Hearing Research 265 (2010) 70–76 Contents lists available at ScienceDirect Hearing Research journal homepage: www.elsevier.com/locate/heares Research paper b-Bungarotoxin application to the round window: An in vivo deafferentation model of the inner ear Björn Palmgren a,b,*, Zhe Jin c, Hongmin Ma a, Yu Jiao a, Petri Olivius a,b a Center for Hearing and Communication Research, Karolinska University Hospital, 171 76 Stockholm, Sweden Department of Clinical Neuroscience, Section of Otorhinolaryngology, Karolinska Institutet, Karolinska University Hospital, 171 76 Stockholm, Sweden c Department of Neuroscience, Uppsala University, Box 593, 751 24 Uppsala, Sweden b a r t i c l e i n f o Article history: Received 1 April 2009 Received in revised form 13 February 2010 Accepted 18 February 2010 Available online 23 February 2010 a b s t r a c t Hearing impairment can be caused by a primary lesion to the spiral ganglion neurons (SGNs) with the hair cells kept intact, for example via tumours, trauma or auditory neuropathy. To mimic these conditions in animal models various methods of inflicting damage to the inner ear have been used. However, only a few methods have a selective effect on the SGNs, which is of importance since it might be clinically more relevant to study hearing impairment with the hair cells undamaged. b-Bungarotoxin is a venom of the Taiwan banded krait, which in vitro has been shown to induce apoptosis in neurons, leaving remaining cochlear cells intact. We wanted to create an in vivo rat model of selective damage to primary auditory neurons. Under deep anaesthesia, 41 rats received b-Bungarotoxin or saline to the round window niche. At postoperative intervals between days 3 and 21 auditory brainstem response (ABR) measurement, immunohistochemistry, SGN quantification and cochlear surface preparation were performed. The results in the b-Bungarotoxin-treated ears, as compared with sham-operated ears, show significantly increased ABR thresholds at all postoperative intervals, illustrating a severe to profound hearing loss at all tested frequencies (3.5, 7, 16 and 28 kHz). Quantification of the SGNs showed no obvious reduction in neuronal numbers until 14 days postoperatively. Between days 14 and 21 a significant reduction in SGN numbers was observed. Cochlear surface preparation and immunohistochemistry showed that the hair cells were intact. Our results illustrate that in vivo application of b-Bungarotoxin to the round window niche is a feasible way of deafening rats by SGN reduction while the hair cells are kept intact. Ó 2010 Elsevier B.V. All rights reserved. 1. Introduction In recent years considerable progress has been made in the treatment of severely hearing-impaired patients. In addition to external hearing aids cochlear implants have made it possible for selected groups of patients to regain a good hearing status. Other types of experimental treatment being explored are administration of neurotrophic substances, gene therapy and cell transplantation to the cochlea or to the auditory nerve (Hildebrand et al., 2008; Hu et al., 2004a; Miller et al., 2007; Yamagata et al., 2004). Abbreviations: ABR, auditory brainstem response; AN, auditory neuropathy; bBuTx, b-bungarotoxin; BDNF, brain-derived neurotrophic factor; DAPI, 40 ,6-diamidino-2-phenylindole; NT-3, neurotrophin 3; PBS, phosphate-buffered saline; PFA, paraformaldehyde; RT, room temperature; SGN, spiral ganglion neuron; SEM, standard error of the mean; TdT, terminal deoxynucleotidyl transferase; TUJ1, antineuronal class III beta-tubulin; TUNEL, terminal deoxynucleotidyl transferasemediated biotin-dUTP nick end labelling; VGN, vestibular ganglion neuron. * Corresponding author. Address: Department of Clinical Neuroscience, Section of Otorhinolaryngology, Karolinska Institutet, Karolinska University Hospital, 171 76 Stockholm, Sweden. Tel.: +46 851770000. E-mail address: bjorn.palmgren@karolinska.se (B. Palmgren). 0378-5955/$ - see front matter Ó 2010 Elsevier B.V. All rights reserved. doi:10.1016/j.heares.2010.02.009 Hearing deteriorates when normal cochlear structures such as hair cells, cochlear nerve and spiral ganglion neurons (SGNs) are compromised. Primary damage to the cochlear nerve can occur, e.g. following neurotrauma, a tumour in the pontine angle or surgery and radiation to the area. It has also been shown that degeneration of the auditory nerve and SGNs can occur following auditory neuropathy (AN) (Starr et al., 2000) or secondary to a hair cell loss that reduces the neurotrophic support of the SGNs (Keithley and Croskrey, 1990; Kusunoki et al., 2004). Various cell transplantation