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Exogenous delivery of neurotrophic factors into the cochlea of deafened animals rescues spiral ganglion cells (SGCs) from degeneration. To be clinically relevant for human cochlear implant candidates, the protective effect of neurotrophins should persist after cessation of treatment and the treated SGCs should remain functional. In this study, the survival and functionality of SGCs were investigated after temporary treatment with brain-derived neurotrophic factor (BDNF). Guinea pigs in the experimental group were deafened, and 2 weeks later, the right cochleae were implanted with an electrode array and drug delivery cannula. BDNF was administered to the implanted cochleae during a 4-week period via a mini-osmotic pump. After completion of the treatment, the osmotic pumps were removed. Two weeks later, the animals were killed and the survival of SGCs was analyzed. To monitor the functionality of the auditory nerve, electrically evoked auditory brainstem responses (eABRs) were recorded in awake animals throughout the experiment. BDNF treatment resulted in enhanced survival of SGCs 2 weeks after cessation of the treatment and prevented the decreases in size and circularity that are seen in the untreated contralateral cochleae. The amplitude of the suprathreshold eABR response in BDNF-treated animals was significantly larger than in deafened control animals and comparable to that in normal-hearing control animals. The amplitude in the BDNF-treated group did not decrease significantly after cessation of treatment. The eABR latency in BDNF-treated animals was longer than normal and comparable to that in deafened control animals. These morphological and functional findings demonstrate that neurotrophic intervention had a lasting effect, which is promising for future clinical application of neurotrophic factors in implanted human cochleae.
A progressive loss of spiral ganglion cells (SGCs) after loss of inner hair cells is presented in several animal models (Spoendlin 1975; Webster and Webster 1981). In these models, electrical stimulation (Lousteau 1987; Hartshorn et al. 1991; Leake et al. 1991) and administration of neurotrophins (Staecker et al. 1996; Miller et al. 1997; Gillespie and Shepherd 2005; Pettingill et al. 2007) via a cochlear implant are effective in preventing SGCs from degeneration after induced deafness. The preservation of SGCs might be important since the success of the cochlear implant is thought to be related to the number of excitable SGCs. In the majority of studies, neurotrophic treatment was started within the first 2 weeks after deafening (see review, Gillespie and Shepherd 2005). It has been shown that neurotrophic treatment is also effective after degeneration has set in, a condition that mimics the clinical situation (Gillespie et al. 2004; Yamagata et al. 2004; Wise et al. 2005; Miller et al. 2007; Agterberg et al. 2008; Glueckert et al. 2008). In these studies, intracochlear infusion of neurotrophins by means of an osmotic pump system was started 2–6 weeks after deafening. Neurotrophic treatment prevented SGC degeneration, but SGCs treated with BDNF were larger than SGCs in normal cochleae and their myelin layers were reduced (Richardson et al. 2005; Shepherd et al. 2005, 2008; Agterberg et al. 2008; Glueckert et al. 2008). These morphological changes might reflect a suboptimal condition. Nevertheless, SGCs treated with neurotrophins remained electrically excitable and animals treated with neurotrophins showed reduced thresholds of electrically evoked auditory brainstem responses (eABRs) as compared to animals in deafened control groups infused with artificial perilymph (Shinohara et al. 2002; Yamagata et al. 2004; Shepherd et al. 2005; Maruyama et al. 2008).
An important question with respect to clinical application of neurotrophins is whether continuous treatment is necessary for lasting protective effects. According to the neurotrophin hypothesis and based upon findings in other sensory systems (Montero and Hefti 1988; Mansour-Robaey et al. 1994), it is predicted that cessation of neurotrophic treatment would result in degeneration of SGCs. This prediction has been addressed in recent studies, which yielded contradictory results. Gillespie et al. (2003) and Shepherd et al. (2008) reported an abnormally rapid degeneration of SGCs after cessation of BDNF treatment. Two weeks after cessation, the number of SGCs was not significantly different from that in deafened, untreated cochleae. In addition, Shepherd et al. (2008) found that chronic electrical stimulation partially reduced the rate of SGC loss in the basal cochlear turn, i.e., the area adjacent to the electrode array. In contrast to these findings with BDNF, Maruyama et al. (2008) demonstrated that SGC survival and electrical responsiveness were well preserved after cessation of intracochlear infusion with glial cell line-derived neurotrophic factor (GDNF).
