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Murine models are ideal for studying cochlear gene transfer as many hearing loss-related mutations have been discovered and mapped within the mouse genome. However, due to its small size and delicate nature, the membranous labyrinth of the mouse is a challenging target for delivery of viral vectors. To minimize injection trauma, we developed a procedure for the controlled release of adeno-associated viruses (AAV) into the scala media of adult mice. This procedure poses minimal risk of injury to structures of the cochlea and middle ear and allows for near-complete preservation of low and middle frequency hearing. In the present study, transduction efficiency and cellular specificity of AAV vectors (serotypes 1, 2, 5, 6, and 8) were investigated in normal and drug-deafened ears. Using the cytomegalovirus (CMV) promoter to drive gene expression, a variety of cell types were transduced successfully, including sensory hair cells and supporting cells, as well as cells in the auditory nerve and spiral ligament. Among all five serotypes, inner hair cells (IHCs) were the most effectively transduced cochlear cell type. All five serotypes of AAV vectors transduced cells of the auditory nerve, though serotype 8 was the most efficient vector for transduction. Our findings indicate that efficient AAV inoculation (via the scala media) can be performed in adult mouse ears, with hearing preservation a realistic goal. The procedure we describe may also have applications for intra-endolymphatic drug delivery in many mouse models of human deafness.
Hearing loss is largely due to degeneration and/or abnormality of sensory hair cells, spiral ganglion neurons, or specific non-sensory cells, such as fibrocytes in the spiral ligament and glial cells in the auditory nerve. In mammalian vertebrates, sensory hair cells and spiral ganglion neurons are unable to regenerate from surviving cell populations. There is evidence that certain non-sensory cells, such as cochlear fibrocytes, are able to repopulate themselves after injury; however, this regenerative ability declines with age.1,2,3 Non-mammalian vertebrates can replace lost hair cells through direct and indirect trans-differentiation of supporting cells (see review from Brignull et al4 and Groves5). Raphael’s group developed an adult guinea pig model of cochlear cell regeneration using adenoviral vectors to drive expression of the Atoh1 transcription factor in the drug-damaged organ of Corti, leading to a remarkable recovery of auditory function and regeneration or repair of drug-damaged hair cells.6 Staecker et al7 found that Atoh1 gene transfer using adenovectors resulted in the recovery of vestibular function and regeneration of macular hair cells. Gubbels et al8 reported that Atoh1 gene transfer is able to induce functional hair cells in the mouse cochlea using in utero electroporation of plasmid DNA. These and other gene transfer studies in the inner ear demonstrate the feasibility of genetic manipulation for cell replacement based on inducing regeneration and/or trans-differentiation of endogenous cochlear cells. In the past 15 years, numerous studies have investigated the functional and structural effects of gene transfer into inner ear tissues.9–43 These reports have provided important information on the advantages and disadvantages of different surgical approaches, gene delivery methods and viral vectors.
AAV has limited immunogenicity and toxicity and has been shown in previous studies to transduce cochlear hair cells and supporting cells (SCs).9,14,24,25,27,32,37,42,43 To date, the majority of these studies have used primary mouse cochlear explants, perinatal mouse cochleas, or adult guinea pig cochleas as models to investigate the transduction efficiency of AAV vectors. Adult mice present an ideal model for cochlear gene transfer because their genetic information is accessible and many hearing loss-related mutations have been discovered. Via a scala tympani approach in normal adult mice, Liu et al25 successfully transduced inner hair cells (IHCs) and other cochlear cell types including cells in the spiral ligament and the spiral ganglion using AAV vectors. However, no outer hair cells (OHCs) and only a few SCs were seen for all 8 AAV serotypes examined.
Further improvement in infection efficiency of OHCs and SCs in the mouse model is desirable, especially since the therapeutic delivery of genes to specific inner ear cells may be required to treat different genetic causes of deafness. In the present study, we examined the efficiency of AAV infectivity of IHCs, OHCs, SCs, auditory nerve fibers, and cells of the lateral wall under various conditions. More specifically, transduction efficiency was assessed in the presence or absence of drug-induced hair cell loss as well as under various conditions (serotypes, volume, rate) of virus delivery into the scala media. The delivery of fluid reagents into the scala media is particularly challenging in the mouse due to its small volume and the need to add an appropriate number of transducing viral particles to ensure efficient infection. To reduce the surgical trauma to cochlear cells associated with bulk fluid injections, we employed a newly-developed nanoliter-level fluid delivery system (Nanoliter Microinjection System, WPI, Sarasota, FL). Using this system, injection trauma can be minimized by optimizing the delivered volume while reducing the speed of injection. A variety of cell types in normal and deafened inner ears were successfully transduced including sensory cells in the organ of Corti, cells in the auditory nerve and the spiral ligament of the cochlear lateral wall, as well as cells in the vestibular end organs.