strategies have been suggested for replacement of the damaged hair cells and/or SGNs (Hu et al., 2004a; Li et al., 2004; Martinez-Monedero and Edge, 2007; Sekiya et al., 2006). However, partially due to the complex structure and physiology of the hair cells several strategies for their replacement have not lead to satisfactory results (Beisel et al., 2008; Iguchi et al., 2004). The method presented here comprises an in vivo technique to deafen animals without traumatizing hair cells. b-bungarotoxin (b-BuTx) is a presynaptic neurotoxin isolated from the venom of the Taiwan banded krait (Bungarus multicinctus). It is composed of two subunits, A and B, where A has a phospholipase A2 activity and B selectively inhibits voltage-dependent potassium channels B. Palmgren et al. / Hearing Research 265 (2010) 70–76 (Liu et al., 2009). The bite of a Taiwan banded krait causes rapid neuromuscular paralysis, which has been associated with depletion of synaptic vesicles from motor nerve terminals and subsequent degeneration of motor nerve terminals and intramuscular axons (Dixon, 1999; Montecucco, 2000). Further, destruction of motor terminals was found to be reversible with extensive reinnervation by sprouting and collateral innervation of muscle fibres already 3–5 days after toxin exposure (Dixon, 1999). The effect on the bipolar SGN is less studied, however, but it seems that the toxin is deafferentating hair cells without traumatizing them (MartinezMonedero et al., 2006). In a combined in vivo and in vitro study on chick embryos and cultured SGNs, the number of SGNs was reduced by the toxin whereas the number of hair cells was not (Hirokawa, 1977). The neuronal effect is considered to be a result of increased intracellular Ca2+ and accumulation of reactive oxygen species leading to apoptosis (Hirokawa, 1977; Shakhman et al., 2003). As stated previously there is a need for models in which possible treatments and pathological mechanisms of injuries and diseases can be evaluated. In the present in vivo study we wanted to investigate whether b-BuTx can be used in a rat model to mimic SGN injury with intact hair cells. Nerve degeneration in deaf patients and aplasia of the cochlear nerve in newborns are examples of conditions that could be studied here. Such a model can also be of assistance in studies of cell replacement therapies where axonal guidance of the implant is of importance. As compared to cochlear damage with destroyed hair cells this would probably provide more appropriate electrophysiological and neurotrophic conditions for studies on a selective auditory nerve lesion. 2. Materials and methods Forty-one female Sprague–Dawley rats (SD) weighing 230– 270 g were obtained from Scanbur (Stockholm, Sweden). The animals were kept in cages under standard colony conditions with food and water available ad libitum. All animal-related procedures were conducted in accordance with local ethical guidelines and approved animal care protocols (approval no: N330/07, N38/09). Ten rats underwent auditory brainstem response (ABR) measurement and cochleas from 19 rats were used for immunohistochemistry and SGN quantifications. The remaining 12 rats were used for cochlear surface preparations and hair cell quantifications (supplementary Table 1). 2.1. Application of b-BuTx to the round window niche The rats were anaesthetized via an intraperitoneal injection of xylazine (10 mg/kg, i.p.) and ketamine (50 mg/kg, i.p.). A retroauricular incision was made, the bulla of the temporal bone opened and the middle ear and round window niche exposed. We applied 5 ll of b-BuTx (0.05 lg/ml, Alexis Biochemicals) or 5 ll of saline (sham-operated animals) absorbed by gel foam to completely fill the round window niche. A piece of fascia was placed to cover the hole in the bulla. Postoperatively the rats received subcutaneous injections of 3 ml 0.9% NaCl and 0.2 ml Temgesic (0.3 mg/ml). The survival period ranged from 3 to 21 days after which the rats were sacrificed by an overdose of pentobarbital (60 mg/ml, i.p.). All animals in the study withstood the surgery and the toxin application well. No animals had to be sacrificed due to the humane endpoints in the animal care protocol above. 