In the present study, we investigated the effects of cessation of BDNF treatment. Treatment was started 2 weeks after deafening, when degeneration had set in. Similar to Gillespie et al. (2003), we used BDNF as neurotrophic factor, and similar to Maruyama et al. (2008), we recorded eABRs to monitor the excitability of the SGCs. Light microscopy was used to determine the SGC packing density and to examine morphological features, such as cell size and shape (circularity). The functionality of the SGCs was assessed with the suprathreshold amplitude (Hall 1990) and latency of the first negative eABR peak.
Twenty albino female guinea pigs (strain: Dunkin Hartley; weighing 250–350 g) were purchased from Harlan Laboratories (Horst, The Netherlands) and housed in the animal care facility of Utrecht University. All animals had free access to both food and water and were kept under standard laboratory conditions. Lights were on between 7:00 am and 7:00 pm. Temperature (21°C) and humidity (60%) were kept constant.
Six guinea pigs were bilaterally deafened and 2 weeks thereafter implanted in the right cochlea with an electrode array and cannula. Consecutively, they received BDNF during a period of 4 weeks. After completion of BDNF treatment, the osmotic pumps were removed. Two weeks after BDNF treatment (i.e., 8 weeks after deafening), these animals were killed and processed for histology. For histological analysis, a comparison was made between BNDF-treated and untreated ears in the same animals; furthermore, the histological data of BDNF-treated ears were compared to data from normal cochleae (n=4 animals) presented in a previous paper (Agterberg et al. 2008).
After implantation, eABRs in BDNF-treated animals were recorded weekly to assess auditory function in relation to electrical stimulation of the treated cochleae. This paradigm is schematically explained in Figure 1 A. The eABR data of the treated animals were compared to eABRs in both normal-hearing and deafened untreated animals. For this, we used three groups of control animals (Fig. 1 B–D). A group of two animals were deafened and implanted in the right cochlea with a regular electrode array (without drug delivery cannula) 2 weeks thereafter, and eABRs were recorded up to more than 8 weeks after deafening (Fig. 1 B). A second group of eight normal-hearing control animals were implanted in the right cochlea with a regular electrode array and eABRs were recorded weekly (Fig. 1 C). In order to examine the effect of deafening on the eABRs in the same animals, these animals were deafened after 4 weeks and killed another 6 weeks later (in one of these animals, eABRs could not be recorded anymore in later sessions due to electrode failure). A third group of four animals were implanted as the second group but killed after 4 weeks while in normal-hearing condition (Fig. 1 D). All surgical and experimental procedures were approved by the Animal Care and Use Committee of Utrecht University (DEC-UMC # 03.04.036).
Animals were anesthetized with Domitor® (medetomidine hydrochloride; 10 mg/kg, i.m.) and Ketanest-S® ((S)-ketamine; 40 mg/kg, i.m.). Before the deafening procedure, acoustically evoked ABRs (aABRs) were recorded to check hearing thresholds. Animals were injected subcutaneously with kanamycin (400 mg/kg, s.c.) followed (15–60 min later) by slow intravenous infusion of furosemide (100 mg/kg, i.v.). This procedure, originally reported by West et al. (1973), has been shown to eliminate almost all cochlear hair cells (Gillespie et al. 2003; Versnel et al. 2007). For the intravenous infusion of furosemide, the right jugular vein was exposed and a catheter was inserted. A successful insertion was confirmed with withdrawal of blood into the catheter.