To optimize the use of AAV vectors for therapeutic treatment of hair cell loss, it was important to compare virus-mediated gene transduction in normal and damaged ears of adult mice. For deafening, we employed a recently published method of rapid hair cell degeneration that used a combination of a single dose of aminoglycoside following by a single dose of a loop diuretic delivered intraperitoneally into adult CBA/CaJ mice.45,49,50 Auditory brainstem responses (ABRs) were measured at 3 days, 7 days, 1 month, or 3–5 months after treatment (Fig. 1a). A significant ABR threshold shift (about 40–70 dB SPL across all tested frequencies) was revealed as early as 3 days after treatment. No signs of functional recovery were seen at 3–5 months after treatment, which was the longest recovery period examined.
Drug-treated mice were examined at 7 days, 1 month and 3 months to evaluate the extent of hair cell destruction. A loss of OHCs was seen in the entire basal turn and most of the apical turn of the examined ears (n=19) (Figs. 1b, 1c, 1d). Only a small number of OHCs remained in the extreme apex. The majority of IHCs were intact but a scattered loss of IHCs was observed in the basal turns of treated mice. These results correspond with previous studies.45,49,50 Most importantly, relatively normal architecture of supporting cells was seen 1–3 months after treatment (Figs. 1d, 1e). Therefore, this deafened model is ideal for testing the transduction efficiency of AAV viruses in surviving supporting cells in the context of nearly complete OHC loss.
The organ of Corti is a delicate epithelium located on the basilar membrane, an acellular structure suspended across a fluid-filled space deep within the temporal bone. Furthermore, delivery of virus directly to the apical surfaces of the sensory epithelium requires a breach of the scala media, a tubular compartment filled with potassium-rich endolymphatic fluid. Excessive leakage of endolymph at the site of injection will degrade the transepithelial endocochlear potential that is an essential driving force for effective mechanosensory transduction. To better preserve the integrity of both the basilar membrane and the organ of Corti, we established a surgical approach to deliver AAV vectors into the scala media through the lateral wall (Fig. 2a, 2b). The bulla was surgically exposed and prepared to permit access to the basal turn of the mouse ear. This approach allows direct visualization of the bony lateral wall of the scala media and the richly vascularized spiral ligament of the stria vascularis that lies just beneath. We were able to minimize injection trauma by optimizing the delivered volume and speed of injection through glass micropipettes using a Nanoliter Microinjection System. Although injection trauma was minimal, 7% of animal subjects showed significant leakage of endolymph at the injection site and thus were not included in the study.
Physiological assessments were used to evaluate whether the surgical procedure itself was potentially damaging to the inner ear. Figures 2c and 2d show that no significant ABR threshold shifts were seen at 4 to 22.6 kHz in a normal mouse 7 days after AAV inoculation. However, a 30–40 dB SPL shift in the ABR threshold did appear at higher frequencies (32, 40 and 45.2 kHz), suggesting a cochlear injury at the extreme basal turn adjacent to the typical injection site. In contrast, deafened mice, which were already missing most OHCs and some IHCs in the basal turn (data not shown), demonstrated no additional high-frequency ABR threshold shifts in response to virus inoculation (Fig. 2e)
Five types of AAV-GFP vectors (serotypes 1, 2, 5, 6, and 8) were delivered to normal or deafened adult mice. Although histological analysis revealed that many cell types could express GFP following AAV inoculation, IHCs were by far the most commonly transfected (Figs. 3 and and4).4). For a semi-quantitative evaluation of the relative transduction efficiency of AAVs under different experimental conditions, we counted IHC expression in two different ways. First, the number of mice showing any GFP expression in IHCs was counted relative to the total number of mice inoculated. Second, the number of mice with more than 100 GFP+ IHCs was counted and compared to the total number of mice inoculated. Both of these measures can be found in Table 1 for the full set of inoculated animals.