2.2. Auditory brainstem response measurements ABR measurements were performed on both ears of sham-operated (n = 4) and b-BuTx-treated (n = 6) animals postoperatively on 71 days 3, 7 and 21. The ABR measurements were conducted under general anaesthesia with ketamine (50 mg/kg, i.p.) and xylazine (10 mg/kg, i.p.). ABR measurements were performed in a soundproof booth using a Tucker-Davis System II (BioSig stimulate/ recording system 2.0, Tucker-Davis Technologies, Alachua, FL, USA). Stimulus intensity was calibrated with a 0.25-in. condenser microphone (model 4135, Brüel & Kjær, Nærum, Denmark). All sound pressure levels were expressed in decibels relative to 20 lPa. The sound stimulation (tone burst 20 stimuli/s; single sinusoidal wave) was applied to the left ear using a high frequency transducer via a flexible tube in the external auditory meatus. Needle electrodes were placed on the vertex and below the recorded ear with the ground electrode placed on the hind leg. The evoked response was amplified 100 000 times and 2048 sweeps were averaged in real time by a digital signal processor (DSP32C, Lucent Technologies) with a time-domain artefact rejection. The initial intensity of the stimulus was 90 dB peak sound pressure level that was decreased in 5 dB steps until the ABR disappeared. The ABR threshold was defined as the lowest intensity at which a visible ABR wave was observed in two averaged runs. The threshold was measured at four frequencies (3.5, 7, 16 and 28 kHz). Statistical analysis of mean values from ABR thresholds at each frequency was performed by Student’s two-tailed t test between sham-operated and b-BuTx-treated ears. 2.3. Immunohistochemistry Under deep anaesthesia (pentobarbital 60 mg/ml, i.p.) the rats were perfused transcardially with 0.9% NaCl followed by 4% of paraformaldehyde (PFA). The cochleas were dissected and a small hole was made in the apex after which they were perfused with PFA (initially 4% and then 0.5%). The cochleas were decalcified in EDTA (ethylenediaminetetraacetic acid) for 7 days. After one day in 20% glucose solution the cochleas were embedded and frozen in OCT compound (Tissue-Tek, Sakura Finetek, Torrance, CA, USA) and 12 lm mid-modiolar cryosections were made for mounting on glass slides. The sections were initially blocked with 10% goat serum, 5% bovine serum albumin (BSA) and 0.2% Triton X-100 in 0.1 M phosphate-buffered saline (PBS) for 1 h at room temperature and then incubated overnight at 4 °C with mouse anti-neuronal class III beta-tubulin (TUJ1) antibody (1:200, Covance Research Products, Berkeley, CA, USA) and rabbit polyclonal anti-myosin VIIa (Myo7a) antibody (1:1000). Following incubation the cryosections were labelled with anti-mouse fluorescein isothiocyanate (1:400) secondary antibodies (Jackson Immunoresearch Europe, Newmarket, Suffolk, UK) and anti-rabbit-Cy3 (1:2000) for 1 h at room temperature. The specimens were visualized and photographed using a fluorescence microscope (Zeiss, Stockholm, Sweden) equipped with a digital camera (Nikon Coolpix 990, Solna, Sweden). Omission of primary antibody served as negative control. Cell nuclei were stained with 40 ,6-diamidino-2-phenylindole (DAPI). 2.4. TUNEL staining The cochlear cryosections used for TUNEL (terminal deoxynucleotidyl transferase biotin-dUTP nick end labelling) staining were prepared as described for immunohistochemistry above. TUNEL assay was performed using ApopTag Plus Peroxidase In Situ Apoptosis Detection Kit (Chemicon, Solna, Sweden) following the manufacturer’s instruction. Sections were post-fixed in ethanol/ acetic acid (2:1) for 5 min at 20 °C. Endogenous peroxidase was quenched in 3% hydrogen peroxide in PBS for 5 min at room temperature (RT). After incubation with terminal deoxynucleotidyl transferase (TdT) enzyme in a humidified chamber for 1 h at 37 °C followed by anti-digoxigenin peroxidase conjugate for 72 B. Palmgren et al. / Hearing Research 265 (2010) 70–76 30 min at RT, sections were developed with diaminobenzidine substrate for 3 min at RT and counterstained in methyl green solution (Vector Laboratories, Orton Southgate, Peterborough, UK) for 1 min at 60 °C. The sections were examined under a light microscope (Zeiss). Substitution of TdT enzyme with distilled water was used as negative control. membrane containing the organ of Corti was dissected into halfturns. Each piece of the basilar membrane was placed in an 8-well microscopic slide, examined under a fluorescence microscope (Zeiss) and documented with a digital camera (Nikon Coolpix 990). 