Animals were anesthetized with Domitor® (10 mg/kg, i.m.) and Ketanest-S® (40 mg/kg, i.m.). In the control animals the right cochleae were implanted with an eight-electrode array (Cochlear®, platinum ring electrodes of 0.3 mm width, inter-electrode distance 0.75 mm). In the experimental animals, the right cochleae were implanted with a six-electrode array (same electrode configuration as eight-electrode array) with drug delivery cannula (~80 mm, ID=0.8 mm) with a tip of ~20 mm (ID=0.12 mm; OD=0.16 mm) designed for guinea pigs (Cochlear®), resembling the array-cannula device described by Shepherd and Xu (2002). The right bulla was exposed retro-auricularly and a small hole was drilled to visualize the cochlea. The array was inserted 3–4 mm through a cochleostomy in the basal turn near the round window. The array cable was fixed onto the bulla with dental cement (Ketac-Cem Aplicap, ESPE dental supplies, Utrecht, The Netherlands) and connected to the skull with a screw (Brown et al. 1993) and dental cement. An Alzet® mini-osmotic pump (model 2004; flow rate 0.25 μl/h; reservoir 200 μl) was attached to the cannula and inserted into a subcutaneous pocket. The incision was closed in two layers with Vicryl®. The cannula and the mini-osmotic pump were filled with BDNF (PeproTech Inc., Rocky Hill, NJ, USA) solution (100 μg/ml). This concentration was chosen because concentrations of neurotrophins in this range (50–100 μg/ml) have proved to be effective in several studies (Gillespie et al. 2003, 2004; Yamagata et al. 2004; Wise et al. 2005; Miller et al. 2007; Agterberg et al. 2008; Shepherd et al. 2008). Bovine serum albumin (1%) was added to the BDNF solution. The pumps were incubated in sterile saline for 48 h at 37°C to guarantee a constant flow rate at implantation.
aABRs and eABRs were measured once or twice a week in awake and freely moving animals. All electrophysiological recordings were performed in a sound-attenuated chamber. The aABRs and eABRs were recorded with three stainless steel screws (8.0×1.2 mm) inserted into the skull bone 1 cm posterior to bregma, 2 cm anterior to bregma, and 1 cm lateral from bregma, respectively (Mitchell et al. 1997). Stimulus generation and signal acquisition were controlled with custom-written software and a personal computer. The stimuli were synthesized and attenuated using a Tucker-Davis Technologies TDT3 system (modules RP2, PA5 (2x) and SA1). The responses were amplified differentially using a Princeton Applied Research 113 pre-amplifier (amplification 5,000; band pass filter 0.1–10 kHz) with the posterior and anterior screws as active and reference electrodes, respectively, and the lateral screw as ground electrode. The amplified signal was digitized by the TDT3 system (module RP2) and made available for off-line analysis.
Broadband click stimuli consisting of monophasic rectangular pulses (width 20 μs; interstimulus interval 99 ms) were presented in free field, using a Blaupunkt speaker (PCxb352; 4 Ω; 30 W) positioned 10 cm above the awake guinea pig. Threshold was defined as the sound level at which the aABR was just visible. The click stimuli were presented from 75 dB above the average threshold of normal-hearing animals (~110 dB peSPL) down to threshold in 10-dB steps. Animals with a threshold shift of >50 dB, measured 14 days after deafening, were included as “deafened”.
