Following operant conditioning with acoustic stimuli, animals were unilaterally ototoxically deafened, inoculated with one of two adenoviral constructs (Ad.BDNF or Ad.Empty), and implanted with a multichannel cochlear implant. Following implantation, psychophysical and electrophysiological thresholds were measured for 80 days. Animals were then euthanized and both cochleae were prepared for histological analysis. A separate group of animals was used to assess BDNF levels in cochlear fluids at one and two weeks following inoculation.
Eight pigmented male guinea pigs (Elm Hill, Chelmsford, Mass, USA) were used in the first part of this study, four animals in an experimental group and four animals in a control group. Animals began psychophysical training with a free-feeding schedule until weights reached 400 g, at which time a restricted diet was instituted. This restricted diet was designed to keep animals at 80 % of free-feeding weight, encourage performance in a food-reward based psychophysical task, and maintain good health. Animal weights at time of inoculation and implantation ranged from 780–1200 g. Pure tone auditory brainstem response thresholds verified normal hearing in all tested animals prior to being placed in one of the treatment groups.
2.3. Adenoviral Vectors
Animals received one of two adenoviral constructs. Animals in the experimental group received an adenoviral vector with a mouse BDNF
gene insert, driven by the cytomegalovirus promoter, labeled as Ad.BDNF
, as previously described (Di Polo et al., 1998
). Control animals received an adenoviral construct containing no gene insert, labeled as Ad.Empty (a gift from GenVec, Inc., Gaithersburg, MD, USA).
All animals were unilaterally deafened in the left ear via an aminoglycoside antibiotic (neomycin) introduced directly into the cochlea. For general anesthesia animals were given a ketamine (40 mg/kg) and xylazine (10 mg/kg) mix (IM), as well as atropine (0.05 mg/kg, SQ), and kept warm using a heating blanket. Lidocaine was used as a local anesthetic, and an incision was made in an arc caudal to the pinna of the left ear. The underlying muscle and tissue were gently pushed back to reveal the bulla and a small hole was made in the bulla with the tip of a scalpel blade. The round window was punctured and some perilymph was absorbed. Neomycin (10 % neomycin sulfate in sterile water, 60 µL) was slowly infused into the scala tympani via the round window using a 100 µL glass syringe and a 30 G needle. Following neomycin infusion, the hole in the bulla was sealed with carboxylate cement and the incision in the skin was stitched in layers. Although other methods of aminoglycoside deafening are effective (West et al., 1973
; Xu et al., 1993
), direct infusion of neomycin into the perilymph was chosen as the ototoxic method in this study to ensure a within-subject control, as this technique does not deafen the contralateral ear. Ototoxic deafening of only one ear also allows the option to implant the contralateral ear and continue behavioral experiments should the implant fail. Neomycin in volumes as low as 10 µL has been shown to not only eliminate hair cells, but also create a flat, uniform epithelium from the organ of Corti within 4 days of injection into the perilymph (Kim and Raphael, 2007
2.5. Inoculation and Implantation
All animals were unilaterally implanted in the deafened left ear. Four days after neomycin deafening, animals were given general and local anesthesia as detailed above, and an incision was made down the midline of the head. Muscle and connective tissue were gently pushed apart to reveal the skull. Six screws were placed in the skull; three for electrically-evoked auditory brainstem response recordings and three screws formed a triangle around bregma and held the head of a restraining bolt, which held the implant in place. A thin coat of acrylic was applied to these screws to aid in securing the restraining bolt. The incision from the deafening procedure was then re-opened and the carboxylate cement removed. For treated animals, a single injection of 5 µL Ad.BDNF (approximately 4×1012 adenoviral particles/mL) was placed through the round window into the perilymph of the basal turn of the scala tympani via a bent tip 30 G needle and a 10 µL glass syringe. In control animals, 5 µL of Ad.Empty was administered in a similar manner. Animals were left undisturbed for 20 minutes to allow permeation of the adenoviral solution. A small cochleostomy was then made apical to the round window to expose the scala tympani, and a multichannel cochlear implant was placed into the scala tympani. All animals received an 8-electrode implant, which resembled the apical end of the Nucleus CI-22™ human implant and was manufactured by Cochlear Corporation (Nucleus Ltd., Lane Cove, Australia). The implants had an approximate center-to-center distance between electrodes of 0.75 mm, which allowed 5–6 electrodes to be inserted into the cochlea. Post-mortem evaluations of implant position revealed an insertion depth range of 2.25 – 5.90 mm, an average of 4.27 +/− 1.09 mm, and no differences between experimental and control groups (Student’s t-test p = 0.63). Only six of the eight implant electrodes were stimulated, and were labeled A through F where A was the most apical. All electrodes were within the first turn of the cochlea. The letter G was used to identify a ground electrode, which was placed in the post-auricular muscle. See for details on the implant dimensions and labeling system.