3. Results 2.5. Quantification of cochlear spiral ganglion neurons 3.1. Increase of ABR threshold in b-BuTx-treated ears from day 3 To verify the effect of b-BuTx on the SGNs, SGN survival was quantified by measuring the neural density in the cochlear sections. The contralateral cochlea served as control. Out of the representative mid-modiolar sections six were randomly selected from each cochlea and used for quantitative analysis. The sections were put in 1% toluidine for 1 min and then rinsed in distilled water for 5 min. After drying, the sections were placed and examined under a microscope. For quantification of the number of surviving SGNs and measurement of the Rosenthal’s canal area the imaging software cell-B (Olympus Life and Material Science Europe, Hamburg, Germany) was used. The data were generated from microscope images. Neuronal profiles with the shape and size of a SGN (cell diameter 10–20 lm and a visible central nucleus) were counted. The borders of Rosenthal’s canal on all cochlear turns were manually marked on the computer and this area was calculated by using the imaging software above. The cell density was generated from the data and statistical analysis was performed using Kruskal–Wallis ANOVA on Ranks test. 2.6. Cochlear surface preparation The cochleas were dissected out from the temporal bone, fixed overnight in 4% PFA at 4 °C and further stored in 0.5% PFA at 4 °C. The contralateral cochleas served as control. After washing with PBS the bony capsule surrounding the cochlea, the cochlear lateral wall and Reissner’s membrane were removed. The remaining part of the cochlea was made permeable with 0.3% Triton X-100 for 10 min and stained with TRITC-phalloidin (1:200 Sigma) for 45 min. After being washed 3 times with PBS the entire basilar On postoperative day 3 at all frequencies tested, the ABR thresholds from the b-BuTx-treated ears were significantly higher than those from the sham-operated ears. At 16 kHz, e.g. the mean value of ABR threshold from the b-BuTx-treated ears was 69 ± 8 dB which was significantly higher than that of the sham-operated ears (32 ± 2 dB) (Fig. 1 and supplementary Fig. 1). The elevated ABR thresholds in the b-BuTx-treated ears were observed until postoperative day 21, which was the latest postoperative survival time examined (Fig. 1). In contrast, ABR thresholds measured from the contralateral ears were not affected and fell within the normal hearing range (data not shown). 3.2. Delayed and close to total loss of SGNs but intact hair cells The microscopic structure of the labyrinth was observed to be intact. Qualitatively, all toluidine-stained SGN sections until postoperative day 14 showed a well-preserved morphology with the SGN nuclei clearly visible, enabling good conditions for neuronal analysis. TUJ1 staining illustrated that all visible SGN profiles had a well-preserved soma with DAPI-positive nuclei. Outer and inner hair cells examined by immunohistochemistry and cochlear surface preparations revealed a proper intact morphology and ensured good conditions for hair cell assessment (Figs. 2 C, F, I and J). Following application of b-BuTx, the immunohistochemical analysis illustrated that on days 3, 7 and 14 the SGNs, including their TUJ1-positive fibres, were well preserved. Qualitatively, at this time of survival there was no reduction in SGN numbers, but Fig. 1. ABR thresholds (mean ± SEM) at frequencies 3.5, 7, 16 and 28 kHz in sham-operated (n = 4) and b-BuTx treated (n = 6) ears at postoperative days 3, 7 and 21 (**P < 0.01 and ***P < 0.001). B. Palmgren et al. / Hearing Research 265 (2010) 70–76 73 Fig. 2. (A–I) Immunohistochemical staining on cochlear cryosections at 21 days postoperative survival time. (A, D, G) TUJ1-positive spiral ganglion neurons (green) in the intact contralateral cochlea are illustrated adjacent to the b-BuTx treated cochlea. (B, E, H) The b-BuTx treated cochleas illustrate a significant reduction in TUJ1-stained spiral ganglion neurons (white arrow). (C, F, I) Myosin VIIa positive outer (arrowhead) and inner (arrow) hair cells appeared normal in the b-BuTx treated cochleas. The nuclei are counterstained with DAPI. (J) A representative image of a basal turn surface preparation from a b-BuTx treated cochlea at day 21 illustrating intact outer and inner hair cells. (K) No TUNEL-positive spiral ganglion neurons were observed in a 17-day control cochlea. (L) In the 17-day b-BuTx treated cochlea apoptotic cells are illustrated with the TUNEL assay in the basal turn (brown, black arrow and inset). The nuclei are counterstained with methyl green. Scale bars: (A–I) 20 lm, (J) 40 lm, (K) and (L) 50 lm, (L) inset: 5 lm. on day 21 there appeared to be a close to total loss of the SGNs (Figs. 2B, E, H). SGN quantification was performed following toluidine staining of the sections (supplementary Fig. 2). In the control sections the mean number of SGNs was 27 ± 1 SGN/10 000 lm2. There was no observed loss of SGNs in the b-BuTx-treated cochleas following 3 (26 ± 1 SGN/10 000 lm2), 7 (23 ± 2 SGN/10 000 lm2) and 14 days (25 ± 2 SGN/10 000 lm2) postoperative survival. However, in the day 21 animals all cochleas showed