After cochlear implantation, eABRs were recorded. Monophasic rectangular pulses generated and attenuated by the TDT3 system were converted to current pulses by a linear stimulus isolator (type A395, World Precision Instruments). The current pulses (width 20 μs; interstimulus interval 99 ms) were presented with alternating polarity to the most apical intracochlear electrode using the lateral screw in the skull as return electrode (monopolar eABR recordings). To define the peaks in the eABRs, they were compared to the peaks in the aABRs (see Fig. 2). The first positive peak (P1) in the eABR recordings was often obscured by the electrical artifact. In approximately half of the animals, in at least some of the recordings, the third positive peak (P3) was influenced by the digastric muscle response (Hall 1990). The first negative peak (N1) and second positive peak (P2) were always clearly visible and not obscured by the electrical artifact or by the digastric muscle response. Therefore, wave N1–P2 was analyzed. The N1–P2 amplitude, N1–P2 threshold, and N1 latency were determined. The N1–P2 amplitude was found to correlate well with SGC density (Hall 1990), and therefore, it was considered an appropriate parameter to examine excitability of BDNF treated SGCs. Thresholds were defined as the stimulus level that evoked a 2.0-μV (determined with interpolation) reproducible waveform. Stimuli were presented from 400 μA down to threshold with 2-dB steps. Higher current levels would evoke a whisker response. The N1–P2 amplitudes for all current levels in the experimental and control animals were analyzed using repeated-measures analysis of variance (rm ANOVA) and t tests were used to analyze measurements at single suprathreshold current levels.
After 4 weeks of BDNF treatment, the animals were anesthetized with Domitor® (10 mg/kg, i.m.) and Ketanest-S® (40 mg/kg, i.m.). After the connection of the cannula with the osmotic pump was checked, the cannula was cut off at about 40 mm from the cochlea and the osmotic pumps were removed. The open end of the remaining cannula was sutured with Vicryl®. The electrode array was left in place to record eABRs after termination of the BDNF infusion. Two weeks after cessation of BDNF treatment, the animals were euthanized for histology.
Immediately after the final eABR measurements, the left and right cochleae were fixed by intralabyrinthine perfusion with a fixative consisting of 3% glutaraldehyde, 2% formaldehyde, 1% acrolein, and 2.5% DMSO in 0.08 M sodium cacodylate buffer (pH 7.4) followed by immersion in the same fixative for 3 h at room temperature. Histological processing of the cochleae was carried out according to our standard protocol (De Groot et al. 1987). After decalcification with EDTA, the cochleae were immersed in 1% OsO4 containing 1% K4Ru(CN)6 for 2 h at 4°C and then rinsed, dehydrated, and embedded in toto in Spurr’s low-viscosity resin. After polymerization, cochleae were divided into two halves along a standardized midmodiolar plane, and these were re-embedded in fresh resin. Semithin (1 μm) sections stained with methylene blue and azur II were used for light microscopical evaluation and quantitative analyses.
The efficacy of the deafening procedure was assessed, in addition to aABR thresholds, by counting the number of remaining inner hair cells (IHCs) and outer hair cells (OHCs) at seven different locations along the basilar membrane at a half-turn spacing (B1, B2, M1, M2, A1, A2, and A3; Fig. 3).
Determination of SGC packing densities was performed as described previously (Versnel et al. 2007; Agterberg et al. 2008). SGC packing densities were determined using digitized light microscopical images of the spiral ganglia taken from five different cochlear locations (B1, B2, M1, M2, and A1; Fig. 3). Using the image analysis program NIH Image (version 1.63; US National Institutes of Health, Bethesda, MD), the bony boundaries of Rosenthal’s canal were outlined and its cross-sectional area (in mm2) was calculated. The number of SGC perikarya was counted at each location. The following populations were included in the counts: (1) all perikarya demonstrating the morphological determinants typical of type-I and type-II SGCs (for details, see Romand and Romand 1984); (2) partial and complete profiles of perikarya; and (3) perikaya with and without evident nucleus or nucleoli. SGC packing density was calculated by dividing the number of SGCs by the cross-sectional area of Rosenthal’s canal and expressed as the mean number of SGCs per square millimeter.
