W. L., Strohl, W. R. Adeno-associated virus (aav) as a vector for gene therapy. BioDrugs. 31 (4), 317-334 (2017).
Ghosh, S., Brown, A. M., Jenkins, C., Campbell, K. Viral vector systems for gene therapy: A comprehensive literature review of progress and biosafety challenges. Appl Biosaf. 25 (1), 7-18 (2020).
Wang, D., Tai, P. W. L., Gao, G. Adeno-associated virus vector as a platform for gene therapy delivery. Nat Rev Drug Discov. 18 (5), 358-378 (2019).
Srivastava, A. Rationale and strategies for the development of safe and effective optimized aav vectors for human gene therapy. Mol Ther Nucleic Acids. 32, 949-959 (2023).
Mukherjee, P., Roy, S., Ghosh, D., Nandi, S. K. Role of animal models in biomedical research: A review. Lab Anim Res. 38 (1), 18 (2022).
Jucker, M. The benefits and limitations of animal models for translational research in neurodegenerative diseases. Nat Med. 16 (11), 1210-1214 (2010).
Vandamme, T. F. Use of rodents as models of human diseases. J Pharm Bioallied Sci. 6 (1), 2-9 (2014).
Anders, H. J., Vielhauer, V. Identifying and validating novel targets with in vivo disease models: Guidelines for study design. Drug Discov Today. 12 (11-12), 446-451 (2007).
Turner, P. V., Brabb, T., Pekow, C., Vasbinder, M. A. Administration of substances to laboratory animals: Routes of administration and factors to consider. J Am Assoc Lab Anim Sci. 50 (5), 600-613 (2011).
Prabhakar, S., Lule, S., Da Hora, C. C., Breakefield, X. O., Cheah, P. S. Aav9 transduction mediated by systemic delivery of vector via retro-orbital injection in newborn, neonatal and juvenile mice. Exp Anim. 70 (4), 450-458 (2021).
Hauff, P., Nebendahl, K., Kiessling, F., Pichler, B. J., Hauff, P. Drug administration. Small Animal Imaging: Basics and Practical. , 127-152 (2017).
Saleem, M., et al. A new best practice for validating tail vein injections in rat with near-infrared-labeled agents. J Vis Exp. (146), (2019).
Resch, M., Neels, T., Tichy, A., Palme, R., Rulicke, T. Impact assessment of tail-vein injection in mice using a modified anaesthesia induction chamber versus a common restrainer without anaesthesia. Lab Anim. 53 (2), 190-201 (2019).
Glascock, J. J., et al. Delivery of therapeutic agents through intracerebroventricular (icv) and intravenous (iv) injection in mice. J Vis Exp. (56), 2968 (2011).
Grames, M. S., Jackson, K. L., Dayton, R. D., Stanford, J. A., Klein, R. L. Methods and tips for intravenous administration of adeno-associated virus to rats and evaluation of central nervous system transduction. J Vis Exp. (126), 55994 (2017).
Yano, J., Lilly, E. A., Noverr, M. C., Fidel, P. L. A contemporary warming/restraining device for efficient tail vein injections in a murine fungal sepsis model. J Vis Exp. (165), 61961 (2020).
Warren, J. S. A., Feustel, P. J., Lamar, J. M. Combined use of tail vein metastasis assays and real-time in vivo imaging to quantify breast cancer metastatic colonization and burden in the lungs. J Vis Exp. (154), 60687 (2019).
JoVE Science Education Database. . Lab Animal Research. Compound administration I. , (2023).
UCSF Office of Research, Institutional Animal Care and Use Program. Lateral tail vein injection in mice and rats (preferred technique for vascular access in mice Available from: https://iacuc.ucsf.edu/sites/g/files/tkssra751/f/wysiwyg/STD%20PROCEDURE%20-%20Misc%20Rodent%20Procedures%20-%20Lateral%20Tail%20Vein%20Injection%20in%20Mice%20and%20Rats.pdf (2023)
Jones, K. Tail vein injections in the mouse and rat sop. UBC Animal Care Guidelines SOP: ACC-2012-Tech03. , (2012).
Liadaki, K., Luth, E. S., Kunkell, L. M. Co-detection of gfp and dystrophin in skeletal muscle tissue sections. Biotechniques. 42 (6), 699-700 (2007).
Burr, A., et al. Allometric-like scaling of aav gene therapy for systemic protein delivery. Mol Ther Methods Clin Dev. 27, 368-379 (2022).
Lang, J. F., Toulmin, S. A., Brida, K. L., Eisenlohr, L. C., Davidson, B. L. Standard screening methods underreport aav-mediated transduction and gene editing. Nat Commun. 10 (1), 3415 (2019).
Rodriguez-Estevez, L., Asokan, P., Borras, T. Transduction optimization of aav vectors for human gene therapy of glaucoma and their reversed cell entry characteristics. Gene Ther. 27 (3-4), 127-142 (2020).
Hatakeyama, S., Yamamoto, H., Ohyama, C. Tumor formation assays. Methods Enzymol. 479, 397-411 (2010).