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Adeno-associated virus (AAV) in a small replication deficient DNA virus belonging to the Parvovirinae family. It has a single-stranded approximately 4.7kb genome. Recombinant AAV (rAAV) is created by replacing the viral rep and cap genes with the transgene of interest along with promoter and polyadenylation sequences. The short viral inverted terminal repeats must remain intact for replication and packaging in production as well as vector genome processing and persistence in the transduction process. The AAV capsid (serotype) determines the tissue tropism of the rAAV vector. In this unit we will discuss serotype selection for lung targeting along with the factors effecting efficient delivery of rAAV vectors to the murine lung. Detailed procedures for lung delivery (intranasal, orotracheal, and surgical tracheal injection), sample collection and post-mortem tissue processing will be described.
Adeno-associated virus (AAV) in a small, (20um) replication deficient DNA virus belonging to the Dependovirus genus. It is a member of the Parvovirinae family with a single-stranded approximately 4.7kb genome. The wild-type AAV genome contains rep and cap genes flanked by short inverted terminal repeat (ITR) sequences. While AAV infections are common in humans, no clinical disease has been associated with AAV. Adeno-associated viruses have large capsid sequence variability with 6 clades of AAVs identified thus far (Gao, 2004). It is the AAV capsid (serotype) that determines the tissue specificity upon natural or experimental infection (Grimm, 2006). Recombinant AAV (rAAV), for use in gene transfer vectors, is created by replacing the rep and cap genes with the transgene of interest along with promoter and polyadenylation sequences. The ITRs must remain intact for replication and packaging in production as well as vector genome processing and persistence in the transduction process. The rep and cap genes are provided in a complementing plasmid, along with adenovirus or herpes simplex virus helper gene products, to successfully package the rAAVs. rAAV persists in non-dividing cells as an extra-chromosomal element (Flotte, 1994 and Afione, 1996).
The lung was the target of the first human rAAV trial (Flotte, 1996). In that trial a rAAV2 vector was used to deliver the CFTR gene to the lung of cystic fibrosis patients. rAAV has been used in multiple human trials subsequently, targeting the lung as well as other organ systems. While the safety of rAAV in human trials has been established, other challenges to efficient long-term gene transfer were encountered (Mueller, 2008 and High, 2011). We will discuss this further in the background section along with suggestions on serotype selection, expression analysis and study design.
This unit will cover methods of delivering rAAV to the murine lung (intranasal, orotracheal intubation and surgical tracheal injection) as well as evaluation of gene transfer (serum collection, bronchoalveolar lavage technique, and post-mortem collection and processing of lung tissue).
CAUTION: According to the NIH Guidelines for Research Involving Recombinant DNA Molecules (April 2000), adeno-associated virus is a Biosafety Level 1 (BSL-1) pathogen because both rAAV and wild-type AAVs are not known to cause disease in healthy human adults. BSL-1 status assumes that the rAAV construct does not encode a toxic or tumorigenic molecule and in produced without a helper virus. Biosafety protocols should be approved by the Institutional Biosafety Committee at the institution where the research is being conducted.
CAUTION: This experiment requires Animal Biosafety Level 1 (ABSL-1) conditions. Biosafety protocols should be approved by the Institutional Biosafety Committee at the institution where the research is being conducted. Protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) or must conform to governmental regulations regarding the care and use of laboratory animals.
The intranasal route provides a straight-forward method of vector delivery with an affordable equipment list and a small learning curve. This leads to easily repeatable results in even the novice murine researcher. The downfall of nasal delivery is loss of vector in the upper airways (nasal passages, sinuses, and pharynx) and gastrointestinal tract rather than having the entire vector dose deposited within the lower airways. This may necessitate larger vector doses to attain the same level of transduction compared to intra-tracheal delivery.