The cellular features perikaryal area and cell circularity—as a measure of perikaryal cell size and cell shape, respectively—were selected for further quantitative analysis. Because there is a considerable regional variation in SGC perikaryal area within the cochlea (Leake et al. 1999), measurements were performed in digitized light microscopical images of the spiral ganglion at one specific location in the basal turn (location B2 in Fig. 3). This location was chosen, because (1) the effect of BDNF on SGC packing densities is most prominent in the basal turn, and (2) it was occasionally not possible to determine SGC packing densities for location B1, which is near to the hook region, due to tangential sectioning of Rosenthal’s canal. Only type-I SGCs with an evident nucleus were measured. Perikaryal area was determined by outlining the myelin sheath surrounding the type-I perikaryon. Cell circularity is a feature that can be measured directly in NIH Image after delineating the cell’s perimeter, i.e., the myelin sheath. It is calculated as follows: 4π×A/L2, where A is the area and L is the perimeter (circularity is 1 for a perfect circle and less than 1 for an imperfect circle, e.g., 0.78 for a square).
Statistical analyses were performed using SPSS® for Windows (version 15.0.1). SGC packing densities at the different locations (B1–A1) were analyzed using repeated-measures analysis of variance (rm ANOVA). Statistical comparisons of perikaryal area and cell circularity at location B2 were made using paired t tests.
Two weeks after the deafening procedure, all control and experimental animals demonstrated threshold shifts of >60 dB for click-evoked aABRs. This severe hearing loss was microscopically confirmed. All hair cells at cochlear locations B1–A1 were lost, except in two animals (one control animal and one experimental animal). In these animals, a few inner hair cells still remained.
Figure 4 shows light micrographs of Rosenthal’s canal at cochlear location B2, providing typical examples of SGCs in cochleae of normal-hearing (A) and deafened animals (B–D), and in the left untreated and right BDNF-treated cochlea of an experimental animal (E, F). In the normal cochlea, Rosenthal’s canal contained the full complement of SGCs and nerve fibers embedded in a matrix consisting of vascularized connective tissue (Fig. 4A). Two weeks after deafening, the cellular distribution within the spiral ganglia was similar to normal (Fig. 4B). Figure 4C illustrates the dramatic loss of SGCs in the left cochlea of a control animal that was examined 6 weeks after deafening. In the right cochlea, which was electrically stimulated for short periods with the purpose to record eABRs, the loss of SGCs was equally dramatic (Fig. 4D). This indicates that brief electrical stimulation did not prevent degeneration. Eight weeks after deafening, a dramatic loss of SGCs was evident in the untreated, contralateral cochlea of a BDNF-treated animal (Fig. 4E). The surviving SGCs had lost their characteristic ovoid shape and, instead, had acquired a more elongated or dendritic appearance. In the BDNF-treated cochlea of the same animal, 2 weeks after cessation of BDNF treatment, no signs of degeneration were observed (Fig. 4F). The amount of SGCs was comparable to that in the normal cochlea, and the SGCs kept their characteristic ovoid shape.
Figure 5 compares SGC packing densities, at cochlear locations from B1 through A1, averaged across animals (n=6), in BDNF-treated cochleae with those in the untreated contralateral cochleae and with normal data (dashed lines). The data confirm the result shown in Figure 4E, F. SGC packing densities in BDNF-treated cochleae were normal at locations B1–M2 and by a factor 3 larger than those in untreated cochleae (rm ANOVA: F(1, 5)=65, p<0.001).
It is possible that electrical stimulation during eABR recordings contributed to the enhanced survival of SGCs (cf. Mitchell et al. 1997). Therefore, within untreated deafened animals, we compared the SGC packing densities in the stimulated implanted ears (see example in Fig. 4D) to those in the nonstimulated contralateral ears (see Fig. 4C). There were no statistically significant differences in SGC packing density between stimulated implanted ears and nonstimulated contralateral ears (rm ANOVA: F(1, 5)=3.2, p>0.1).
