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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Curr Protoc Microbiol. Author manuscript; available in PMC 2014 February 1.
Published in final edited form as:
PMCID: PMC3643812

Retinal gene delivery by rAAV and DNA electroporation


Ocular gene therapy is a fast growing area of research. The eye is an ideal organ for gene therapy since it is immune privileged, easily accessible, and direct viral delivery results primarily in local infection. Because the eye is not a vital organ, mutations in eye specific genes tend to be more common. To date, over 40 eye specific genes have been identified which harbor mutations that lead to blindness. Gene therapy with recombinant Adeno Associated Virus (rAAV) holds the promise to treat patients with such mutations. However, proof-of-concept and safety evaluation for gene therapy remains to be established for most of these diseases. This unit describes the in vivo delivery of genes to the mouse eye by rAAV-mediated gene transfer and plasmid DNA electroporation. Advantages and limitations of these methods are discussed, and detailed protocols for gene delivery, required materials, as well as subsequent tissue processing methods are described.

Keywords: Retina, eye, gene therapy, gene transfer


Gene therapy in the eye thus holds promise to treat many genetic and age-related blinding diseases. The eye is considered a prime target for gene therapy since it is a relatively isolated and immune privileged organ. The field of ocular gene therapy received its biggest boost in 2001. Dogs, whose eyes are comparable to humans in size, were successfully treated for Leber’s congenital amaurosis-2 with a rAAV vector (Acland et al., 2001). The disease is caused by a mutation in the retinal-pigmented epithelium (RPE) protein 65. One of the dogs, named Lancelot, became a media star when he visited Congress.

Delivery of rAAVs to the eye is accomplished either by intravitreal or by subretinal injection. Intravitreal injections infect preferentially cells closer to the ganglion cell layer, such as ganglion cells and inner nuclear layer (INL) cells, while subretinal injections tend to infect photoreceptors (PR) and RPE cells. INL cells can also be targeted by subretinal injections depending on the titer and serotype. The protocol describes both, the subretinal and intravitreal injection methods for delivery of rAAV to the mouse eye. Injection methods into both early postnatal (Basic Protocol 1, Alternate Protocol 1) and adult (Basic Protocol 3) mouse eyes are described. In addition, delivery of plasmid DNA by electroporation at postnatal day 0 is also described (Matsuda and Cepko, 2004; Basic Protocol 2). This method targets mainly dividing cells, therefore only cells born from the time of electroporation onwards will be transduced. This includes rod photoreceptor, bipolar cells, Muller Glia cells, amacrine cells and at very low frequency horizontal cells. Electroporation of plasmid DNA circumvents viral production and can thus be used as a fast method to test the promoter activity of the viral construct or to test if overexpression of a protein may have a beneficial effect in a retinal degenerative disease model. However, it is not a viable therapeutic approach to treat humans, nor is it useful to test cell tropism of rAAVs. The protocols presented here discuss advantages and disadvantages of these different methods and describe injection tools that accommodate different budgets (Alternate Protocol 1). Additionally, tissue preparation (Support Protocols 1 and 2) and processing for immunofluorescence (Basic Protocols 5 and 7) and in situ hybridization analyses (Basic Protocols 6 and 8) on either whole mount or cryo- and paraffin sections are described. Basic Protocol 4 describes the use of a fundus scope to monitor the transduced retinal cells.


  1. 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 humans. BSL-1 status assumes that the rAAV construct does not encode for a gene that is toxic or cancerogenic and that it is produced without a helper virus. Follow all appropriate guidelines and regulations for the use and handling of pathogenic microorganisms. The Institutional Biosafety Committee at the institution where the research is being conducted should approve all Biosafety protocols.
  2. rAAV-mediated gene transfer to the eye is an Animal Biosafety Level 1 (ABSL-1) procedure. Protocols using live animals must be reviewed and approved by the Institutional Animal Care and Use Committee (IACUC) and must adhere to governmental regulations regarding the use and care of animals. The protocols described here have been approved by the IACUC committee of the University of Massachusetts Medical School and conform to officially approved procedure for proper care and use of laboratory animals.

BASIC PROTOCOL 1 Delivery of rAAV by subretinal & intravitreal injection into eyes of newborn mice

This protocol describes the delivery of rAAVs to the subretinal or intravitreal space of newborn mice. The advantage of subretinal delivery of virus or plasmid DNA at birth is that PR outer segments are not formed yet. This means that the subretinal space, between the RPE and the ONL, is an actual space in which fluid can be injected and can spread. The biggest advantage of this method is thus that the entire retinal surface area can be infected. However, at the same time this poses the following disadvantages. First, the eye is smaller in size and targeting the subretinal space correctly may be more challenging. Second, since the retina is still developing, injections that result in too much damage may complicate the interpretation of your results if development is interrupted. For example, injections with phosphate buffered saline (PBS) in a retinal degeneration model may result in a protective effect (delay of photoreceptor cell death), from the physical damage to the tissue. Thus if you study photoreceptor degeneration, it is important to perform enough control injections to account for technical variations of the procedure. An extreme case of neuroprotection occurs when too much fluid is injected into the subretinal space. In such a case, the adult retina can take on the shape of a cone instead of a half sphere. This tends to lead to a more profound protective effect. Both procedures, intravitreal and subretinal, can result in cataracts, and in the worst case in an arrest of eye development.

Intravitreal injections tend to be easier since the targeting area is larger. Injections can be performed with glass needles or metal needles (Hamilton) and in both cases the injection route can be either at the intersection of the cornea and sclera or through the sclera. Injections with a glass needle through the sclera targets directly the subretinal space. If you prefer that route for vitreal delivery, you need to push the needle through the retina. Injecting at the junction of the cornea and sclera targets the vitreous. However, the same route can also be used for subretinal injections. Basic Protocol 1 will detail injections with glass needles either through the sclera for subretinal injections or through the cornea-scleral margin for subretinal and vitreal injections. This is the most effective route and the one we recommend for delivery. The proper use of fine glass needles results in normal retinal morphology, since it is the least invasive procedure. The alternative protocol will present different tools such as Hamilton syringes and different routes of injection. As you develop your skills, you may prefer one method over the other.


  • Dissecting Scope with appropriate light source
  • Ice bucket
  • Glass needles (Humagen: Custom O)
  • Heating plate/mat
  • Weight trays
  • Injection pump (Eppendorf: FemtoJet)
  • Pipette
  • Microloader (pipette tips to load the glass needle)
  • 30-gauge disposable needles
  • 1 pair of forceps (student Dumont #5 work well)
  • Cotton swabs
  • Tissue towel
  • Fast-Green solution (0.05% in H2O)
  • 70% Ethanol
  • Betadine
  • Gloves
  • Buprenorphine
  • Insulin injection needle
  • Virus: rAAV can be produced in-house (Current Protocols Microbiol Chapter 14: Unit14D 11) (Mueller et al., 2012), or obtained through a local vector core or via a commercial manufacturer.

Narrative summary of neonatal injection procedure: Inject neonatal pups subcutaneously with Buprenorphine (0.1 mg/kg) one hour prior to the procedure. After one hour anesthetize pups by hypothermia by placing the pup onto a dry rubber glove over ice. After 2–3 minutes place the pup onto a clean paper towel under the dissecting scope. Clean the skin over the eyelid with Betadine, followed by water and 70% ethanol using cotton swabs. Cut the skin over the eye with a sterile 30-gauge needle in the area where the future eyelid develops. If performed properly the incision will not result in bleeding as this region is undergoing cell death. Push back the skin gently to the side with a pair of sterile forceps to expose the eyeball. Inject by inserting a beveled glass needle directly into the eyeball. Close the eyelid gently with a cotton swab soaked with Betadine and place the pup onto a warm heating mat until fully recovered and then return it to the mother.

  • 1
    Remove mouse pups from the mother (all at once) and inject each pup subcutaneously with Buprenorphine (0.1mg/kg) using an insulin injection needle.

    Buprenorphine is an analgesic that alleviates pain during and after the procedure.

  • 2
    Place pups in a weight balance tray on a warm (37°C) heating mat for 1 hour.
  • 3
    Meanwhile set up your injection station (Movie 1).
  • 4
    Prepare virus mix: Per 10μl virus solution add 2μl of 0.05% Fast Green in a centrifuge tube.

    The titer should be at least 5×1011 genome copies/ml, however we recommend injecting with a titer of 1012 – 5×1013 genome copies/ml, depending on how many cells you intend to transduce.