The epithelium of a deafened animal is likely to differ from that of a normal animal, and these changes might influence the number or types of cells infected under the two conditions. Furthermore, an acutely deafened organ of Corti may not necessarily be infected with similar efficiency to that which is chronically deafened. To examine the effects of post-deafening recovery time, AAV was delivered into the scala media of the deafened cochlea following either short term (3–7 days, acutely-deafened) or long term (1–6 month, chronically-deafened) recovery periods after kanamycin and furosemide treatment. There is a trend showing that AAV transduction is more effective in acutely-deafened mice than chronically-deafened mice (Table 1). About 87% (32/37) of the acutely-deafened mice expressed GFP protein, as compared to 68% (17/25) of the chronically-deafened mice. This compares to 88% (35/39) of normal animals that were transduced with GFP. However, these numbers do not take into account the rather large differences in infection efficiency across viral serotypes.
For each AAV serotype, we initially examined 3–4 animals per group. The overall transduction efficiency showed that GFP transduction was best achieved with serotypes 2 and 8, secondarily with serotype 6 and lastly with serotypes 1 and 5. The initial results led us to focus on serotypes 2 and 8 to further examine the effect of survival time after virus inoculation on the transduction efficiency (Tables 1 and and2).2). The transduction patterns of AAV vectors in sensory cells of normal and deafened ears were also characterized using vector serotypes 2 and 8 (Figs. 5 and and66).
The effect of recovery period on transduction efficiency of AAV was compared for serotypes 2 and 8 at seven days and one month after surgery (Table 2). The percentage of inoculated mice showing any GFP expression in IHCs ranges from 88–100%, with no clear advantage of one serotype over the other using this measure. The percentage of inoculated mice with >100 GFP+ IHCs shows a large range from 41–64%, with the highest percentage seen at one month for serotype 2. Qualitatively, there was no clear trend towards a larger number of GFP-expressing cells per ear with increased recovery time.
For all five AAV serotypes, IHCs were the most effectively transduced cell type in the organ of Corti. Robust expression of GFPin IHCs was seen in both normal and deafened mice, especially for serotypes 2 and 8 (Figs. 3 and and4).4). Figure 4a shows a deafened mouse with near complete IHC transduction 7 days after serotype 2 virus inoculation. OHCs were also shown to express GFP in the normal ear (Fig. 3). However, robust GFP+ OHCs were only seen in the apical turns of four normal mice (out of 25 mice that were inoculated with serotype 2 and 8 viruses).
A small number of GFP+ SCs were seen in normal and deafened mice for serotypes 2, 6 and 8 (Figs. 3d, 3e, ,4e,4e, 5a–f). Sox2 was used as a marker to identify SCs. A previous study has indicated that Sox2 can be expressed in the nuclei of all cochlear SC subtypes including Deiters’, Hensen’s, inner and outer pillars, inner phalangeal, and border cells.45 Figure 5 reveals that some of the GFP+ cells were co-labeled with Sox2. However, there are some GFP+ cells in the SC region (outer pillar cell region) of the deafened mice that were not stained for Sox2 (Figure 5). It is possible that these GFP+/Sox2− cells are fibroblasts or macrophages that appeared in response to the loss of cochlear OHCs.
We were also interested in whether scala media inoculation could yield GFP gene transduction in vestibular organs. The examination of frozen sections of the inner ears of normal and deafened mice revealed GFP expression in both a subset of HCs and SCs of the sensory epithelia of vestibular organs including the macula and ampulla (Figure 6). Oesterle et al45 reported that Sox2 protein was expressed in SCs and type II HCs of both the striolar and extra-striolar regions. We found that a majority of the GFP+ HCs were co-labeled with Sox2, suggesting these cells are type II HCs (Fig. 6c–h). GFP expression patterns were similar in normal and deafened ears. A few GFP+ cells were also seen within the stromal region of the vestibular organs (data not shown).