Figure 6 shows the mean values of the perikaryal area (Fig. 6A) and circularity (Fig. 6B) of SGCs at cochlear location B2. The perikaryal area of BDNF-treated SGCs was larger than that of SGCs in the untreated contralateral cochleae with a difference of 25% (paired t test, p<0.05). Circularity of BDNF-treated SGCs was larger than that of SGCs in the contralateral cochleae with a difference of 8% (paired t test, p<0.05). Perikaryal area and circularity of BDNF-treated SGCs were not statistically different from perikaryal area and circularity of SGCs in normal-hearing guinea pigs. SGCs in the implanted cochleae that were electrically stimulated during eABR recordings were larger than those in the unstimulated contralateral ears (data not shown, 12%, paired t test, p<0.05). There was no significant difference in circularity of SGCs in the implanted ears as compared to the unstimulated contralateral ears (paired t test, p>0.2).
Figure 7 shows eABR recordings in two animals (top and middle row) in the normal (A, C) and deafened (B, D) condition. The decrease in N1–P2 amplitude after deafening was prominent in the recordings after deafening for current levels of 318 and 400 μA. The thresholds in these examples did not change after deafening. The P3 of the eABR recordings in gp-sp02 (A, B) at stimulus intensities of 400 and 318 μA was influenced by the digastric muscle response (arrow). Figure 7E, F depicts eABR recordings in gp-mr02 after 4 weeks of BDNF treatment (i.e., 6 weeks after deafening) and 2 weeks after cessation of BDNF treatment (i.e., 8 weeks after deafening). The N1–P2 amplitude did not decrease after cessation of BDNF treatment, and the threshold did not change.
Figure 8 shows the input–output curves of eABRs in normal-hearing animals, animals 6 weeks after deafening, animals after 4 weeks of BDNF treatment (i.e., 6 weeks after deafening), and animals 2 weeks after cessation of BDNF treatment. The amplitude of the N1–P2 complex was decreased in deafened animals for stimulus intensities of 252 μA (unpaired t test, p<0.05), 318 μA (unpaired t test, p<0.01), and 400 μA (unpaired t test, p<0.001) as compared to the amplitude in normal-hearing animals. At lower stimulus currents (100–200 μA), the amplitude differences were smaller and not significant (p>0.2). The amplitudes in BDNF-treated animals were near the amplitudes found in normal-hearing animals (unpaired t test, p>0.4) and significantly larger than in the deafened animals for stimulus intensities of 318 μA (unpaired t test, p<0.05) and 400 μA (unpaired t test, p<0.01). There was no significant change in amplitude at any current level after cessation of treatment (paired t test, p>0.1).
Figure 9 shows the mean time course of eABR amplitudes elicited with 400 μA pulses for the BDNF-treated animals and the control animals. In both groups, amplitudes were increasing during the first 2–3 weeks after implantation. The control animals showed a gradual and significant decrease in amplitude after deafening. Six weeks after deafening, the amplitude was at 60% of the original value. The amplitude did not change significantly after cessation of BDNF treatment (p>0.6). The eABR recordings in these animals also yielded threshold and latency data which will be described in the following sections.
Figure 10 shows a moderate decrease in eABR threshold in the control group during the first weeks after implantation (rm ANOVA: p<0.05). The threshold in these animals did not change significantly during 6 weeks after deafening (rm ANOVA: p>0.5). The eABR threshold significantly decreased over time in the group temporarily treated with BDNF (rm ANOVA: p<0.01). After cessation of BDNF treatment, the eABR thresholds did not demonstrate any significant changes.
Figure 11 shows the time course of N1 latencies of eABRs to 400-μA pulses for both the BDNF-treated and control animals. Deafening caused a prominent latency change, which occurred mostly in the first 2 weeks: first a decrease and then an increase. In the deafened condition, latencies were longer by about 0.15 ms than in the normal condition (paired t test 6 weeks after deafening versus normal, p<0.001). In general, animals treated with BDNF demonstrated latencies longer than normal. During the treatment period, the variance among animals was large, with some animals having longer latencies (as compared to the deafened controls) and others having shorter latencies (as compared to controls 1 week after deafening). After cessation of treatment, all animals had longer latencies, very similar to deafened controls and significantly longer than normal-hearing controls (unpaired t test, p<0.01). Statistical analysis of the latencies performed on all animals (as in Fig. 8) showed similar outcomes.