  • 5
    Spin mixture at maximum speed for 2′ at RT. This will remove any small debris that could clog the glass needle.
  • 6
    Transfer supernatant into a fresh centrifuge tube and place on ice.
  • 7
    After 1 hour anesthetize one pup on ice for 2–3′ by placing it on a rubber glove over ice. Ice anesthesia has multiple advantages. Pups recover faster and reducing the body temperature results in vasoconstriction, which reduces potential bleeding from the procedure. Additionally, reducing the body temperature results in an opaque lens, which helps visualizing the eyeball under the skin.
  • 8
    Meanwhile load the glass needle with a microloader (load 10μl of virus solution) and mount onto handheld injection devise.
  • 9
    Once the pup is anesthetized move it onto a paper towel under the dissecting scope, placing the pup on its side with one eye facing up.
  • 10
    Clean the skin over the eye with a cotton swabs soaked in Betadine, followed by water and the 70% Ethanol.
  • 11
    With the thumb and the index finger of your left hand (if you are right handed) stretch the skin over the eye while gently moving a 30-gauge needle with your right hand over the scar tissue of the future eyelid (Movie 1). Hold the needle such that the beveled edge functions as knife (you can also use a scalpel for this step). It is not necessary to cut completely through the skin, as stretching the skin with your left hand will open up the cut.

    If you cut to deep you may damage the cornea and if you cut too close to the edge of scar the tissue may bleed.

  • 12
    Once you have a small opening, insert a pair of forceps in a closed position and let it open gently. This will further open the initial incision to the edge of the future eyelid without bleeding (Movie 1).

At this point there are two options depending on the injection route and injection needle. You may pop out the eyeball to better expose the sclera for injections into the subretinal space from the sclera (recommended). You may also choose to use the same procedure to inject into the vitreous by pushing your needle through the retina. To use these procedures continue with step 13 (Movie 1).

For direct injections into the vitreous without damaging the retina you don’t need to pop out the eyeball, you can just push the skin to the side with your thumb and index finger. You can also use this procedure to inject into the subretinal space. To perform the procedure without popping out the eyeball continue with step 19.

  • 13
    Injections by popping out the eyeball: Push the skin to the side with your thumb and index finger and with the forceps in your right hand push the skin further to the side to expose the eyeball. You can either move your forceps from the left to the right of the eye by pushing down on the skin or open the forceps and push simultaneously left and right. Once the eyeball pops out, hold the skin pushed down with your thumb and index finger (Movie 1).
  • 14
    Inject into the subretinal space with your right hand. Hold the needle tip perpendicular to the sclera to optimize the force at the tip of the needle. Gently push the needle into the sclera. If an air pump is used for injections and the back flow pressure is set to 0, after entering the eyeball the solution in the needle (which appears blue due to the Fast Green) will be slightly pushed back by the intraocular pressure. This indicates that you are in the eye (Movie 1).
  • 15
    Inject slowly 0.5–1μl of virus solution.

    IMPORTANT: Follow visually the (blue) solution when injecting. If the center of the eye turns immediately blue, then your needle entered too far and the needle tip crossed the retina, meaning you are injecting into the vitreous (Fig. 1a). If you see the sclera bulging in a blue color then your needle tip did not cross the entire sclera and you are not in the subretinal space yet. If the solution is spreading slowly across the eyeball as seen through the lens then you are in the subretinal space (Fig. 1b). If you inject too much volume, once you pull out the needle, the pressure in the eye may push back some of the injected material. This may be the case if you attempt to transduce the entire retina. TRICK: Before injecting the virus poke a hole into the sclera with the glass needle. Afterwards, reposition the needle and perform the injection. The first hole will allow some of the pressure to escape while you are injecting.

    Figure 1
    Basic Protocol 1. Schematic of injection routes for newborn mouse pups using glass needles. (A–D) Cartoons of mouse eyes showing sclera, choroid, RPE, cornea, lens, retina, subretinal space, and injected solution (blue). (A, B) Example of injection ...
  • 16
    After pulling back the needle close the eyelid with a cotton swabs soaked with Betadine.
  • 17
    Place the pup back onto a tray on a heating mat at 37°C. Mice will start to gasp within 2–3 minutes after removing them from the ice. The color of the anesthetized pup turns purple and once the pup recovers it turns pink. It takes less than 1 minute to inject one eye. Viral injections can be performed into both eyes. If the pup is waking up while you are attempting to inject the second eye place it briefly back on ice.
  • 18
    Repeat the procedure with the entire litter. Once all pups are injected and fully recovered return them to the mother all at once.
  • 19
    Injections without popping out the eyeball: Push the skin to the side with your thumb and index finger of your left hand to expose the cornea and part of the sclera.
  • 20
    To inject into the vitreous insert the needle with your right hand at the margin of the cornea and the sclera (Fig. 1c). Inject slowly using titers and volumes as described above. The Fast Green dye should immediately cover the space under the lens rather than spreading slowly through the eye. Hold the needle such that it passes by the lens without damaging the lens. After injecting remove the needle and follow steps 16–18.
  • 21
    To inject into the subretinal space either target the visible area of the sclera or the margin between the cornea and the sclera (Fig. 1d). Hold the needle at the appropriate angle to target the subretinal space without damaging the lens (Fig. 1d). Inject slowly using titers and volumes as described above. The Fast Green dye should spread slowly across the eyeball. After injecting remove the needle and follow steps 16–18.

BASIC PROTOCOL 2 Delivery of plasmid DNA by subretinal injection into eyes of newborn mice

This protocol describes the transduction of retinal cells by electroporation of plasmid DNA. Since electroporation of plasmid DNA into mitotic cells is more efficient than into postmitotic cells the DNA needs to be delivered in proximity to dividing cells before retinal development is completed. Dividing cells in the retina are located close to the subretinal space thus the same procedure as described in Basic Protocol 1 can be used. At postnatal day 0 mitotic cells give rise to rods, Muller Glia, bipolars, and amacrines. Electroporation at postnatal day 0 will thus result in transduction of only these cell types once retinal development is completed. Because most of the postnatal cells are born within a short window of time after birth, electroporation of plasmid DNA needs to be performed ideally within the first 24h after birth. Electroporations at postnatal day 3 will barely yield any transduced cells. This contrasts transduction of cells by rAAV infection, which can be performed at any time.


Same material as described in Basic Protocol 1 with the following exceptions:

  • Replace the virus mixture with a mixture of DNA and Fast Green. Use DNA at a concentration of 2μg/μl.
  • PBS (Phosphate buffered saline)
  • Square-filed Electroporator (e.g. Harvard Apparatus, cat. # 450052)
  • Tweezer electrodes (e.g. Harvard Apparatus, cat. # 405166)
  1. Follow basic Protocol 1 until step 15. Inject a volume of 0.5–1μl of 2μg/μl plasmid DNA mixed with Fast Green. Use the procedure for subretinal injections. Try to inject the DNA as close as possible to the central region of the retina.
  2. Close the eyelid with a cotton swab soaked in PBS.
  3. Wet the tweezer electrodes in PBS (PBS increases conductivity between the electrode and the skin) and place them around the head of the pup such that each electrode pad covers one eye. When placing the tweezer electrodes pay attention to the position of the electrode pads relative to the site of DNA delivery (Fig 2a). The DNA will move in the direction of the electrical field that is generated between the electrode pads and move along the path of least resistance. If you injected your DNA in the periphery of the eye, and the electrode pads are placed straight over the eye then the DNA may move along the subretinal space instead of moving into the retina (Fig. 2b). If you inject in the center of the eye then placing the electrode pads straight over the eye is the correct approach (Fig. 2a). If you injected laterally,
    Figure 2
    Basic Protocol 2. Positioning of tweezer electrodes for DNA electroporation at postnatal day 0. (A) Shows correct position of electrode pads when DNA is injected into the central region of the retina. (B) Shows wrong position of electrode pads for peripheral ...
  4. then rotate the electrode pads around to generate an electrical field that is perpendicular to the retina and the injection site (Fig. 2c). ATTENTION: the plus pole (+) of the electrode needs to be over the eye you injected with the DNA.
  5. Apply electrical field: 5 pulses at 80V. Each pulse should be 50msec long with an interval of 950msec.
  6. Follow steps 16–18 described in Basic Protocol 1.

    ATTENTION: In contrast to viral injection it is recommended that for electroporation of plasmid DNA only one eye be injected. Injecting and electroporating the second eye generates an electrical filed in the opposite direction to the first eye and may reduce the efficiency of electroporation of the first eye.

BASIC PROTOCOL 3 Delivery of rAAV by subretinal & intravitreal injection into the eye of adult mice

This protocol described the delivery of rAAVs into the eye of adult mice. The delivery procedure is similar to the one described in Basic Protocol 1. Since the eye is already exposed, popping out the eyeball is not required. Subretinal injections into adult wild-type mice will always lead to retinal detachment, since the PR outer segment and RPE interactions that exist in adult mice are disrupted by the fluid that is injected. This contrasts injections into newborn mice, as the retina is not attached to the RPE at that age because outer segments are not developed yet. Injecting mice with PR degeneration reduces the amount of retinal detachment caused by the injection, as part of the retina may already be detached due to the disease. However, this increases the efficiency of the viral spread. Targeting the vitreous of adult mice is straightforward. Injecting adult mice has the advantage that development is completed, which reduces some of the undesired procedural effects. Nonetheless, any injection, even PBS, can result in the release of various neuroprotective growth factors. Therefore, when injecting a virus that should result in a neuroprotective effect, enough control injections need to be performed to account for artifacts.