a significant reduction of SGNs in all examined cochlear turns (1 ± 0.3 SGN/10 000 lm2) (Fig. 3B). The SGN degeneration occurred between days 14 and 21, illustrating a delayed effect of the b-BuTx on neuronal morphology as compared to the effect on hearing thresholds. There was a tendency for the effect of b-BuTx to be more severe in the basal cochlear turns than in the apical turns (Figs. 3A, a1, a2) although this difference was not statistically verified. No morphological sign of SGN degeneration in the b-BuTx-treated cochleas was observed before day 14 (supplementary Fig. 2). In the b-BuTx-treated cochleas at postoper- ative survival times 17, 19 and 21, as shown by TUNEL staining (Fig. 2L and inset), too few remaining SGNs were TUNEL positive to allow statistical analyses between the groups. In the sham-operated cochleas no TUNEL-positive labelling was observed (Fig. 2K). Further, cochlear surface preparation at postoperative day 21 illustrated no loss of hair cells (Fig. 2J). Immunohistochemistry with the hair cell marker myosin VIIa revealed intact inner and outer hair cells (Figs. 2C, F, I). Qualitative analysis of the vestibular ganglion neurons (VGN) did not reveal any signs of degeneration at days 3 and 7, whereas at day 21 some of the VGNs displayed swelling of the soma (supplementary Fig. 3). 4. Discussion In mammals it is known that hair cell damage can initiate a retrograde degeneration of SGNs but also that the supporting cells play a vital role in intercellular regulations (Bichler et al., 1983; 74 B. Palmgren et al. / Hearing Research 265 (2010) 70–76 Fig. 3. (A) One 21-day mid-modiolar cross-section from the b-BuTx-treated cochlea. In the apical turn (a1) merely a few surviving spiral ganglion cells (arrowhead) reside inside the Rosenthal’s canal. At the basal turn (a2) almost all spiral ganglion cells have degenerated. The cross-section is stained with toluidine blue and eosin. Scale bars: (A) 100 lm, (a1) and (a2) 20 lm. (B) Box plot graph showing values of spiral ganglion cell density in b-BuTx-treated cochleas at postoperative days 3, 7, 14 and 21 (n = 12). The data were calculated from the mean value of all cochlear turns. Values from the contralateral cochlea at day 21 served as control. Ipsilaterally there was a significant decrease in spiral ganglion cell density after 21 days (**P < 0.01). The ends of the boxes define the 25th and the 75th percentile with a line at the median value. The error bars define the 10th and the 90th percentile. Black dots represent outliers. Spoendlin, 1975; Suzuka and Schuknecht, 1988). Several endogenous neurotrophic factors affect the proliferation, differentiation, migration and survival of both innate and implanted cells. Brainderived neurotrophic factor (BDNF) and neurotrophin 3 (NT-3) are expressed in the inner ear sensory epithelia providing neurotrophic support to the inner ear neurons (Ernfors et al., 1995; Farinas et al., 2001). Further, BDNF, NT-3 and glial cell line-derived neurotrophic factor stimulate stem cells towards migration, neural proliferation and differentiation (Rask-Andersen et al., 2005). Thus, by preserving the hair cells the inner ear would retain a major part of its trophic support. In order to provide an animal model for conditions caused by selective pathology to the auditory nerve viable hair cells are of importance. Such a model may also be beneficial for cell transplantation studies, where it, by maintaining neurotrophic support from hair cells, could stimulate differentiation and migration of implanted cells. In the present paper we have used b-BuTx as a means of damaging SGNs while preserving the hair cells. Our results show that b-BuTx has a significant effect on hearing as well as on SGN numbers. The ABR curves were affected after three days whereas the number of SGNs was not reduced until after 14 days, illustrating that the increase in hearing thresholds preceded the SGN loss. As compared to one previous in vitro study (Martinez-Monedero et al., 2006) we applied b-BuTx at a rather low concentration. Even though dose–effect responses from in vitro studies cannot be directly transferred into in vivo studies, we speculate that following a higher concentration of the toxin, our observed discrepancy between reduced auditory function and loss of SGN would be diminished. However, our observation that functional reduction precedes structural degeneration was also shown previously (Lubka et al., 2008; Megerian et al., 2008; Takahashi et al., 1999). The apoptosis of SGNs after postoperative day 14, here confirmed by TUNEL staining, is consistent with earlier findings where b-BuTx-treated