Scala media inoculation can also produce efficient GFP transduction in cells within Rosenthal’s canal, although this region was not specifically targeted. Based upon their morphological features, GFP+ cells within Rosenthal’s canal are mostly glial cells (Fig. 7). The proportion of normal and deafened animals showing GFP expression as a function of viral serotype is shown in Table 3. Transduction was best achieved with serotype 8. GFP+ auditory nerve cells were seen in 60% of inoculated mice for serotype 8, 43% for serotype 1, 42% for serotype 3, 38% for serotype 6 and 22% for serotype 5. Overall, we observed a trend towards higher transduction efficiency in deafened ears as compared to normal ears.
GFP+ cells were also seen in the spiral ligament of normal and deafened mice that received serotypes 2 and 8 viruses (Fig. 8). In most cases, GFP was expressed in the outer sulcus epithelial cells as well as type II and IV fibrocytes of the spiral ligament. GFP+ fibrocytes were mostly located in the basal turns of both normal and deafened mice. No GFP+ cells were seen in the stria vascularis of the specimens examined.
We established a scala media delivery procedure that is able to target a variety of cell types in normal and deafened adult mouse ears. The cell types transduced include sensory HCs and SCs in the auditory and vestibular organs as well as glial cells in the auditory nerve and fibrocytes of the spiral ligament. Most importantly, this procedure minimizes surgical trauma and preserves hearing, especially at low and middle frequencies. Compared to previous studies that have used a scala media approach, there were two major improvements in our procedure which played an important role in reducing cochlear injury and preserving hearing. First, a nanoliter-level fluid delivery system was used to control the precisely delivered volume and speed of virus injection. This system allows us to deliver controlled amounts of fluid as small as 4.6 nl. In the adult mouse, the volume of the endolymphatic space in the scala media was reported to be 0.19 μl.51 In our procedure, multiple injections were performed and the fluid volume delivered into the scala media was approximately 46 nl per injection. There was a one-minute recovery period after each injection, allowing the host cochlear cells to adjust to the change in endolymphatic pressure. A slow-infusion technique with a minor disruption to the cochlear fluid environment during virus inoculation is critical for delivering virus into the endolymph compartment with minimal tissue trauma. A similar strategy was used for fluid delivery into the perilymph compartment of the adult mouse cochlea.52, 53 Infusion of an artificial perilymph solution into the scala tympani at a rate of 16–32 nl/min resulted in good preservation of hearing function.
Second, our surgical method was adopted from a standard non-invasive procedure of endocochlear potential measurement in adult mice.54 The bony lateral wall of the scala media was thinned by a dental drill but the membranous lateral wall was left intact. The endolymph-filled cavity was sealed completely without an opening hole during the surgical procedure. This preparation allows the sharp tip of a glass micropipette to penetrate the membranous lateral wall of the scala media without endolymph leakage. By the end of the virus delivery procedure, the penetrated site was able to completely close after the micropipette was removed.
A great advantage of the scala media approach is the highly efficient transduction of cochlear sensory cells.6, 21,24 The improved delivery methods developed in this study largely reduced cochlear injury and preserved low and middle frequency hearing well following scala media virus inoculation. This delivery procedure may potentially be used for intra-endolymphatic drug delivery in various mouse models of human deafness.
AAV vectors have often been used for gene delivery into the central nervous system as they express genes in post-mitotic neuronal cells for long periods with minimal to no immunogenicity and toxicity (see review from Hester et al.55). An interesting finding in this study is that IHCs, the post-mitotic sensory cells of the inner ear, were most effectively transduced by AAV vectors. This finding is of value for further molecular and physiological studies of sensory hair cells and for planning gene therapy strategies when deafness is caused by a gene deficit in sensory hair cells that does not affect their survival but perturbs their function.
GFP+ IHCs were seen in ears inoculated with all five serotypes of AAV vectors (serotypes 1, 2, 5, 6 and 8). In particular, AAV2 and AAV8 were the most efficient vectors for IHC transduction. The divergence of cellular tropism observed in AAV serotype vectors may contribute to the differential characteristics of the vector-cell interaction. It has been known that transduction of cells with AAV2 is mediated by heparan sulphate proteoglycan receptor, fibroblast growth factor receptor 1, aVb5.56–58 Like AAV2, AAV8 also needs a laminin receptor for cell entry.58 In contrast to those two serotypes, AAV1, AAV5 and AAV6 do not need to bind a receptor for cell entry.58,59 Interestingly, our study found that a majority of the GFP+ HCs in vestibular organs are type II HCs (Sox2+) suggesting that type II vestibular HCs may share a common AAV8 receptor (such as a laminin receptor) with cochlear IHCs.