  • Dissecting Scope with appropriate light source
  • Glass needles (Humagen: Custom O)
  • Injection pump (Eppendorf: FemtoJet)
  • Pipette
  • Microloader (pipette tips to load the glass needle)
  • Cotton swabs
  • Tissue towel
  • Fast-Green solution (0.05%)
  • Virus
  • 70% Ethanol
  • Corneal lubricant ointment
  • Betadine
  • Gloves
  • Ketamine
  • Xylazine
  • Buprenorphine
  • Insulin injection needle
  • Heating plate/mat
  1. Anesthetize mouse by an intraperitoneal (IP) injection of a Ketamine/Xylazine (100 mg/kg and 10 mg/kg) mixture. Test depth of anesthesia by a sharp tail pinch.
  2. Place mouse on a clean paper towel under the dissecting scope and apply corneal lubricant ointment to protect the cornea. Then clean the eyelid with Betadine, followed by water and 70% ethanol.
  3. Gently push down the skin of the eyelid to better expose the eyeball.
  4. Insert the glass needle from the scleral side if you intend to inject into the subretinal space. Inject virus as described in Basic Protocol 1 with the same recommended titer.
  5. If you intend to target the vitreous, insert the needle at the margin of the sclera and cornea to inject the virus.
  6. After injection clean the eyelid and inject mouse subcutaneously with Buprenorphine (0.1 mg/kg) to alleviate pain.
  7. Place mouse back into its cage and place cage onto a warm heating plate (37°C) until the mouse is fully recovered.

ALTERNATE PROTOCOL Subretinal & intravitreal injection with Hamilton syringes

This protocol describes an alternate injection tool and route of delivery for injection into newborn mice. The procedure is very similar to the one described in Basic Protocol 1 and can also be used for electroporation of plasmid DNA or for vitreal injections into adult mice. A less costly way to perform the experiment as described in Basic Protocol 1 is to use a handheld pipetteman that allows mounting a glass needle. Alternatively, a Hamilton syringe, which is also less costly, can be used instead of a glass needle with an injection pump. Here we discuss the use of a Hamilton syringe in combination with a blunt metal needle. Pointy (beveled) metal needles are also available for Hamilton syringes. These needles can be used to perform the injection as described in Basic Protocol 1. The advantage of metal needles is that they do not break easily. However, given the larger size of the needle there is more damage to the tissue. The procedure below explains the use of the Hamilton syringe with a blunt metal needle. This requires a slightly modified procedure to the one explained in Basic Protocol 1.


  • Dissecting Scope with appropriate light source
  • Ice bucket with ice
  • Hamilton syringe with blunt needle
  • Heating plate/mat
  • Weight trays
  • 30-gauge disposable needles
  • 1 pair of forceps (student Dumont #5 work well)
  • Cotton swabs
  • Tissue towel
  • Fast-Green solution (0.05%)
  • Virus or DNA
  • If DNA: Square-filed Electroporator, Tweezer electrodes and PBS
  • 70% Ethanol
  • Betadine
  • Gloves
  • Buprenorphine
  • Insulin injection needle
  1. Follow Basic Protocol 1 until step 12.
  2. Push the skin to the side with your thumb and index finger of your left hand to expose the cornea of the eye and part of the sclera.
  3. With the 30-gauge needle poke a small hole at the margin of the sclera and cornea. This step is necessary because the Hamilton syringe has a blunt needle tip.
  4. Insert the needle loaded with virus or DNA into the hole and move the needle tip passed the lens towards to central part of the eye (Fig. 3a). Inject virus if the vitreous needs to be targeted. After injecting the virus pull back the needle and continue with Basic Protocol 1 step 16.
    Figure 3
    Alternative Protocol. Schematic of vitreal and subretinal injection routes into newborn mouse pups using a Hamilton syringe with a blunt needle. (A, B) Cartoons of mouse eyes as shown in Fig 2. (A) Example of injection route for vitreal injection. (B) ...
  5. If you want to target the subretinal space push the needle further in until it crosses the retina (Fig. 3b). Because the needle is blunt it will not cross the sclera. This means that once you hit resistance you can inject the fluid, which will disperse into the subretinal space. This procedure causes more damage as you physically create a hole in the retina through which the virus or DNA is injected.
  6. After pulling back the needle continue with Basic Protocol 1 step 16 if you are injecting a virus or with Basic Protocol 2 step 2 if you are injecting plasmid DNA.

BASIC PROTOCOL 4 Fundoscopy examination to monitor infection area

This protocol describes the use of a fundus scope to visualize retinal cells that have been transduced with a green fluorescent protein (GFP) (Fig. 4a). The technique is non-invasive and allows acquiring retinal photographs of mice that are anesthetized with a Ketamine/Xylazine mixture. Visualizing the area of infection or electroporation is only possible if the expression cassette of your transgene also co-expresses GFP. The advantage of this protocol is that it allows selecting well infected or electroporated animals for further analysis. For example, 30 mice are injected with an rAAV that should delay PR death. Behavioral tests and/or electroretinograms need to be performed to test if the viral transgene leads to improved vision. Preselecting the 10 best-infected mice reduces the overall workload. Additionally, fewer mice need be kept for extend periods of time if long-term effects of the viral transgene are to be tested. However, this procedure is not required to perform histological analyses of GFP transduced retinas. We do not recommend purchasing such equipment to perform the gene delivery protocols described here. Its use is recommended if your institute owns such equipment and if the experimental design benefits from such use.

Figure 4
Basic Protocol 4. Transduction efficiency as seen by fundoscopy. (A) Fundus image showing large area of GFP positive cell that were infected with a GFP expressing rAAV(2/5). Similar images can be seen after electroporation of a GFP expressing plasmid. ...


  • Fundus Scope equipment:
    • Appropriate light source
    • Platform to position mouse
    • Computer system and software
  • Phenylephrine
  • Tropicamide
  • Gloves
  • Ketamine
  • Xylazine
  • Insulin injection needle
  • Heating plate/mat
  1. Dilate pupils using one drop of Phenylephrine (quick and short effect) and one drop of Tropicamide (slow and long effect).

    Since anesthesia decreases the body temperature of the mouse, which can cause the lens to become temporarily opaque, dilating the pupil in advance allows starting immediately after the mouse is anesthetized.

  2. Once the pupils are dilated (5 to 10 minutes later) anesthetize mouse by an intraperitoneal injection of a Ketamine/Xylazine (100 mg/kg and 10 mg/kg) mixture. Test depth of anesthesia by a sharp tail pinch.
  3. Place mouse on fundus scope platform (Fig. 4b). A heating pad can be used to keep the mouse warm, which helps keeping the lens transparent. This extends the period of time to perform a better examination. However, with some experience the procedure is performed within minutes. Therefore, keeping the mouse warm during the procedure is not required.
  4. Place camera directly on one eye by moving the platform into the appropriate position
  5. Select appropriate wave-length and focus image.
  6. Moving the platform around will allow you to see different areas of the retina.
  7. Acquire movie or individual images.
  8. Place mouse back into its cage and place cage onto a warm heating plate (37°C) until the mouse is fully recovered.

BASIC PROTOCOL 5 Whole mount immunofluorescence analyses

This protocol describes the processing of the retina for whole mount analysis. There are two ways the retina can be dissected and prepared depending on the cell type that needs to be visualized. Both methods will be introduced at the beginning. Retinal cells can then be visualized either by whole mount immunofluorescence or immunocytochemistry. These methods will be discussed in this protocol and rely on the use of an antibody that is either cell type specific or directed against the protein that is over-expressed as a result of the transduction of retinal cells. If no antibody is available, whole mount in situ hybridizations can be used to detect either the viral RNA or the mRNA of a cell type specific gene. This procedure will be described in Basic Protocol 6.


  • CO2 Chamber for euthanasia
  • Dissecting scope with light source
  • Tissue towels (Kimwipes)
  • Petri dish (6cm)
  • Two pairs of forceps (student Dumont #5 work well)
  • Small Spring scissor
  • Transfer pipettes
  • Pipettes and pipette tips
  • PBS (Phosphate Buffered Saline; pH 7.4)
  • 4% PFA/PBS (Fixative: 4% paraformaldehyde in PBS; pH 7.4)
  • PBT (PBS with 0.3% Triton-X100)
  • PBTB (PBT with 5% BSA)
  • Cover glass
  • Glass slides
  • Primary and secondary antibodies
  • Mounting media: Gel mount, Fluoromount, etc

Protocol steps

Dissect eye in PBS by removing the retina from the rest of the tissue. If target cells are photoreceptors, leaving the retina attached to the lens allows fixing the retina such that it retains its shape as a cup. The lens can be left on during the entire procedure and removed prior to mounting the retina. This avoids curling of the retina and results in better accessibility of the antibody at the periphery of the tissue. To target inner nuclear layer cells and ganglion cells the lens has to be removed. If removed after fixation, most of the curling is prevented. Alternatively, if INL cells are targeted the retina can be left onto the lens and the incubation with the primary and secondary antibody can be performed over a period of 2–3 days each at 4°C. A longer incubation time allows for better penetration of the antibody to the cell in the center of the tissue. However, the length of time needs to be established for each individual antibody. If not otherwise indicated, all steps are performed at room temperature.