chick embryos demonstrated SGN degeneration by postoperative day 14 (Hirokawa, 1977). Apoptosis of the SGNs has also been verified in earlier in vitro studies (Martinez-Monedero et al., 2006). Further, the slightly more severe effect of the toxin on the SGNs in the basal turn, as observed by us, may be due to a basal to apical concentration gradient of the toxin. This is consistent with earlier findings on the effect of round window application of other drugs and toxins (Plontke et al., 2007; Plontke et al., 2008). B. Palmgren et al. / Hearing Research 265 (2010) 70–76 Previous techniques generating iatrogenic damage to the auditory nerve include intracochlear injection of ototoxic substances (Hu et al., 2004b; Miller et al., 1997). Aminoglycosides (McFadden et al., 2004) and cisplatin (Ding et al., 1999; Rybak et al., 2007) affect the afferent SGNs but also the hair cells. In one study on gerbils it was shown that application of ouabain to the round window leads to a partial to complete loss of the auditory nerve function whereas the hair cells were kept relatively intact (Schmiedt et al., 2002). There are indications that ouabain induces apoptosis in type I SGNs while most type II neurons survive (Lang et al., 2005). Others have shown that ouabain application to the round window leaves the inner hair cells intact but causes degeneration of outer hair cells and limbal fibrocytes (Hamada and Kimura, 1999). A different model of selective nerve damage is compression of the auditory nerve. To access the nerve a suboccipital craniotomy is performed followed by compression of the nerve and the labyrinthine artery between the brainstem and the temporal bone at the internal auditory meatus. The results have been promising but include rather traumatizing surgery (Sekiya et al., 2000). In vitro rat cochlear studies also report that sodium salicylate can selectively induce auditory neuronal degeneration (Zheng and Gao, 1996). Disorders to the auditory nerve include AN (Matsumoto et al., 2008; Starr et al., 2000). In this condition, the pathophysiological mechanisms are not yet fully understood but it has been shown that ABR thresholds are significantly elevated, a finding which does not correspond to the relatively normal otoacoustic emissions (Harrison, 1998). Hearing impairment in AN has further been suggested to occur either in isolation or due to a generalized neuropathic process (Starr et al., 1996). Along the peripheral auditory pathway the affected locations may vary, but they include outer hair cells, SGN synapses and auditory nerve fibres (Starr et al., 2000). We suggest that the present b-BuTx model may be suitable for AN studies. It has previously been shown that the permeability of the round window membrane affords a good locus for administration of drugs to the inner ear (Ito et al., 2005; Saber et al., 2009). Our surgical approach, using application of b-BuTx to the round window niche, is less traumatic for the animals as compared to surgery directly into the inner ear or to the auditory nerve. Direct injection to the inner ear can cause structural damage and change homeostasis and auditory function (Borkholder, 2008). Because of the risk of side effects and possibly complications due to the blood–inner ear barrier, systemic administration of toxins may not be suitable (Juhn and Rybak, 1981). We did not examine the effect of b-BuTx on the cochlear supporting cells. Neither was the effect of b-BuTx on utricular and saccular macula specifically analysed. Some signs of soma swelling were observed in the VGNs on day 21 although the rats showed no sign of dizziness or disorientation during the entire 21 days postoperative survival time. We cannot exclude the possibility that with longer survival times or with higher concentrations of b-BuTx the VGNs could be affected. Our interpretation is that the present experimental set-up is not suitable for studies on vestibular disorders even though this could be re-evaluated with altered toxin concentrations and survival times. Further, in our SGN quantifications we did not differentiate between SGN types I and II. However, since there was a close to total loss of SGNs it is likely that both types of neurons were affected. The aim of the present study was to create an in vivo inner ear injury model resulting in selectively auditory neuronal damage, i.e. to destroy the SGNs but to keep the hair cells intact. Our results show that b-BuTx application to the round window niche is a feasible way of deafening rats when intact hair cells are important for the study. For selected types of diseases this model provides basic conditions that are closer to the clinical practice, as compared to 75 surgery on normal hearing rats or rats that have had their inner ears more severely disrupted. The present method could, for example, be used for implantation studies in order to investigate the effect of application of stem cells on the deafferentated SGNs. Patients who may suffer from various diseases including genetic disorders, tumours and trauma, could benefit from such a study. Acknowledgements This work was supported by the Swedish Scientific Research Council, The Marianne and Marcus Wallenberg Foundation, The Foundation Tysta Skolan and the Organization for Hard of Hearing People. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.heares.2010.02.009. References Beisel, K., Hansen, L., Soukup, G., Fritzsch, B., 2008. Regenerating cochlear hair cells: quo vadis stem cell. Cell Tissue Res. 333, 373–379. Bichler, E., Spoendlin, H., Rauchegger, H., 1983. Degeneration of cochlear neurons after amikacin intoxication in the rat. Arch Otorhinolaryngol. 237, 201–208. Borkholder, D.A., 2008. State-of-the-art mechanisms of intracochlear drug delivery. Curr. Opin. Otolaryngol. Head Neck Surg. 16, 472–477. Ding, D.L., Wang, J., Salvi, R., Henderson, D., Hu, B.H., McFadden, S.L., Mueller, M., 1999. Selective loss of inner hair cells and type-I ganglion neurons in carboplatin-treated chinchillas. Mechanisms of damage and protection. Ann. N. Y. Acad. Sci. 884, 152–170. Dixon, R.W., 1999. Nerve terminal damage by beta-bungarotoxin: its clinical significance. Am. J. Pathol. 154, 447–455. Ernfors, P., Van De Water, T., Loring, J., Jaenisch, R., 1995. Complementary roles of BDNF and NT-3 in vestibular and auditory development. Neuron 14, 1153– 1164. Farinas, I., Jones, K.R., Tessarollo, L., Vigers, A.J., Huang, E., Kirstein, M., de Caprona, D.C., Coppola, V., Backus, C., Reichardt, L.F., Fritzsch, B., 2001. Spatial shaping of cochlear innervation by temporally regulated neurotrophin expression. J. Neurosci. 21, 6170–6180. Hamada, M., Kimura, R.S., 1999. Morphological changes induced by administration of a Na+, K+-ATPase inhibitor in normal and hydropic inner ears of the guinea pig. Acta Otolaryngol. 119, 778–786. Harrison, R.V., 1998. An animal model of auditory neuropathy. Ear Hear. 19, 355– 361. Hildebrand, M.S., Newton, S.S., Gubbels, S.P., Sheffield, A.M., Kochhar, A., de Silva, M.G., Dahl, H.H., Rose, S.D., Behlke, M.A., Smith, R.J., 2008. Advances in molecular and cellular therapies for hearing loss. Mol. Ther. 16, 224–236. Hirokawa, N., 1977. Disappearance of afferent and efferent nerve terminals in the inner ear of the chick embryo after chronic treatment with beta-bungarotoxin. J. Cell Biol. 73, 27–46. Hu, Z., Ulfendahl, M., Olivius, N.P., 2004a. Central migration of neuronal tissue and embryonic stem cells following transplantation along the adult auditory nerve. Brain Res. 1026, 68–73. Hu, Z., Ulfendahl, M., Olivius, N.P., 2004b. Survival of neuronal tissue following xenograft implantation into the adult rat inner ear. Exp. Neurol. 185, 7–14. Iguchi, F., Nakagawa, T., Tateya, I., Endo, T., Kim, T.S., Dong, Y., Kita, T., Kojima, K., Naito, Y., Omori, K., Ito, J., 2004. Surgical techniques for cell transplantation into the mouse cochlea. Acta Otolaryngol. Suppl., 43–47. Ito, J., Endo, T., Nakagawa, T., Kita, T., Kim, T.S., Iguchi, F., 2005. A new method for drug application to the inner ear. ORL J. Otorhinolaryngol. Relat. Spec. 67, 272– 275. Juhn, S.K., Rybak, L.P., 1981. Labyrinthine barriers and cochlear homeostasis. Acta Otolaryngol. 91, 529–534. Keithley, E.M., Croskrey, K.L., 1990. Spiral ganglion cell endings in the cochlear nucleus of young and old rats. Hear. Res. 49, 168–177. Kusunoki, T., Cureoglu, S., Schachern, P.A., Baba, K., Kariya, S., Paparella, M.M., 2004. Age-related histopathologic changes in the human cochlea: a temporal bone study. Otolaryngol. Head Neck Surg. 131, 897–903. Lang, H., Schulte, B.A., Schmiedt, R.A., 2005. Ouabain induces apoptotic cell death in type I spiral ganglion neurons, but not type II neurons. J. Assoc. Res. Otolaryngol. 6, 63–74. Li, H., Corrales, C.E., Edge, A., Heller, S., 2004. Stem cells as therapy for hearing loss. Trends Mol. Med. 10, 309–315. Liu, W.H., Kao, P.H., Lin, S.R., Chang, L.S., 2009. Membrane-damaging activity with A chain and B chain of beta-bungarotoxin. Toxicon 53, 262–268. Lubka, M., Muller, M., Baus-Loncar, M., Hinz, M., Blaschke, K., Hoffmann, W., Pfister, M., Lowenheim, H., Pusch, C.M., Knipper, M., Blin, N., 2008. Lack of Tff3 peptide 76 B. Palmgren et al. / Hearing Research 265 (2010) 70–76 results in hearing impairment and accelerated presbyacusis. Cell Physiol. Biochem. 