Ophthalmic ointment (such as Puralube, Webster Item #: 07-888-2572)
3/10 cc syringe with 31 gauge needle (8mm length needle) (such as: BD Ultra-Fine II Short Needle Insulin Syringes)
100 or 200 microliter micro-pipette tip
Orotracheal intubation provides the ability to directly instill vector into the lung without the risks of a surgical procedure or loss of vector in the upper airways. This means that the dose of vector reaching the lungs is more tightly controlled than with intranasal instillation because essentially the entire amount delivered reaches the lung. Orotracheal intubation does have an increased equipment investment and a steeper technique learning curve compare to intranasal instillation.
Additional Materials (also see Basic Protocol 1)
20 gauge 1.25 inch catheter (BD Angiocath, Becton Dickinson, Sandy Utah)
2% Lidocaine HCl Jelly (30 ml) (Webster Item #: 07-835-7610)
1 ml tuberculin slip tip syringe
Surgical injection of vector into the trachea has the advantage of improved lung vector delivery compared with intranasal instillation and decreased equipment costs and ease of learning compared with orotracheal intubation. There is increased risk associated with a surgical procedure including hemorrhage, incision dehiscence, infection, and longer recovery. Because of the small size of the murine trachea there is also a risk of injecting the vector through the trachea and targeting the peritracheal tissues or esophagus rather than the lung.
Materials (also see Basic Protocol 1)
#15 scalpel blade and handle or 4 ½ inch curved tissue scissors
(2) Adson tissue forceps (or other delicate tissue forcep)
4-0 monofilament absorbable suture on a cutting needle(such as PDSII) or tissue glue (such as Dermabond)
Needle holder (for suturing skin)
Depilatory cream (such as Nair), scissors, or clippers to remove hair from surgical sight
Chlorhexadine or betadine scrub and 70% alcohol
IV butterfly catheter with 23 or 25 gauge needle (alternative option)
This protocol offers an alternative approach to ortracheal intubation to that described in Alternate Protocol 1. The equipment costs are less than those for speculum aided intubation (unless your laboratory already owns an otoscope) and the system boasts a smaller learning curve. The disadvantages include the lack of a magnifying glass associated with the system (this is present in the otoscope) which may increase the difficulty level in visualizing the larynx appropriately, especially in smaller mice, as well as the inability to move soft tissue out of the way (such as the soft palate) using the speculum blade in order to better visualize the larynx. The system consists of a fiber optic illuminator and fiber optic stylet rather than an otoscope with speculum and separate stylet used for Alternate Protocol 1. The fiber optic stylet allows simultaneous visualization and cannulation of the larynx, without magnification. The intubation catheter used is identical to that used in Alternate Protocol 1. See Alternate Protocol 1 for specific pros and cons of orotracheal intubation as a delivery route for viral vectors. See Internet Resources section for a video demonstrating the BioLITE procedure.
Additional Materials (also see Alternate Protocol 1)
Preparing rAAV Vector for Delivery
The following are the steps we follow to prepare rAAV vectors for delivery to the mouse.
Pipette tips (200ul)
Parafilm or sterile petri dish
Delivery syringe or pipette (see particular delivery protocol)
Bronchoalveolar lavage collection allows for quantification of rAAV gene products within the distal airways. It is a relatively easy procedure to perform and requires minimal equipment.
Additional Materials (also see Alternate Protocol 2)
3 ml syringe
1.5 ml eppendorf tube
Proper removal of blood from the lungs will decrease autofloresence when performing immunofluorescent staining. Fixing the lungs while inflated will aid in histologic evaluation of the lung because the alveoli will have a more normal architecture rather appearing collapse, allowing identification of individual epithelial cells.
Additional Materials (also see Alternate Protocol 2 and Support Protocol 2)
5 ml syringe
27 gauge needle
10 ml conical tube
Neutral buffered formalin
|Xylazine (20mg/ml stock concentration)||0.25 ml (10 mg/kg dose to mouse)|
|Ketamine (100mg/ml stock concentration):||0.5 ml (100mg/kg dose to mouse)|
|Sterile Isotonic Saline:||5 ml|
|Total Cocktail:||5.75 ml|
Dose of cocktail to mouse: 0.10 ml/10 g, intraperitoneally
Weigh all animals to determine anesthetic dose. Place the animal in a quiet cage following anesthetic administration and allow at least 5 minutes to pass before checking anesthetic depth by toe pinch. If adequate anesthetic depth is not present allow the animal another 5 minutes in the quiet cage. If still not sufficiently anesthetized a 50 mg/kg dose of ketamine alone should be administered. If that does not result in sufficient anesthesia then a 0.05 ml/10 g dose of the xylazine/ketamine cocktail can be re-administered.