  1. Euthanize mouse with CO2 or alternative procedure that has been approved by the IACUC of your institute. Perform a double-kill procedure (e.g. cervical dislocation).
  2. Remove the eyeball (enucleate) by first pushing down the skin around the eyeball. Then, with a pair of forceps in open position slide each arm of the forceps around the eyeball all the way down to the optic nerve. Close the forceps to hold the eye at the bottom (optic nerve) and pull out the eyeball. Part of the optic nerve and some muscle and connective tissue that regulate eye movement may be attached to the eyeball. Place eye in a 6cm petri dish filled with PBS. I would recommend submerging the eye completely in PBS. At any time during procedure try to avoid squeezing the eyeball as this may damage the retina (Movie 2).
  3. Steps 3–5 are performed under a dissecting scope. With your right hand (if you are right handed) poke a hole at the margin of the sclera and cornea using one arm of the forceps as needle. You may also use a needle for this step. To hold the eyeball stable, with your left hand open the forceps and place it behind the optic nerve. While you are pushing with your right hand, the eye will not roll away as the forceps in your left hand will hold it in place (Movie 2).
  4. Once one arm of the forceps has entered into the eyeball close the forceps. Insert one arm of the other forceps (left hand) into the hole and close forceps. Gently move forceps apart to tear the eyeball along the sclera-corneal margin. Reposition the forceps along the tear and continue until cornea is removed.
  5. Insert one arm of each forceps between retina and sclera. Close forceps and gently tear the sclera apart to expose the retina. Once the retina is exposed up to the optic nerve pinch off the optic nerve without damaging the retina. Continue to remove the sclera until the retina is completely exposed. If the procedure worked well your retina will stay attached to the lens (Movie 2).
  6. Transfer tissue to a 4% PFA solution in PBS (4%PFA/PBS) using transfer pipettes and fix for 30 min at RT. This step can be done in a 1.5ml centrifuge tube or scintillation vial. For this and all following incubation steps it is recommended that the tube/vial be in gentle motion (rocker, nutator, etc).
  7. Wash 3 × 10′ in PBS.
  8. If your antibody staining will target photoreceptors, you can leave the lens attached. In this case proceed with step 10.
  9. If you need to target ganglion cells remove the lens from the retina. Place the tissue back in to a petri dish with PBS. With both forceps pinch the outer membrane of the lens (lens capsule) and tear open the capsule to remove the lens. Then gently remove the rest of the tissue (lens capsule and ciliary margin) from the retina (Movie 2). Place the retina back into a centrifuge tube or scintillation vial.
  10. Permeabilize tissue for 30′ in PBT.
  11. Block for 1h in PBTB.
  12. Incubate over night at 4°C with primary antibody diluted in PBTB.
  13. Wash 3 × 20′ in PBTB.
  14. Incubate for 2h with secondary antibody diluted in PBTB.
  15. Wash 4 × 30′ in PBTB.
  16. If your secondary antibody requires a colorimetric reaction (e.g. antibody is coupled to alkaline phosphatase or horseradish peroxidase) proceed according to manufactures recommendation for such reaction. Otherwise mount retina as described in step 17.
  17. If you have not removed the lens yet, remove the lens as described in step 5. After removing the lens place the retina on a cover glass (photoreceptor side down, ganglion cell side up). Ensure the retina is covered with enough fluid (large drop). Under a dissecting scope, with a small scissor make 4 incisions at an angle of 90° (Movie 2). Incision should be half the distance from the optic nerve head to the periphery. To flat mount the tissue you will need to roll each quadrant outward. For this, decrease first the volume of the drop by holding a kimwipe into the drop. Remove fluid until retina is barely covered. Then take two kimwipes and roll the corner of each sheet between your fingers to generate a pointy tip. Wet the tip with PBT and roll again to remove excess fluid. Using the two kimwipe tips gently roll the retina flat. Once the retina is flat and looks like a clover leaf, add mounting media to one edge of the cover glass. Do not add it directly on top of the retina as the quadrants may curl back. Use a second cover glass and slowly move it downwards starting from the edge where you placed the drop. The retina is now mounted between two cover glasses. Depending on the cell type you need to image, place the cover glass sandwich on a glass slide with cell type of interest facing up. To prevent the cover glass sandwich to move around, wet the glass slide. Seal the edge of the cover glass if desired with nail polish. (To prevent the retina from being squeezed too much between the cover glasses you can also break a third cover glass and place a piece of glass left and right of the retina. The glass functions as a spacer between the two cover glasses).
  18. Analyze retina with a fluorescent upright or inverted microscope. Depending on the intensity of the staining a minimum magnification of 10X and higher may be required. If the fluorescent staining is intense with a low background the fluorescent flat mount can also be imaged on a fluorescent dissecting scope.

BASIC PROTOCOL 6 Whole mount in situ hybridizations

This protocol is very similar to Basic Protocol 5 however, gene expression is revealed by in situ hybridization. We recommend removing the lens the latest after the hybridization step even if PRs are targeted. This avoids background from probe that is trapped between the lens and the retina. Similar to the antibody staining protocol, the incubation steps work best if the tissue is in motion during incubation. Make sure that all your solutions up to the hybridization step are free of RNAse. Treat PBS over night with DEPC (Diethylpyrocarbonate, dilution 1:1000) and autoclave next day. Make all your solutions for Day 1 in DEPC treated PBS (except 4% PFA/PBS). If not otherwise noted all steps are performed at room temperature.


Reagents for RNA probe synthesis

  • Nucleotide labeling mix
  • RNA polymerase (T7, T3, SP6)
  • RNAse inhibitor
  • DNAse (RNAse free)
  • TE (10mM Tris pH8, 1mM EDTA)
  • Ethanol 100%
  • H2O (nuclease free)
  • LiCl 4M (DEPC treated)
  • HB (Hybridization buffer)
  • Dissecting scope with proper illumination
  • CO2 Chamber for euthanasia
  • Tissue towels (Kimwipes)
  • Petri dish (6cm or 10cm)
  • Two pairs of forceps (student Dumont #5 work well)
  • Small Spring scissor
  • Transfer pipettes
  • 1.5 ml centrifuge tubes
  • Scintillation vials
  • Heating Block
  • Rocker or Nutator
  • Water bath or incubation oven
  • Cover glass
  • Glass slide
  • PBS (Phosphate Buffered Saline; pH 7.4)
  • 4% PFA/PBS (Fixative: 4% paraformaldehyde in PBS; pH 7.4)
  • Methanol 100%
  • Proteinase K (Stock: 10mg/ml; keep at −20°C)
  • PBT (PBS that is treated with DEPC with 0.1% Tween-20)
  • HB (Hybridization buffer: 50% Formamide; 1.3x SSC; 5mM EDTA; 0.2% Tween-20; 0.5% CHAPS, 100μg/ml Heparin; 50μg/ml Yeast tRNA)
  • 20x SSC (Saline-Sodium Citrate)
  • Tris-HCl 1M pH9.5
  • PIPES 1M pH6.8
  • MAB (100mM Maleic acid pH7.5; 150mM NaCl)
  • MABT (MAB; 0.1% Tween-20)
  • α-Digoxigenin antibody coupled to alkaline phosphatase (α-DIG-AP)
  • NTMT (100mM NaCl; 100mM Tris-HCl pH9.5; 50mM MgCl2; 0.1% Tween-20)
  • Dimethylformamide (DMF)
  • NBT: 4-Nitro blue tetrazolium chloride; Stock (50x): 50mg/ml in 70% DMF (store at −20°C)
  • BCIP: 5-Bromo-4-chloro-3-indolyl phosphate; Stock (50x): 25mg/ml in water (store at −20°C)
  • Staining solution: NTMT; 1x BCIP; 1x NBT
  • EDTA 200mM pH8
  • Glycerol

RNA probe synthesis

Preparation of linearized DNA or PCR product is not described here, nor are the reagents and the equipment needed listed. Purify linearized DNA either by precipitation or by gel electrophoresis. Purify PCR product by gel electrophoresis. For probe synthesis we recommend to use PCR products that have been amplified from a standard cloning vector with a combination of the following primers: T7, T3, SP6.