The use of the CMV promoter may also contribute to the high transduction pattern seen in cochlear IHCs. A previous study showed that replication-defective (E1−, E3−, pol−) adenovirus vectors containing CMV-driven LacZ can transduce about 99% of the IHCs in the basal turn and 90% of the IHCs in the apical turn of adult guinea pigs.23 Liu et al.25 reported that AAV serotype 3 vectors containing chicken β-actin promoter can transduce adult mouse IHCs with high efficiency. It will be interesting to examine whether AAV3 with the CMV promoter can produce a similar gene transduction pattern.
We expected the infection efficiency of SCs to be enhanced by drug treatment, since the absence of HCs should allow the apical surfaces of SCs to expand and thus present a larger apical surface area for binding of viral particles. While GFP+ SCs were seen in both normal and deafened mouse ears inoculated with AAV2, AAV6 and AAV8, the transduction efficiency of SCs was low in both conditions. Furthermore, there was no difference in SC infectivity comparing short-term (acutely deafened ears) and long-term (chronically deafened ears) recovery periods after kanamycin and furosemide treatment. Thus, OHC loss does not appear to increase gene transduction efficiency into SCs. These results suggest that either receptor availability is limiting, or perhaps the CMV promoter is not an ideal promoter for expression in adult SCs. Previous studies reported robust transgene expression in cochlear SCs of several models including mouse cochlear explants, postnatal mouse ear and adult guinea pig ear. Stone et al26 found that the GFAP promoter with AAV serotypes 1 and 2 vectors can result in strong transgene expression in SCs of the cultured organ of Corti taken from E13 or P0-1 mice. Iizuka et al32 observed an efficient transgenesis in Deiter’s cells, IHCs, and lateral wall cells after AAV serotype 5 vectors with a CMV promoter were delivered into P0 mouse ears. Shibata et al37 observed that bovine AAV with a β-actin promoter transduced SCs efficiently in both normal and deafened adult guinea pig cochleas. Our data and these previously reported data demonstrate that the SC transduction efficiency is likely dependent upon the choice of promoters, virus serotypes, in vitro or in vivo inoculation, animal species, and animal age. Further investigation into the proper promoter may be necessary to increase the efficiency of targeting SCs in the adult mouse ear with AAV vectors.
Another finding of this study is that scala media delivery of vectors can transduce GFP gene expression in fibrocytes of the spiral ligament of the cochlear lateral wall. Non-sensory cells such as fibrocytes, along with sensory hair cells and neurons, form the highly complex microarchitecture of the inner ear. Non-sensory cells not only provide an essential environment for the health of sensory hair cells and neurons, but they also play an important role in the maintenance of normal auditory functions.60,61,62 The ability to transfer genes into the cells of the cochlear lateral wall is important for better understanding the function of cochlear fibrocytes.
This study also found that the scala media approach can transduce the GFP gene in glial cells of the auditory nerve. We compared the transduction efficiency of 5 serotypes of AAV vectors in the auditory nerve. Serotype 8 was found to be optimal for gene transfer in the auditory nerve under these experimental conditions. In addition, there is a trend toward an increased transduction efficiency into the nerve in deafened mice compared to normal mice. Local transduction of cochlear glial cells with a secreted protein, e.g., a growth factor, could prove beneficial for preventing the degeneration of spiral ganglion neurons after HC loss or with age. A recent study showed that transgene expression of the neurotrophin BDNF in cochleas with prior hair cell degeneration led to a robust re-growth of auditory nerve fibers into the sensory epithelium.63
Purified AAV-GFP vectors of serotypes 1, 2, 5, 6, 8 were purchased from the Harvard Gene Therapy Initiative (Harvard Medical School, Boston, MA). Viral stocks were generated by tripartite transfection (AAV-RE/CAP expression plasmid, adeno-miniplasmid pHGTI-Adeno1 and AAV transfer vector plasmid pAAV-GF) into HEK293 cells. GFP expression in these vectors is driven by the CMV promoter. Serotype 2 particles were purified by heparin column chromatography. All other serotypes were purified by iodixanol density gradient followed by Q sepharose column chromatography. Purified vector particles were dialyzed extensively against PBS, concentrated by Amicon spin columns, and titered by dot blot hybridization. Concentrated virus titers for serotypes 1, 2, 5, 6 and 8 were 6.7 × 1012, 3.2 × 1012, 5.3 × 1012, 7.5 × 1012 and 1 × 1013 genome copies per ml, respectively. With total injection volumes of ~322 nl per cochlea (see below), approximately 1–3 × 109 genomes were delivered to each ear. Previous work has shown that genome copies per ml, calculated using quantitative polymerase chain reaction, can overestimate by ~5 log units the number of AAV transducing units/ml, based on LacZ or eGFP transgene expression following limiting dilutions on cultured cells.44 Thus, we estimate that we are delivering approximately 10,000–30,000 transducing units to each ear.