The lung was the first site for rAAV gene therapy in humans. The initial gene therapy trials involving the lung have shown rAAV to be a safe vector for human gene therapy, with only transient moderate adverse events being reported in any of the clinical trials published thus far. However, efficient gene therapy in the lung has encountered other obstacles. The original cystic fibrosis trial used rAAV2 to deliver the CFTR gene, with only low levels of expression seen within the lung. It was discovered that rAAV2 requires heparin sulfate proteoglycans (HSP) for efficient cell entry (Summerford, 1998). HSPs are only located on the basolateral surface of airway epithelial cells, not the apical surface. This necessitates disruption of the tight junctions between airway epithelial cells in order to allow the rAAV2 viral vector to gain access to the HSP receptors. Subsequently other serotypes have shown improved lung transduction, without the necessity for tight junction disruption, due to the presence of apical surface receptors. Currently the most promising vectors for lung-targeted gene therapy include AAV5, AAV6, AAV9 and AAV6.2. The choice of serotype should be based on the cell type that is being targeted (see Table 1). It should be noted however, that Table 1 focuses on transduction efficiency in murine models, it has been shown that murine and lower primate models may not reflect the transduction efficiency in higher primates, specifically chimpanzees and humans (Liu, 2007 and Flotte, 2010). Specifically, AAV5 showed higher efficiency in mouse and lower primate models, whereas AAV1 had improved efficiency in human airway epithelials cells and chimpanzees airways (Lui, 2007 and Flotte, 2010).
An additional problem encountered in rAAV mediated gene transfer to the lung is the lack of transgene persistence in the dividing epithelial cell population. This is due to the fact that, as mentioned above, rAAV is present in transduced cells as an extra-chromosomal element (Afione, 1996). This lack of persistence is compounded by the fact that repeated dosing of rAAV leads to inefficient transduction, depending on serotype, as a result of neutralizing antibodies directed against the AAV capsid (Sinn, 2009). Transgene expression in the murine lung following a single vector administration has been detected up to 217 days after dosing, with a decrease in transgene expression occurring between day 49 and 77 and vector genomes between day 28 and 90 (Limberis, 2006).
Neutralizing antibodies directed against AAV capsid are the main immune impediment to successful rAAV transduction, although T cell and innate immune responses, such as alveolar macrophages and certain pattern recognition receptors also pose a potential challenge (Zaiss, 2008 and High, 2011). Preexisting immunity to AAV is of particular concern when selecting a serotype for experimentation with future consideration of translation to human subjects. The prevalence of preexisting antibodies in humans is largely serotype dependent, although significant cross-reactivity exists between some serotypes (Boutin, 2010). Neutralizing antibiodies in humans are most prevalent against AAV serotypes 1 and 2, with lower prevalence and titers for 5, 8, and 9 (Boutin, 2010). In cystic fibrosis patients, AAV2 neutralizing antibody titers were slightly higher than titers against AAV5 and AAV6 and adults demonstrated significantly higher titers than children (Halbert, 2006). It is also possible to have an immune response directed toward the delivered transgene (Moore, 2010 and Zaiss, 2008). This has further reaching implications for patient trials as it could preclude other treatments such as protein replacement therapy (Zaiss, 2008). Neutralizing antibody titers have also been shown to increase in patients following rAAV2 vector delivery to the lung, especially evident following repeat dosing (Moss, 2004 and 2007).