  1. For RNA probe synthesis mix the following reagents and incubate at 37°C for 2h:
    1. 2μl of 10x Transcription Buffer (Roche)
    2. 2μl of 10x Nucleotide Labeling Mix, (Roche: DIG labeling mix)
    3. Xμl of Template (~1μg of linearized plasmid DNA or 300ng of PCR product; not shorter than 200bp, ideally between 500bp-1000bp)
    4. 0.5μl of RNase Inhibitor (Roche)
    5. 1μl of RNA Polymerase (Roche: T7, T3, SP6, depending on the PCR primers used to generate the PCR product; make sure to take the correct RNA polymerase to synthesize antisense RNA)
    6. Yμl of H2O to fill up the volume to 20μl total. Make sure that H2O nuclease free.
  2. After 2h run 1 μl on an agarose gel to ensure that the synthesis reaction worked.
  3. Add 1μl of DNAse (RNAse free; Roche) to remaining 19 μl and incubate at 37°C for 15′.
  4. Add 100μl of DEPC treated TE pH8 (10mM Tris pH8, 1mM EDTA).
  5. Add 10μl of DEPC treated 4M LiCl.
  6. Add 300μl of 100% Ethanol, mix well and incubate over night at −20°C.
  7. Centrifuge at 13,500rpm at 4°C for 15′.
  8. Wash pellet in 70% Ethanol and air-dry pellet for 5–10′.
  9. Resuspend pellet in 20μl of nuclease free H20.
  10. Add 80 μl of warm (65°C) HB, vortex and store at −80°C.
  11. Before each use warm up probe briefly at 65°C and vortex.

in situ hybridization

Tissue preparation

Follow Basic Protocol 5 to dissect the retina (steps 1–5). Fix over night at 4°C or at RT for 3h.

  1. Wash 3 × 10′ in PBT (Attention: Tween-20).
  2. Wash 3 × 5′ in 100% Methanol (Dehydration of tissue).
  3. Store at −20°C for at least 1h (permeabilization of tissue). Better results may be obtained if the tissue is stored for 1 week at −20°C in 100% Methanol. (Tissue can be stored up to several months in Methanol).

Day 1: Pretreatment and hybridization

  1. Rehydrate tissue in decreasing concentrations of Methanol and increasing concentrations of PBT (Methanol concentrations: 75%, 50%, 25%, 0%). Perform each gradient step for 2 × 5′ and 1 × 10′.
  2. Treat tissue with Proteinase K (10μg/ml) in PBT for 20′.
  3. Rinse 2 X with PBT.
  4. Post-fix tissue for 20′ with 4% PFA/PBS.
  5. Wash 4 × 5′ in PBT.
  6. Wash 2 × 5′ in hybridization buffer (HB).
  7. Incubate for 2h in HB at 65°C.
  8. Add 1 μl of probe per 100μl of pre-warmed HB (65°C), vortex tube and replace HB solution with HB solution containing the probe.
  9. Hybridize over night at 65°C.

Day 2: Post-hybridization washes

  1. Remove probe with hybridization buffer and store at −80°C. Probe can be reused 5–6 times.
  2. Wash 2 × 30′ in HB at 65°C.
  3. Wash 10′ in 75% HB and 25% 2x SSC at 65°C.
  4. Wash 10′ in 50% HB and 50% 2x SSC at 65°C.
  5. Wash 10′ in 25% HB and 75% 2x SSC at 65°C.
  6. Wash 10′ in 2x SSC at 65°C.
  7. Wash 30′ in 0.2x SSC at 65°C.
  8. Wash 10′ in 0.2x SSC at room temperature (from here on room temperature).
  9. Wash 10′ in 10mM PIPES, 500mM NaCl.
  10. Wash 10′ in MAB.
  11. Block for 2h in MAB containing 2% blocking reagent (Roche).
  12. Incubate over night at 4°C in MABT with 2% blocking reagent and 1° antibody (α-DIG-AP) diluted at 1:5000.

Day 3: Post washes and detection

  1. Wash 6 × 15′ in MABT
  2. Wash 30′ in 100mM Tris-HCl pH9.5 + 0.1% Tween-20.
  3. Incubate in staining solution in the dark. Depending on the quality of the probe, the expression level of the gene, and the site that was targeted for hybridization, the in situ hybridization signal can be visible after 30′ or can take several hours. Incubation can be extended over night at 4 °C. After signal has been detected, proceed with the following washes to inactivate the enzyme and clear the tissue.
  4. Wash for 10′ in PBST.
  5. Wash for 5′ in PBS with 10Mm EDTA.
  6. Postfix for 20′ in 4% PFA/PBS.
  7. Wash 3 × 5′ in PBT.
  8. Wash for 15′ in 30% Glycerol and 70% PBT
  9. Wash for 15′ in 60% Glycerol and 40% PBT
  10. Wash for 15′ in 90% Glycerol and 10% PBT
  11. Flat mount retina as described in Basic Protocol 5 step 17. Use glycerol as mounting media.

SUPPORT PROTOCOL 1 Dissection and tissue processing for cryo-sectioning

This protocol describes the processing of the retina for cryo-sectioning. There are two ways the retina can be dissected and prepared depending on the need to retain the RPE attached to the retina. If the RPE is not needed, we recommend the dissection method described in Basic Protocol 5, which leaves initially the lens attached. However after fixation, the lens needs to be removed prior to performing the sucrose gradient and the embedding. Leaving the lens attached ensures a nice cup shaped retina. If you need to retain the RPE attached, follow the dissection method described here. Detection of gene expression by immunofluorescence and in situ hybridization on sections will be presented in Basic Protocol 7 and 8 respectively.


  • CO2 Chamber for euthanasia
  • Dissecting scope with light source
  • Cryostat
  • Tissue towels
  • Petri dish (6cm)
  • Two pairs of forceps (student Dumont #5 work well)
  • Small Spring scissor
  • Transfer pipettes
  • 1.5 ml centrifuge tube
  • Scintillation vials
  • Embedding molds
  • Rocker or Nutator
  • PBS (Phosphate Buffered Saline; pH 7.4)
  • 50% Sucrose
  • 4% PFA/PBS (Fixative: 4% paraformaldehyde in PBS; pH 7.4)
  • OCT embedding media (Tissue-TeK)
  1. Euthanize mouse, remove eyeball and dissect retina with lens attached as described in Basic Protocol 5 if RPE is not needed. Then go to step 5. Alternatively if RPE-PR outer segment interactions need to be preserved go to step 2.
  2. Poke a hole at the margin of the sclera and cornea as described in step 3 of Basic Protocol 5. The hole can also be poked within the cornea, which leaves the very peripheral retina intact.
  3. Transfer eyeball to a 4% PFA solution in PBS (4%PFA/PBS) and fix for 10′. This step can be done in a 1.5ml centrifuge tube or scintillation vial. Place tube on rocker or nutator. Poking a hole before fixing the tissue helps to prevent shrinkage of the eyeball and results in faster exposure of the retina to the fixative.
  4. . Transfer eyeball back into a petri dish with PBS and remove cornea and lens under dissecting scope. You can either perform the same procedure as described in Basic Protocol 5 by tearing gently with two forceps the margin between the cornea and the sclera apart. However, we recommend the following procedure. With a small spring scissor enter into the hole and cut gently along the margin between the cornea and sclera until the cornea is removed (Movie 2). If the very peripheral margin of the retina needs to be preserved, we recommend cutting on the corneal side along the margin such that a small rim of the cornea remains attached to the sclera. Try not to touch the sclera, rather hold the eyeball at the optic nerve or any residual tissue that is attached to the eyeball. While cutting with your right hand, hold the eye with a pair of forceps in your left hand. Once the cornea is removed gently pull out the lens. Prefixing the eyeball ensures that the retina does not detach. Removing the cornea by cutting rather than by tearing the tissue results in less retinal detachment that can occur due to the procedure and therefore in better overall morphology.
  5. Fix tissue over night at 4°C in 4% PFA/PBS.
  6. Wash tissue 3 × 10′ in PBS.
  7. Equilibrate tissue in sucrose gradient.
    1. 10% sucrose in PBS for 2–4 hours at 4°C
    2. 20% sucrose in PBS for 2–4 hours at 4°C
    3. 30% sucrose in PBS over night at 4°C
  8. Equilibrate tissue for 10′ in a 1:1 mixture of OCT : 30% sucrose in PBS.
  9. Equilibrate tissue in OCT for 10′. Make sure the eyecup is filled with OCT by pipetting with a P1000 OCT into the eyecup to push out the mixture of OCT and 30% sucrose in PBS. Frozen OCT is harder than the 1:1 mixture of OCT and 30% sucrose in PBS. The difference in density may result in poor overall morphology when sectioning.
  10. Transfer into a mold with OCT.
  11. Freeze mold with OCT and tissue on a mixture of dry ice/isopropanol.
  12. Store block at −80°C until needed. Make sure to store it air-tight.