A colony of adult CBA/CaJ mice was established with original breeding pairs purchased from the Jackson Laboratory (Bar Harbor, ME) and bred in-house in a low-noise environment at the Animal Research Facility of the Medical University of South Carolina (MUSC). About 120 mice of both genders, aged 2 to 12 months and weighing 16–35g, were used in the study. All aspects of the animal research were conducted in accordance with the guidelines of the Institutional Animal Care and Use Committee of MUSC. Prior to data acquisition, mice were examined for signs of external ear canal and middle ear obstruction. Mice with any symptoms of ear infection were excluded from the study.
Cochlear lesions were induced according to a previous description by Oesterle et al.45 Kanamycin sulfate (Sigma-Aldritch, St. Louis, MO) was dissolved in physiological saline to a concentration of 45 mg/ml. Mice were deafened with a single dose of kanamycin (1,000 mg/kg, subcutaneous) followed 45 min later by a single intraperitoneal injection of furosemide (400 mg/kg) (American Regent Laboratories, INC).
Mice were anesthetized by an intraperitoneal injection of xylazine (20mg/kg) and ketamine (100mg/kg) and placed in a head holder in a sound-isolation room. Young adult CBA/CaJ mice underwent physiological measurements before and after the kanamycin and furosemide treatment as well as before and after virus delivery. Auditory brainstem responses were recorded via customized needle electrodes inserted at the vertex (+) and test-side mastoid (−), with a ground in the control-side leg. The acoustic stimuli were generated using Tucker Davis Technologies equipment III (Tucker-Davis Technologies, Gainsville, FL, USA) and a SigGen software package. ABRs were evoked at half octave frequencies from 4 to 45 kHz with 5 ms duration tone pips with cos2 rise/fall times of 0.5 ms delivered at 31/s. At each sound level, 300–500 responses were averaged, using an “artifact reject” whereby response waves were discarded when peak-to-peak amplitude exceeded 50 mV. Physiological results were analyzed for individual frequencies, and then averaged for each of these frequencies from 4.0 to 40 kHz.
Normal and deafened mice were anesthetized as described above. The bulla was exposed through a post-auricular approach and a small perforation was created to expose the basal portion of the cochlea. The bony cochlear lateral wall of the scala media in the basal turn was thinned carefully using a dental drill with a micro-burr. Note that a small area (about 30–40 μm in diameter) of the bony lateral wall at the basal turn was mostly removed by the dental drill, but the membranous lateral wall was intact. Glass micropipettes (WPI, Sarasota, FL) were pulled and the tips were broken to a diameter of 15–20 μm. The Nanoliter Microinjection System (WPI, Sarasota, FL) was used to deliver controlled amounts of fluid. The injection speed was set for a rate of 46 nl/sec according to the manufacturer’s protocol (http://www.wpiinc.com/index.php/vmchk/B203XVY.html). For each mouse, a total of seven injections were completed with a one-minute recovery period after each injection. The duration of one injection was about 1 sec, delivering a total of ~322 nl into the inner ear.