The present study showed that temporary treatment with BDNF prevented degeneration of SGCs, which is commonly observed after inner hair cell loss in ototoxically deafened guinea pigs (Ylikoski et al. 1974; Webster and Webster 1981). Two weeks after cessation of BDNF treatment, SGC packing densities were as in cochleae of normal-hearing guinea pigs and three times greater than in the untreated contralateral cochleae (Fig. 5). Amplitudes of eABR wave N1–P2 in BDNF-treated animals were comparable to those in normal-hearing animals and larger than in untreated deafened animals.
In contrast to our findings, Gillespie et al. (2003) and Shepherd et al. (2008) reported that the number of SGCs was similar to that in untreated, contralateral cochleae, already 2 weeks after withdrawal of BDNF. In addition, Shepherd et al. (2008) reported that electrical stimulation after BDNF treatment only partially prevented degeneration of SGCs in the basal turn. The differences between their studies and our study that may contribute to the contrasting results are: (1) recordings of the eABRs, (2) concentration of BDNF, (3) method of cessation of BDNF treatment, and (4) period of deafness before the start of the neurotrophic treatment.
Considering the four issues addressed above, we propose eABR recordings as the most likely candidate to explain the discrepancy between our data and those of Gillespie et al. (2003). However, we cannot exclude any of the other three arguments.
Although the lasting effect of the neurotrophic treatment is evident in the present study (Fig. 5), this effect is only evaluated 2 weeks after cessation. Degeneration after deafening is not pronounced after 2 weeks (~80% of normal density, Versnel et al. 2007). Also the decline in eABR amplitude after deafening is rather slow (Fig. 9). Therefore, to investigate, both histologically and functionally, whether degeneration after cessation of BDNF treatment is slower than degeneration after deafening without intervention, it is important to apply longer periods of survival.
Immediately after BDNF treatment, SGCs are larger than SGCs in normal-hearing animals (Shepherd et al. 2005; Richardson et al. 2005; Agterberg et al. 2008). We found that 2 weeks after cessation SGCs had a normal size (while larger than in the untreated contralateral cochlea). This implies that the perikaryal area decreased after cessation from larger-than-normal, to normal values. The question remains if the back-to-normal area is an indication of a normal condition. On the other hand, the decrease of cell size following cessation may be an early indication for initiation of a degenerative process (Staecker et al. 1996; Leake et al. 1999; Agterberg et al. 2008). To answer this last question, the survival of SGCs needs to be investigated for longer periods after cessation of the neurotrophic treatment.
We applied suprathreshold amplitudes as a measure of excitability of the auditory nerve, since the amplitude reflects the summed neural firings of a large population of SGCs, whereas the more commonly used parameter threshold reflects the function of only the most sensitive auditory nerve fibers. Indeed, at least in case of monopolar stimulation, the amplitude of early eABR waves has been found to correlate well with the number of SGCs (Smith and Simmons 1983; Hall 1990) whereas correlations between eABR threshold and nerve survival were weak (Smith and Simmons 1983). Note that for bipolar stimulation a better correlation was found for threshold measures than for amplitude measures (Miller et al. 1994).
The normal eABR amplitude in BDNF-treated animals indicates that treated SGCs responded as normal to electric stimuli. It should be noted that the amplitude after deafening was smaller, probably because less SGCs were present, thus implicating that the remaining untreated cells responded normally. Our data agree with the results of Maruyama et al. (2008) who reported that amplitudes in neurotrophically treated animals were larger than in untreated animals and were stable after cessation of the treatment.