    Dehydration of the block causes the OCT to take on a rubber like consistency over time.

  13. Section on a cryostat at desired thickness. The average diameter of photoreceptors is around 5–6μm most other retinal cells are slightly larger. We therefore routinely section at a thickness of 14–20μm for frozen sections. Collect sections on pretreated glass slides for cryo-sections.
  14. Air dry sections for at least 30′ and then either proceed with step 16 or store sections at −80°C for later use. If only immunofluorescence analyses are performed sections can also be stored −20°C.
  15. Acclimate sections for 20′ at RT before use.
  16. Rehydrate section by washing 3 × 10′ in PBS.
  17. If you want to perform an antibody staining follow Basic Protocol 7. Alternatively if you need to detect your gene of interest by in situ hybridization follow Basic Protocol 8.

    ATTENTION: If the gene of interest will be detected by in situ hybridizations use for all steps DEPC treated PBS. If after rehydration of the slides the tissue has holes reduce the incubation time of the sucrose gradient. Sucrose leads to swelling and bursting of cells. Adjusting the window of time of fixation and the sucrose gradient will remedy this problem.

SUPPORT PROTOCOL 2 Dissection and tissue processing for paraffin sectioning

This protocol describes the processing of the retina for paraffin sectioning. Perform dissections as described in Support Protocol 1 according to your needs to retain the RPE attached to the retina. The protocol starts after the initial fixation step of Support Protocol 1 but prior to the over night fixation for cryo-sections.


  • CO2 Chamber for euthanasia
  • Dissecting scope with light source
  • Microtome for paraffin sections
  • Incubator/oven (up to 65°C for paraffin)
  • Petri dish (6cm)
  • Two pairs of forceps (student Dumont #5 work well)
  • Small Spring scissor
  • Transfer pipettes
  • 1.5 ml centrifuge tube
  • Scintillation vials
  • Embedding molds
  • Rocker or Nutator
  • PBS (Phosphate Buffered Saline; pH 7.4)
  • 4% PFA/PBS (Fixative: 4% paraformaldehyde in PBS; pH 7.4)
  • Xylene
  • Paraffin
  • Ethanol
  1. Fix tissue harvested either from step 2 or step 5 of Support Protocol 1 for 30′ in 4% PFA/PBS.
  2. Wash 3 × 10′ in PBS.
  3. Dehydrate tissue to 100% Ethanol by increasing concentrations of Ethanol. Scintillation vial work best.
    a. 25% Ethanol/PBS10′
    b. 50% Ethanol/PBS10′
    c. 75% Ethanol/H2O10′
    d. 100% Ethanol10′
    e. 100% Ethanol10′

    If needed tissue can be stored at −20°C for several months in 100% Ethanol.

  4. Clear tissue in Xylene for 2 × 5′.
  5. Incubate for 30′ in a 1:1 mixture of Paraffin/Xylene at 60°C (temperature depends on melting point of paraffin used).
  6. Wash 4 × 30′ with 100% paraffin at 60°C.

    These washes are important to remove all residual Xylene from the tissue. If Xylene is not removed properly, when stretching the tissue in a warm water bath the lower melting point of the Xylene will leave holes in your section.

  7. Incubate over night at 60°C with fresh 100% paraffin.
  8. Mount tissue in appropriate mold with fresh paraffin.
  9. Paraffin blocks can be stored for several months a 4°C.
  10. Section blocks at a thickness of 14–20μm and transfer section into a water bath at 45°C to allow the paraffin to stretch.
  11. Collect section on pre-treated glass slides.
  12. Dry glass slides with sections over night at 37°C and store at 4°C until needed.
  13. When ready, bake slides at 60°C on a heating plate for 1h.
  14. Remove from plate and allow slides to cool down to room temperature for 5′.
  15. Dewax slides for 2 × 5′ in Xylene.
  16. Rehydrate to PBS
    a. 100% Ethanol2 × 5′
    b. 75% Ethanol/H2O5′
    c. 50% Ethanol/PBS5′
    d. 25% Ethanol/PBS5′
    e. PBS2 × 5′

For antibody stainings follow Basic Protocol 7, for in situ hybridizations follow Basic Protocol 8. ATTENTION: If the gene of interest will be detected by in situ hybridizations use for all steps DEPC treated PBS.

BASIC PROTOCOL 7 Immunofluorescence analysis on cryo- or paraffin sections

This protocol describes immunofluorescence analysis on retinal cross-section. The protocol works equally well for cryo- and paraffin sections.


  • Humidified incubation chamber
  • PBS (Phosphate Balanced Solution; pH 7.4)
  • PBT (PBS with 0.3% Triton-X100)
  • PBTB (PBT with 5% of BSA)
  • Primary and secondary antibodies
  • Mounting media: Gel mount, Fluoromount, etc.
  1. Permeabilize tissue with PBT for 30′.
  2. Block with PBTB for 30′.
  3. Incubate with 1° antibody diluted in PBTB either over night at 4°C or for 2h at RT.
  4. Wash 3 × 20′ with PBTB.
  5. Incubate with 2° antibody diluted in PBTB either over night at 4°C or for 2h at RT
  6. Wash 3 × 20′ with PBTB. Add nuclear DAPI stain in the first wash if desired.

    Alternatively, DAPI can also be added with the secondary antibody.

  7. Apply mounting media and cover slide.

BASIC PROTOCOL 8 In situ hybridization analysis on cryo- or paraffin sections

This protocol describes in situ hybridizations on sections. It differs from Basic Protocol 6 since it is optimized for section in situ hybridizations. The same RNA probe synthesis procedure as described in Basic Protocol 6 can be used for section in situ hybridizations. The recipes for solutions that are the same between both protocols are not described here. ATTENTION: The hybridization buffer (HB) for section in situ hybridizations differs from the whole mount in situ hybridization buffer.


  • Acetic acid anhydride
  • TEA (Triethanolamine 1M pH8.0)
  • HB (Hybridization buffer: 50% Formamide; 10mM Tris-HCl pH7.5; 600mM NaCl; 1mM EDTA; 0.25% SDS; 10% Dextran Sulfate, 1X Denhardt’s; 200μg/ml Yeast tRNA)
  • TNE (10mM Tris-HCl pH7.5; 500mM NaCl; 1mM EDTA)
  • HISS (Heat inactivated sheep serum)

Day 1: Pretreatment and hybridization

  1. Post-fix tissue for 10′ in 4% PFA/PBS.
  2. Wash 3 × 5′ in PBT (Tween-20).
  3. Treat tissue for 10′ with Proteinase K (1μg/ml) in PBS.
  4. Wash 2 × 5′ in PBT.
  5. Post-fix in 4% PFA/PBS for 5′ (Re-use PFA from first post-fix)
  6. Wash 3 × 5′ in PBT.
  7. Perform acetylation for 10′ (add 625μl of acetic acid anhydride to 250ml of 100mM triethanolamine.

    Use solution immediately after mixing. Do not prepare in advance!

  8. Wash 3 × 5′ in PBT.
  9. Air dry slides for 10′.
  10. Meanwhile pre-warm probe and hybridization buffer to 70°C.
  11. Mix 1–3μl of RNA probe with 120μl of HB, vortex and add to slide.
  12. Cover glass slides either with glass cover slides or with home-made cover slips from polypropylene bags.

    Plastic cover slides are easier to remove and do not sheer the sections.

  13. Place slides in humidified slide box (use paper towels soaked with water and place them at the bottom of the box. Seal box with plastic tape. Hybridize in oven over night at 65°C.

Day 2: Post-hybridization washes & detection (Use pre-warmed solutions for washes at 65°C)

  1. Remove cover slides by immersing slides in 5x SSC at room temperature.
  2. Wash for 30′ in a 1:1 mixture of 1x SSC and Formamide at 65°C.
  3. Wash for 10′ in TNE at 37°C.
  4. Treat for 30′ with RNase A (20μg/ml in TNE) at 37°C.
  5. Wash for 10′ in TNE at 37°C.
  6. Wash for 20′ in 2x SSC at 65°C.
  7. Wash 2 × 20′ in 0.2x SSC at 65°C.
  8. Wash 2 × 5′ in MABT.
  9. Block for 30′ in MABT with 20% HISS.
  10. Incubate in humidified chamber over night at 4°C or for 2h at room temperature with MABT containing 20% HISS and 1° antibody (α-DIG-AP) diluted at 1:2500.
  11. Wash 4 × 15′ in MABT
  12. Wash for 10′ in NTMT pH9.5
  13. Add staining solution to slides and incubate in the dark. Signal may take 30′ to several hours to develop. If necessary, continue incubation over night at 4°C or incubate from the beginning over night at 4°C.
  14. Rinse with NTMT pH9.5
  15. Post-fix for 30′ in 4% PFA/PBS to inactivate AP.
  16. Wash 2 × 5′ in PBS.
  17. Mount in desired mounting media (e.g. Gelvatol).