For morphological observation of the deafened ears, the anesthetized animals (7 days, 1 or 3 months after kanamycin and furosemide treatment) were perfused via cardiac catheter first with 10 ml of normal saline containing 0.1 % sodium nitrite and then with 15 ml of a mixture of 4% paraformaldehyde and 2% glutaraldehyde in 0.1M phosphate buffer, pH 7.4. After removing the stapes and opening the oval and round windows, 0.5 ml of fixative was perfused gently into the scala vestibuli through the oval window. The inner ears were dissected free and immersed in fixative overnight at 4 °C. Decalcification was completed by immersion in about 50 ml of 120 mM solution of ethylenediamine tetracetic acid (EDTA) for 2–3 days. The tissues were post-fixed with 1% osmium tetroxide for 1 hour, dehydrated and embedded in Epon LX 112 resin. Semi- thin sections approximately 1 μm thick were cut and stained with toluidine blue.
For direct observation of GFP-gene transduction and immunohistochemical analysis, the anesthetized animals were perfused via a cardiac catheter first with 10 ml of normal saline containing 0.1 % sodium nitrite in normal saline and then with 15 ml of 4% paraformaldehyde in 0.1M phosphate buffer, pH 7.4. The procedures for observation of surface preparations of the mouse cochleas have been described previously.47 The cochlear sensory epithelium, modiolus, stria vascularis and spiral ligament were carefully dissected from the fixed cochlea. GFP+ cells were directly observed and counted using either a Zeiss Axio Observer fluorescent microscope or a Zeiss LSM5 Pascal confocal microscope (Carl Zeiss Inc., Jena, Germany). All virus-injected right ears (n=101) were processed for morphological and/or immunohistochemical observation and are listed in Table 1. To investigate the possibility of contralateral transduction, 38 uninjected (left) ears were randomly selected from mice in the serotypes 2 and 8 groups and were also processed for direct observation of GFP-gene transduction. However, no GFP+ cells were found in these cochleas. These uninjected ears were excluded from Tables 1, ,22 and and33.
In some cases, the cochlear sensory epithelia were processed for dual-immunostaining for GFP and Sox2. These specimens were prepared according to the method previously described.47 Frozen sections or surface preparations of cochlear tissue were blocked with 0.25% BSA and incubated overnight at 4°C with primary antibodies. The primary antisera were rabbit anti-GFP (1:200, A11120, Molecular Probes, Eugene, OR) and goat anti-Sox2 (1:200, sc17320, Santa Cruz Biotechnology, Santa Cruz, CA). Biotinylated secondary antibodies were detected with avidin D coupled to FITC (1:150) or Texas-red (1:150) (Vector, Burlingame, CA). Nuclei were counterstained with bisbenzimide or propidium iodide (PI) in specimens stained with only one primary antibody. Negative controls included omission of the primary antibody or substitution with similar dilutions of non-immune serum of the appropriate species. No regionally specific staining was detected in any of these control experiments.
Anti-GFP rabbit polyclonal and mouse monoclonal antibodies raised against GFP isolated directly from the jellyfish Aequorea victoria have been used for detection of native GFP, GFP variants and most GFP fusion proteins48. The goat polyclonal antibody for Sox2 was raised against a peptide corresponding to C-terminus aa 227–293 of human origin (manufacturer’s technical information). Western blot analysis revealed a single band at 34 kDa in human and mouse embryonic stem cell lysates.
The sections were examined either with a Zeiss Axio Observer or a Zeiss LSM5 Pascal confocal microscope (Carl Zeiss Inc., Jena, Germany). The captured images were processed using Image Pro Plus software (Media Cybernetics, MD), AxioVison 4.8 (Carl Zeiss Inc., Jena, Germany) and Zeiss LSM Image Browser Version 2,0,70 (Carl Zeiss Inc., Jena, Germany). Adobe Photoshop CS2 was employed to adjust brightness, contrast, and sharpness of images with identical setting for all panels. Alterations were not performed on images used for quantitative purposes.
Grant sponsors: National Institutes of Health; Grant number: DC00422 (H.L.); Grant number: DC07506 (H.L.); Grant number: DC00713 (B.A.S.); Grant number: DC002756 (D.M.F); American Academy of Otolaryngology-Head and Neck Surgery; Grant number: CORE 130165 (L.A.K.).
The authors thank Vinu Jyothi for her assistance in ototoxic drug exposure and ABR measurements; Manna Li, Juhong Zhu, Nancy Smythe and James Nicholson for their help with histological observations; Bradley A. Schulte and Richard A. Schmiedt for their critical comments and invaluable discussion over the course of this study.
Conflict of interest
The authors declare no conflict of interest.