21, 437–444. Martinez-Monedero, R., Edge, A.S., 2007. Stem cells for the replacement of inner ear neurons and hair cells. Int. J. Dev. Biol. 51, 655–661. Martinez-Monedero, R., Corrales, C.E., Cuajungco, M.P., Heller, S., Edge, A.S., 2006. Reinnervation of hair cells by auditory neurons after selective removal of spiral ganglion neurons. J. Neurobiol. 66, 319–331. Matsumoto, M., Sekiya, T., Kojima, K., Ito, J., 2008. An animal experimental model of auditory neuropathy induced in rats by auditory nerve compression. Exp. Neurol. 210, 248–256. McFadden, S.L., Ding, D., Jiang, H., Salvi, R.J., 2004. Time course of efferent fiber and spiral ganglion cell degeneration following complete hair cell loss in the chinchilla. Brain Res. 997, 40–51. Megerian, C.A., Semaan, M.T., Aftab, S., Kisley, L.B., Zheng, Q.Y., Pawlowski, K.S., Wright, C.G., Alagramam, K.N., 2008. A mouse model with postnatal endolymphatic hydrops and hearing loss. Hear. Res. 237, 90–105. Miller, J.M., Le Prell, C.G., Prieskorn, D.M., Wys, N.L., Altschuler, R.A., 2007. Delayed neurotrophin treatment following deafness rescues spiral ganglion cells from death and promotes regrowth of auditory nerve peripheral processes: effects of brain-derived neurotrophic factor and fibroblast growth factor. J. Neurosci. Res. 85, 1959–1969. Miller, J.M., Chi, D.H., O’Keeffe, L.J., Kruszka, P., Raphael, Y., Altschuler, R.A., 1997. Neurotrophins can enhance spiral ganglion cell survival after inner hair cell loss. Int. J. Dev. Neurosci. 15, 631–643. Montecucco, C., 2000. How do presynaptic PLA2 neurotoxins block nerve terminals? Trends Biochem. Sci. 25, 266–270. Plontke, S.K., Mynatt, R., Gill, R.M., Borgmann, S., Salt, A.N., 2007. Concentration gradient along the scala tympani after local application of gentamicin to the round window membrane. Laryngoscope 117, 1191–1198. Plontke, S.K., Biegner, T., Kammerer, B., Delabar, U., Salt, A.N., 2008. Dexamethasone concentration gradients along scala tympani after application to the round window membrane. Otol. Neurotol. 29, 401–406. Rask-Andersen, H., Bostrom, M., Gerdin, B., Kinnefors, A., Nyberg, G., Engstrand, T., Miller, J.M., Lindholm, D., 2005. Regeneration of human auditory nerve. In vitro/ in video demonstration of neural progenitor cells in adult human and guinea pig spiral ganglion. Hear. Res. 203, 180–191. Rybak, L.P., Whitworth, C.A., Mukherjea, D., Ramkumar, V., 2007. Mechanisms of cisplatin-induced ototoxicity and prevention. Hear. Res. 226, 157–167. Saber, A., Laurell, G., Bramer, T., Edsman, K., Engmer, C., Ulfendahl, M., 2009. Middle ear application of a sodium hyaluronate gel loaded with neomycin in a Guinea pig model. Ear Hear. 30, 81–89. Schmiedt, R.A., Okamura, H.O., Lang, H., Schulte, B.A., 2002. Ouabain application to the round window of the gerbil cochlea: a model of auditory neuropathy and apoptosis. J. Assoc. Res. Otolaryngol. 3, 223–333. Sekiya, T., Hatayama, T., Shimamura, N., Suzuki, S., 2000. An in vivo quantifiable model of cochlear neuronal degeneration induced by central process injury. Exp. Neurol. 161, 490–502. Sekiya, T., Kojima, K., Matsumoto, M., Kim, T.S., Tamura, T., Ito, J., 2006. Cell transplantation to the auditory nerve and cochlear duct. Exp. Neurol. 198, 12– 24. Shakhman, O., Herkert, M., Rose, C., Humeny, A., Becker, C.M., 2003. Induction by beta-bungarotoxin of apoptosis in cultured hippocampal neurons is mediated by Ca(2+)-dependent formation of reactive oxygen species. J. Neurochem. 87, 598–608. Spoendlin, H., 1975. Retrograde degeneration of the cochlear nerve. Acta Otolaryngol. 79, 266–275. Starr, A., Sininger, Y.S., Pratt, H., 2000. The varieties of auditory neuropathy. J. Basic Clin. Physiol. Pharmacol. 11, 215–230. Starr, A., Picton, T.W., Sininger, Y., Hood, L.J., Berlin, C.I., 1996. Auditory neuropathy. Brain 119, 741–753. Suzuka, Y., Schuknecht, H.F., 1988. Retrograde cochlear neuronal degeneration in human subjects. Acta Otolaryngol. Suppl. 450, 1–20. Takahashi, K., Osawa, N., Ohmura, M., Kitamura, K., 1999. Evaluation of inner ear histology and auditory brainstem response in Wriggle Mouse Sagami. Acta Otolaryngol. 119, 767–772. Yamagata, T., Miller, J.M., Ulfendahl, M., Olivius, N.P., Altschuler, R.A., Pyykko, I., Bredberg, G., 2004. Delayed neurotrophic treatment preserves nerve survival and electrophysiological responsiveness in neomycin-deafened guinea pigs. J. Neurosci. Res. 78, 75–86. Zheng, J.L., Gao, W.Q., 1996. Differential damage to auditory neurons and hair cells by ototoxins and neuroprotection by specific neurotrophins in rat cochlear organotypic cultures. Eur. J. Neurosci. 8, 1897–1905.