Reagents and solutions are described in each protocol. The following stock solutions are recommended:

  • 10x PBS (DEPC treated; to each liter of 10x PBS add 1ml of DEPC, stir over night, remove magnetic stirrer and autoclave).
  • H2O (DEPC treated; as described above)
  • 10% PFA (Paraformaldehyde)
  • 20% Triton X-100 (in H2O)
  • 20% Tween-20 (in H2O)
  • 20% SDS (Sodium dodecyl sulfate)
  • 10% CHAPS
  • Tris-HCl 1M pH9.5
  • Tris-HCl 1M pH8
  • Tris-HCl 1M pH7.5
  • MgCl 1M
  • NaCl 5M
  • 10N NaOH
  • LiCl 4M (DEPC treated; as described above)
  • EDTA 200mM pH8
  • TEA 1M pH8 (Triethanolamine)
  • PIPES 1M pH8
  • Proteinase K (10mg/ml; store at −20°C)
  • 50% Sucrose (sterile filter)
  • 50x Denhardt’s (commercially available)
  • 50% Dextran Sulfate (commercially available)
  • 20x SSC
    In 800ml distilled water add the following reagents:
    • 175g of NaCl
    • 88.2g of sodium citrate
    • Adjust pH to 7 with 1M HCl
    • Adjust volume to 1L, confirm pH, and then autoclave.
  • 10x MAB
    In 800ml distilled water add the following reagents:
    • 116.1g of Maleic acid
    • Adjust pH to 7.5 with 10N NaOH (until pH is adjusted Malic acid will not enter well into solution)
    • Add 30ml of 5M NaCL
    • Adjust volume to 1L, and confirm pH.
  • HB (hybridization buffer) for whole mount in situ:
    In a 50ml conical falcon tube add the following reagents:
    • 3.25ml of 20x SSC
    • 1.25ml of 200mM EDTA pH8
    • 17.5ml of H20 (nuclease free water)
    • 2.5mg of Yeast tRNA
    • 5mg of Heparin
    • Vortex to dissolve
    • 2.5ml of CHAPS (10%)
    • 500μl of Tween-20 (20%)
    • 25ml of Formamide
    • Mix gently and heat to 65°C if necessary to dissolve tRNA and Heparin
    • Store at −20°C and heat to 65°C and mix well before each use
  • HB (hybridization buffer) for section in situ:
    In a 50ml conical falcon tube add the following reagents:
    • 500μl of Tris-HCl 1M pH7.5
    • 6ml of 5M NaCl
    • 250μl of 200mM EDTA pH8
    • 1ml of Denhardt’s 50x
    • 625μl of 20% SDS
    • 10ml of Dextran sulfate from 50% stock
    • 1ml of H20 (nuclease free water)
    • 10mg of Yeast tRNA
    • Mix gently to dissolve the tRNA
    • 25ml of Formamide
    • Mix gently and heat to 65°C if necessary to dissolve tRNA
    • Store at −20°C and heat to 65°C and mix well before each use


Background Information

Gene therapy has long been viewed as one of the tools of modern molecular medicine to treat many human disease conditions. However, less than a decade after the first clinical trials which began in 1990, the field suffered a major setback. In 1999, an 18-year-old boy died only 4 days after receiving an injection of a therapeutic Adenovirus. His death was likely caused by a severe innate immune response to the virus. The rAAVs used today elicit a minimal immune response and are thus much safer. Compared to the first generation of rAAVs, which were based on rAAV2, the ones used today achieve sustained and efficient gene expression. This new generation of rAAVs and their successful use to treat blind dogs (Acland et al., 2001) has paved the road for the future ocular gene therapy in humans.

The retina is a thin neuronal tissue at the back of the eye, which initiates the process of vision. Three distinct nuclear layers characterize it. Each nuclear layer is composed of a subset of specialized neurons (Fig. 5) (Masland, 2001, 2011). The outer nuclear layer (ONL) harbors rod and cone photoreceptors (PR), the cells that absorb photons. Rods are 1000 times more sensitive to light than cones and function primarily in dim light, while cones are used for daylight, color, and high-acuity vision. Although humans are diurnal and mice are nocturnal, in both species rods outnumber cones 20:1 (Masland, 2001), with the exception of a small region in the human eye referred to as the fovea. The fovea is composed only of cones and is the center for high acuity vision in humans. Outside the fovea the mouse and human retina are alike. Upon absorption of a photon, PRs hyperpolarize and signal to bipolar cells, which reside in the inner nuclear layer (INL). Bipolar cells connect to ganglion cells in the third nuclear layer, which send their axons through the optic nerve to the visual centers of the brain (Masland, 2001). In addition to bipolar cells, the INL is also populated by amacrine and horizontal cells (Masland, 2001), which modulate the signal, and Muller Glia cells, which are the only non-neuronal cell type in the retina and form the retinal blood barrier. Nutrition for retinal cells is provided by the retinal vasculature and the retinal-pigmented epithelium (RPE). The RPE is in intimate contact with PR outer segments. It provides nutrients and oxygen for PRs and is involved in the visual cycle (Parker and Crouch, 2010; Wang and Kefalov, 2011).

Figure 5
Retinal morphology. Cross section stained with H&E showing cartoons of different retinal cell types. Cells located in the inner nuclear layer (INL) and ganglion cell layer (GCL) are shown in white, while cones and rods located in the outer nuclear ...

Retinal degeneration is a major cause for blindness in the industrialized world. The degeneration affects either ganglion cells or PRs. Loss of ganglion cells results in glaucoma while loss of PRs is associated with a variety of retinal degenerative diseases such as dry and wet age-related macular degeneration, diabetic retinopathy, Retinitis Pigmentosa (RP), Leber’s congenital amaurosis (LCA), etc. Retinitis Pigmentosa and LCA are inherited retinal degenerative diseases. In such cases, rAAV mediated gene therapy entails either the replacement of a nonfunctional gene or the knockdown of a dominant allele. In contrast, age-related macular degeneration, diabetic retinopathy, and glaucoma are caused by a combination of environmental and genetic factors. rAAV mediated gene therapy is still possible for these diseases however, it requires an understanding of the molecular mechanisms that lead to the disease pathology. For example, in wet age-related macular degeneration and diabetic retinopathy, neovascularization of either the choroidal or the retinal vasculature, respectively, causes leakage of fluid into the retinal proper, which then results in PR death. This neovascularization is stimulated by the vascular endothelium growth factor (VEGF) and overexpressing the soluble form of the VEGF receptor-1 (sFLT-1) reduces the incidence of new blood vessel formation (Lai et al., 2005). Retinal gene therapy can thus be applied to a wide range of genetic and non-genetic eye diseases. Recently a new treatment strategy for PR degenerative diseases has emerged, referred to as optogenetics. This strategy uses the endogenous remaining retinal circuit after PRs have died to reactivate vision. rAAVs are engineered to overexpress light sensitive ion channels such as channel rhodopsin2 in bipolar cells and/or ganglion cells. Such an approach is independent of the initial insult that resulted in PR death and can thus in principle be applied to almost all PR degenerative diseases. Finally, the most successful rAAV mediated gene therapy in the eye of humans thus far targeted the RPE cells. LCA-2 is an early onset disease caused by a mutation in the RPE protein RPE65 (Gu et al., 1997; Marlhens et al., 1997). Delivery of the RPE65 gene by rAAV to the RPE of individuals suffering from LCA-2 has restored vision in blind people (Bainbridge et al., 2008; Maguire et al., 2008), giving hope to many others suffering from vision loss. Since every retinal cell type, including the RPE, is a potential target for gene therapy, a large arsenal of rAAVs capable to infect all different cell types is needed. Cell tropism of rAAV serotypes in the eye is only known for the most commonly used serotypes (Stieger et al., 2011). Knowing the tropism is particularly important since serotypes that preferentially infect only a subset of cells can reduce unwanted side effects. In combination with cell type specific promoters or microRNA regulation (Xie et al., 2011), such rAAVs can potentially restrict transgene expression to a specific cell type.

The procedure for subretinal delivery of viruses in rodents was initially described by the Cepko laboratory (Price et al., 1987) using a replication incompetent retrovirus for lineage tracing. The same laboratory also published the DNA transduction technique of retinal cells in newborn rodents by electroporation (Matsuda and Cepko, 2004) and the embryonic transduction of retinal cells (Punzo and Cepko, 2008; Turner et al., 1990). Since then, many laboratories have used these procedures successfully and modified them according to their needs.