A decrease of eABR threshold was found the first weeks after implantation in the normal-hearing condition. This might be due to mechanisms as tissue growth around the electrodes and recovery from insertion-induced trauma (Su et al. 2008). We assume the threshold decrease found in the animals treated with BDNF was rather due to these mechanisms than to the preservation of SGCs mediated by the BDNF.
Our threshold data do not agree with other investigators who reported lower thresholds (by 4–9 dB) in deafened guinea pigs treated with neurotrophins as compared to deafened and untreated guinea pigs (Shinohara et al. 2002; Yamagata et al. 2004; Shepherd et al. 2005; Chikar et al. 2008; Maruyama et al. 2008). In these studies, the decrease in eABR threshold corresponded with enhanced survival of SGCs after neurotrophic treatment. Contrary to these studies, in which eABR thresholds were often assessed on the basis of visual inspection of peak 3 and recordings were performed in anesthetized animals, we assessed the eABR threshold on the basis of interpolation of N1–P2 amplitudes and we recorded in awake animals. Possibly, the difference in peaks analyzed explains the different threshold data as Mitchell et al. (1997) reported that the threshold of the late eABR peaks in deafened animals decreased over time, while early peak thresholds did not significantly change. Further, the state of the animal plays a role in that the threshold increases with anesthesia as was found for aABRs in mice by Van Looij et al. (2004). The effects of anesthesia may be nonlinear and differ between various experimental conditions.
The eABR N1 latencies were longer in deafened and BDNF-treated animals than in normal-hearing animals (Fig. 11). The initial latency decrease after deafening may be caused by degeneration of dendrites and a loss of spontaneous activity, and the subsequent increase can be attributed to a reduction of the number of myelin layers, which follows dendritic degeneration (Leake and Hradek 1988). The BDNF-treated nerve cells have a similar degree of myelination as untreated cells (Agterberg et al. 2008; Glueckert et al. 2008), and this might explain the similar latencies of BDNF-treated and untreated animals.
Our results demonstrate that SGC survival in deafened animals, in both morphological and functional sense, is not reduced after cessation of intracochlear application of BDNF. The enhanced survival of SGCs is a promising finding for cochlear implant candidates since there are currently no potential methods available to provide neurotrophins for periods that extend the order of months. Lifelong neurotrophic treatment via an osmotic pump is not clinically preferable because of repetitive invasive manipulations needed to replace the pump and high infection risks.
Before neurotrophic treatment can be considered a clinical option, it is important to investigate in animal studies whether neurotrophic treatment does actually improve the sound perception performance with a cochlear implant. A major topic that needs to be further explored is the assumption that more and better functioning SGCs would result in a better performance of cochlear implant users. Alternative to this hypothesis, it might be that a minimum number of SGCs is required for electrical hearing, and that an increase above this minimum would have little functional impact (Blamey 1997). Clinical data do not support the assumption that more SGCs would lead to a better behavioral performance (Nadol et al. 2001; Khan et al. 2005; Fayad and Linthicum 2006; Nadol and Eddington 2006). These data indicate that abnormalities in the central auditory pathways are at least as important as SGC survival in limiting the performance of cochlear implant users (Nadol et al. 2001). Furthermore, the overall performance of cochlear implant users improves markedly within weeks as a result of implant use, confirming the role of the central auditory system and its plasticity (Kral and Eggermont 2007). Still, we, and others, think that it is unlikely that SGC loss does not affect the performance with a cochlear implant and that the human data sets are still too limited to conclude that the extent of SGC survival does not influence cochlear implant performance (Khan et al. 2005; Leake et al. 2008). Moreover, it should be considered that with improved cochlear implant technology the number of SGCs might become more important (Green et al. 2008).
The authors would like to thank Leone Nijman and Rik Mansvelt-Beck for their excellent practical and technical assistance, Ferry Hendriksen for assisting with histology, and René van de Vosse for developing the recording and analysis software. We thank Victor Wiegant for his suggestions on the manuscript. This study was supported by the Heinsius-Houbolt Foundation.
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