Physical barriers to efficient vector delivery also exist in the lung, including loss in the upper and conducting airways and gastrointestinal tract following intranasal delivery (Southam, 2002). Barriers within the lung include the mucociliary clearance system, the luminal epithelial glycocalyx, and as mentioned above, tight junction between airway epithelial cells (Kolb, 2006). There are also disease specific impediments to vector lung access, such as the viscous mucus layer that coats the respiratory epithelium in cases of cystic fibrosis (Hida, 2011).
All these factors must be taken into account when planning gene transfer experiments in the lung. When choosing a reporter or therapeutic rAAV gene construct a plan for determining transgene expression efficiency must be carefully designed. Some important factors to consider are whether the transgene is an intracellular or secreted protein. Are there diagnostic tests readily available to detect the transgene product such as ELISA, immunohistochemistry, Westernblot? It has been established that a significant amount of vector reaches the liver following respiratory tract delivery (Limberis, 2006, Mueller, manuscript in preparation). If the transgene product is secreted, is there a way to determine whether production is occurring due to lung transduction or transduction of another organ such as the liver? Immunohistochemistry or Westernblot of the lung to detect gene product within lung cells or detection of gene product within bronchoalveolar lavage fluid can aid in proving lung origin gene product. It is also important to determine whether gene product expression levels are due to serotype transduction efficiency versus gene or promoter efficiency. Determining vector genomes within the lung tissue by quantitative real-time PCR can help parse out this difference. For example, if vector genomes present are high, but gene product levels are low, the low level of gene product could be attributable to multiple factors that affect transgene expression, other than poor vector distribution, such as abnormalities in viral intracellular trafficking, nuclear targeting, uncoating, and DNA processing (Sanlioglu, 2001 and Ding, 2005).
In conclusion, several rAAV vectors have been shown to target the lung, but serotype specific, immune, and physical barriers to transduction should be considered when planning a lung gene transfer experiment. While a detailed description of transgene product detection procedures is beyond the scope of this unit, we hope that the above commentary will aid you in selecting an appropriate detection method.
The main pitfall encountered with intranasal delivery will involve poor distribution to the lung. Ensuring that vector is not lost by allowing adequate time for inhalation while delivering the vector solution is critical. Poorly anesthetized animals are also more likely to swallow the vector rather than inhaling it into the lungs (Southam, 2002). Poor lung deposition will also result if the volume delivered is less than 50ul total (Southam, 2002). If needle and syringe are used to deposit the vector, poor lung delivery could result if the vector is inadvertently injected into the nasal epithelium rather than into the nasal passage.
Inadvertent catheter placement into the esophagus is the most common mistake that occurs during the orotracheal intubation process. Quickly identifying that the catheter is in the wrong location will prevent inadvertent esophageal delivery of the vector. In order to avoid catheterizing the esophagus it is imperative to see the stylet entering the laryngeal opening. It may be necessary to bend or extend the stylet slightly in order to allow better visualization of entry. Repositioning the animal so that the head and neck are straight and in-line will also aid in adequate visualization of the larynx. Adequate anesthesia will also decrease swallowing and provide easier access to the larynx.
If the catheter is properly positioned within the trachea, not in the esophagus, it is normal to notice a change in breathing pattern when delivering the vector and subsequent air.
As mentioned in the protocol, death of the mouse can occur secondary to laryngeal trauma if more than 2 intubation attempts are made in one animal. If it is critical to dose that animal, it is advisable to allow 24 hours for recovery if any reddening or swelling is visualized.
Potential problems associated with surgical vector delivery include trauma to critical structures in the neck including, carotid arteries, jugular veins, and esophagus. The carotid arteries and jugular veins are located along side the trachea and the esophagus is deep to it. Careful dissection down to the trachea should prevent inadvertent damage to any of these structures. Adherence to aseptic technique during the surgical procedure should prevent any infections. This involves proper hair removal, disinfection of the skin, and sterile surgical instruments. Some animal care and use committees may also recommend the use of a sterile drape, sterile surgical gloves, surgical mask, and preoperative antibiotics to prevent infection. Another common complication in any murine surgery is the animals removing the skin sutures. If this occurs it may be necessary to reanesthetize the animal and replace the sutures. If several days have passed since the surgery enough healing may have occurred such that the sutures will not need to be replaced. Some surgeons prefer tissue glue to prevent issue of suture removal, however some incision inflammation has been described with this product.