Critical Parameters & Troubleshooting

Viral injections yield in general, successful infections even for beginners. The simple act of inserting a needle into the eye and injecting virus will result in infection. The amount of infected target cells will increase with experience. The most important variable regarding viral injections is the viral titer and the infectivity of the viral preparation. rAAV titers are generally determined by genome copies. While this number reflects the actual amount of virus particles, the number of infectious particle can be quite different and as much as 100–1000 times lower. If little infection is seen with a high titer virus, then most likely the infectivity of the viral preparation is low. A rough assessment of infectivity can be performed in cell culture by adding 5μl of the viral preparation to one well of a 6-well culture plate with HEK239 cells. This is only possible if a) the viral cassette uses a broad expressing promoter such as CMV, b) if the cassette overexpresses an easy detectable marker such as GFP, and c) if the serotype used is capable of infecting HEK293 cells. In such a case, if 2 days post infection many cells are GFP positive, the infectivity of the viral preparation should be sufficient to successfully transduce many retinal cells. However, not all rAAV serotypes will lead to such a fast expression. This method is not meant to replace the standard quantification method. It relies on the fact that super-infection of cells with more than 100 virus particles per cell will lead to a fast expression. While a high enough viral titer is usually the most important concern regarding expression in a tissue, a too high titer can lead to unwanted effects in the retina. For example, we have observed that injections with a high titer virus 5 x1013 preparation, which shows also good infectivity, can lead to PR death. However, it remains unclear if this is solely due to a high number of virus particles per PR cell. If PR death occurs due to a high titer, diluting the virus or re-purifying it over a CsCl gradient or specialized exchange columns may help mitigate the problem.

Electroporation of plasmid DNA tends to be more complicated than viral injections. Simply inserting a needle into the eye and injecting DNA will not lead to GFP positive cells. Although the procedure is technically the same, the biggest hurdle is targeting the subretinal space properly, delivering enough DNA into that space and placing the electrode pads adequately. It may take a couple of mouse litters to master the technique. When dissecting the retina, if GFP is seen only on top of the PR outer segments but not in PRs itself, it suggests that the DNA was targeted correctly to the subretinal space but not electroporated efficiently. Macrophages that enter the retina and clean up the remaining plasmid DNA will be GFP positive due to the plasmid they took up. In such a case, either the electroporator is not delivering enough current or the electrode pads were not placed properly. The plus pole of the tweezer electrodes tends to oxidize over time, which will result in a reduction of the electric field. The electrode pads need to be kept clean and the tweezer electrode needs to be replaced every so often. The most critical parameters for this procedure are the quality and concentration of the DNA, the proper targeting of the subretinal space, the electroporator, and the tweezer electrodes. We recommend practicing subretinal injections into newborn mice with the CD1 mouse strain. CD1 mice have large litters and are albino, which helps initially to visualize the spread of the injection solution.

In situ hybridizations on retinal whole mount or sections can be quite challenging. In general, the quality of the probe and the target sequence that was chosen for hybridization are the most critical parameters. If the whole mount in situ hybridization is not working, we recommend testing the probe on sections. After resuspending the probe with H2O, we recommend adding 80μl of the section in situ HB. Diluting the probe afterwards in the whole mount HB for the actual hybridization does not affect the whole mount in situ hybridization, while the probe can still be used for section in situ hybridizations. This allows testing the probe on sections if whole mounts are not working. If the probe does not work on sections we recommend to either resynthesize the probe, or to choose a different target region of the gene. As a positive control, we recommend to start with a gene that is expressed at high levels such as one of the PR specific opsins. If there is too much background, increasing the number of washes and the wash temperature may mitigate the problem. Incubation with the antibody alone will determine if the background stems primarily from the antibody being trapped in the tissue or from unspecific trapping of the probe. A sense probe can be used as a control for the hybridization. However, many genes have coding sequence of another gene running in the antisense direction. Thus it is important to determine on the UCSC genome browser if there is a gene running in the opposite direction prior to designing and synthesizing a sense control probe. We also recommend using the UCSC genome browser to determine if there are alternative splice isoforms of your gene of interest. In such a case designing a probe against the most common region may be a good starting point.

Antibody staining tends to be straightforward when it works and difficult to troubleshoot when it does not work. The antibody and the epitope that is recognized by the antibody are the most important factors, which are unfortunately difficult to control. Here are some tips if the staining does not work. In general, more antibodies tend to work better on cryo-sections than on paraffin section. However, there are exceptions and some antibodies work better on paraffin sections. If your antibody does not work, adding SDS at a final concentration of 0.01%–0.03% in PBTB may help. Alternatively an antigen retrieval protocol can help. There are many different protocols for antigen retrieval and it is difficult to predict which ones works best for a specific antibody. If the signal is weak by immunofluorescence, then immunocytochemistry may yield a better result. In such a case we would recommend to use a secondary antibody coupled to horseradish peroxidase. If such an approach is used the tissue needs to be pretreated with H2O2 to inactivate the endogenous peroxidase in order to reduce background.

Anticipated Results

Subretinal injections at postnatal day 0 take some time to master. Viral injections tend to yield sooner positive results than electroporations, as injecting a virus into the eye will result in infection even if only few of the desired target cells are hit. Most people tend to have positive transduction with both methods after injecting 3 litters. With some practice, 50–70% of electroporated retinas will be successfully transduced. In such cases, the retinal surface area that is positively transduced ranges anywhere between 5%–50%, with 25% being the norm for most people (Fig. 6). Viral injections yield similar results however, it is easier to transduce a larger area of the retina. With some experience, it is possible to transduce even the entire retinal surface area with a single injection (Fig. 6). This may take quite some practice and even with a lot of experience only 10–20% of virally infected retinas will show infection across the entire retinal surface area.

Figure 6
Expected results after viral infection or electroporation of plasmid DNA at postnatal day 0. (A–D, K, L) Show retinal whole mounts and (E–J) show retinal cross sections of GFP transduced cells. (A–H) Show transduction of retinal ...

The time point of electroporation determines the cell types that can be transduced efficiently. To broaden the range of cells types by electroporation, the procedure needs to be performed at embryonic time points. Such a procedure has been described previously (Punzo and Cepko, 2008). In contrast, viral injections with rAAVs can target all retinal cell types at any age. While the cell type that is efficiently infected is dependent on the serotype, the viral titer and the infectivity of the viral preparation can influence the perceived tropism. Additionally, the route of delivery may also influence the cell types that are preferentially infected. Figure 6 shows examples of subretinal injections with a rAAV(2/5) that carries nuclear GFP. At a low titer and low infectivity, preferentially cone PRs and some rods are infected. With increasing titer and infectivity, INL cells appear infected as well. It is thus important to determine if the tropism of a virus has been tested to its maximum potential before concluding which cells can be infected with a specific serotype. Performing both subretinal and intravitreal injections with a high titer virus that shows good infectivity should yield a comprehensive picture of the cell tropism of a specific rAAV serotype. Of note, the tropism observed in the mouse retina with a specific serotype may change when performing injections with the same viral preparation in a different organism.

Immunofluorescence or in situ hybridization analyses depend on the antibody and probe used. A compilation of immunofluorescence and in situ hybridization data is shown in Figure 7.

Figure 7
Expected results for immunofluorescence and in situ hybridization analyses. Compilation of immunofluorescence stanings (A, C, C′) or in situ hybridizations (B, D, D′) on retinal whole mounts (A, B) or retinal cross-sections (C–D′) ...

Time Considerations

Viral injections or electroporations at postnatal day 0 take roughly 1½ to 2 hours per litter depending on your experience. This includes the 1 hour lag time, in which pups are injected with Buprenorphine. The individual injection of a pup takes only 2–3 minutes. Viral injections into adults take a similar amount of time. Transduction of retinal cells either by rAAV or by electroporation differs mainly in the time it takes for gene expression to occur. Gene expression upon electroporation can be detected as early as 2 days post electroporation and stays stable for several months. In contrast, gene expression from rAAV vectors may take 1–3 weeks, however expression tends to persist for years. Two variables determine the speed of expression for rAAVs. One is the viral serotype and the other is the multiplicity of infection (MOI), which depends on the amount of virus that is injected in a particular area and the actual infectious titer of the virus. In general, when more than one virus particle enters a cell, expression tends to start earlier. In our hands, the rAAV(2/5) shows robust GFP expression within 10 days post infection. Preparation of the virus is not discussed here but takes generally 1–2 weeks including, cell line expansion, purification and virus titering.

Retinal analysis by immunofluorescence or in situ hybridization may take 2 days to 2 weeks depending on the protocol and if sections or whole mounts are processed. The specific time windows for the different procedures are indicated in the individual protocols. When performing an antibody staining, we recommend to incubate with the primary antibody over night until you have determined that the antibody works well, at which point that step can be shortened to 2 hours.

Supplementary Material

Video S1

Movie 1: Subretinal space injections into newborn mouse pups (Basic Protocol 1 & 2).

Video S2

Movie 2: Dissections and tissue processing for whole mount and section analysis (Basic Protocols 5–6, Support Protocols 1–2).


This work was supported in part by grants from the National Institutes of Health to G.G (UL1RR031982, 2P01 HL059407, 1P01AI100263-01, and 2R01NS076991-01).

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