Problems of vector delivery could also occur. Ensuring that the needle is in the trachea can be done by drawing back on the syringe prior to and after delivery of vector to be sure that air is obtained. The animals should also be adequately anesthetized to ensure that the vector is inhaled and not coughed up and swallowed. Directing the needle toward the lung and having the animal positioned at an angle with the head up should also increase lung deposition.
See speculum/otoscope section for issues related to adequate visualization of the larynx.
Suturing the catheter in place will prevent inadvertent catheter removal during delivery and removal of the fluid. Do not insert the catheter too far into the tracheal (no more than 2–3mm) or it can push through the distal trachea into the mediastinum resulting in no return of fluid.
A poor flush of the lungs can results from inadvertently flushing through the left ventricle, pushing the needle too far into the heart, or delivering the PBS solution too slowly.
Poor inflation of the lungs with fixative will occur if the catheter is advanced too far and punctures the trachea, if the lungs are cut upon opening of the chest cavity, or the trachea is not closed well following catheter removal. Do not insert the catheter too far into the tracheal (no more than 2–3mm) to prevent tracheal puncture. Making a small nick in diaphragm before cutting diaphragm or chest wall further will allow the lungs to collapse and decrease chance of cutting the pulmonary tissue. Wait to inflate the lungs until after the thoracic cavity has been opened to prevent lung lacerations. Using blunt tipped scissors to open the thorax can also decrease the chance of puncturing the lungs.
If inflation and formalin fixation is not providing enough tissue fixation, such that the lungs still collapse when being sectioned, then 2.5% glutaraldehyde can be used in place of formalin for the procedure described above. If the lung tissue is going to be frozen in OCT, the lungs can be inflated with OCT instead of formalin.
All of the lung delivery techniques described above should result in widespread lung transduction, depending on serotype and promoter used. The duration to maximum expression of the delivered transgene will depend on multiple factors, including serotype, promoter, and whether vector is single stranded or self-complementary. Expression should be expected within 7 days with peak expression likely between 3–6 weeks.
This procedure can be quickly learned and dosing an animal should take less than 1 hour, with 5–10 minutes for anesthesia, 5 minutes or less for dosing, and 20–30 minutes for anesthesia recovery. Multiple animals can be anesthetized and recovered simultaneously, meaning that the largely limiting factor for number of animals dosed per hour will be actual vector delivery time.
Once this technique is learned dosing a single animal takes approximately 5–10 minutes for catheter placement and vector instillation. Times for anesthesia and recovery are similar to intranasal delivery. This technique can take several attempts, meaning multiple animals, before the catheter can be placed in the trachea successfully in less than 2 attempts. It is advised to learn the procedure on non-experimental animals.
Anesthesia and recovery times will be similar to intranasal delivery. Surgery and delivery time will depend on the operator’s surgical experience. Inexperienced investigators may take as long as 20 minutes to complete the technique from incision to closure, while experience investigators may be able to complete the entire technique in around 5 minutes. For longer surgical times it may be necessary to redoes the anesthetic which will prolong the recovery time.
Detailed procedural video demonstrating the orotracheal intubation technique. Note minor modification from technique described above. We do recommend the 20 gauge catheters listed in the materials list over those presented in the video because they are disposable and a new catheter can be used for each mouse. They are also a somewhat softer material and therefore less likely to induce trauma on placement.
Detailed procedural video demonstrating the BioLite intubation technique.
Torres-Gonzalez, E., Helms, M. N., Goodson, P., Rojas, M. Direct Tracheal Instillation of Solutes into Mouse Lung. J. Vis. Exp. (42), e1941, DOI: 10.3791/1941 (2010).
Detailed procedural video demonstrating the surgical tracheal injection technique
Video demonstrating blood collection from facial vein/artery using lancet
Video demonstrating blood collection from facial vein/artery using needle