Diagram of multichannel implant and electrode configurations
2.6. Psychophysical assessment
Animals were trained in a go-no go positive reinforcement auditory stimulus detection task, similar to that described (Miller et al., 1995
; Prosen et al., 1978
; Su et al., 2008
). Briefly, animals were placed in a wire cage inside a sound-attenuating chamber and trained to respond to acoustic stimuli (including both pulse trains and sinusoids) by releasing a button. A food reward was given for correct responses. The method of constant stimuli was used to vary sound pressure levels, and threshold was defined as the level at which correct releases occurred on 50 % of the trials. An animal was considered trained when 10 consecutive thresholds were within a range of +/− 5 dB SPL.
The day after implantation, animals began testing with the same paradigm as described above, but with electrical stimulation from the cochlear implant. Animals were tested for thresholds to cochlear implant stimulation in one 90-minute session per day, 5 days/week, for 11 weeks. In our psychophysical detection paradigm, it can take up to 40 days for animals to obtain stable thresholds to electrical stimulation. We continued our experiment for double this time (80 days) in order to obtain enough threshold values in each electrode configuration for valid statistical tests. Testing began with the most apical monopolar electrode configuration (labeled B–G, see ), which was tested until a threshold was obtained. Thresholds for each remaining configuration were then obtained on a random schedule for the remainder of the testing period. Six electrode location and configuration combinations were tested on each of the eight animals used in the study. These electrode combinations consisted of apical and basal end of the implant tripolar configurations (electrodes A–B–C and C–D–E), apical and basal bipolar configurations (A–B and D–E), and apical and basal monopolar (B–G and D–G) configurations.
The electrical stimulus for psychophysical testing was a train of 100Hz sinusoid bursts (200 msec on, 100 msec off), generated by a Tucker-Davis Technologies (TDT) digital signal processor. The stimuli were sent to a TDT programmable attenuator, then to a controlled-current stimulator and finally to the animal’s implant. To monitor any changes in implant status, electrode impedances were measured daily using a 1 µA rms, 1 kHz sinusoidal stimulus.
Electrically-evoked auditory brainstem responses (EABRs) were recorded every two weeks, beginning one week post-implantation. Electrical stimuli were generated by a TDT digital processor, sent to a programmable attenuator, fed to a controlled-current stimulator, and delivered to the implant through a percutaneous connector mounted on the animal’s skull. Neural activity was recorded using alligator clips attached to screws that were placed in the skull 2 cm anterior to bregma, 1 cm lateral of bregma on the implanted side, and 1 cm posterior to bregma. A bipolar and monopolar electrode configuration was tested at both apical and basal locations on the implant, giving four stimulating electrode pairs (A–B, B–G, D–E, and D–G, see ). Stimuli were 50 µsec phase duration monophasic alternating polarity square pulses. EABR threshold was defined as the lowest level at which there was a repeatable P3 wave, as agreed upon by two unbiased observers.
Eighty days post-implantation, the animals were deeply anesthetized and perfused intracardially with 2 % glutaraldehyde and both cochleae were removed. Tissue processing was completed as previously described (Yagi et al., 2000
), modified as described below. Each cochlea was locally perfused with 4 % paraformaldehyde and placed in EDTA solution to decalcify until sufficiently soft for sectioning (approximately 1 month). Once decalcified, the implant was removed from the left ears, and each cochlea was embedded in JB-4 resin and sectioned in the mid-modiolar plane, which provided 6 measurable profiles of Rosenthal’s canal (Kanzaki et al., 2002
). Sections were 3 µm thick, and every third section was kept and stained with 1 % Toluidine Blue in 1 % Sodium Borate. These sections were evenly divided into a caudal, middle and rostral group and one slide from each of these groups was chosen by a random number generator for evaluation. Each profile of Rosenthal’s canal in each of these three slides was evaluated by an observer who was blinded to the treatment groups, using SPOT Imaging™ software for data acquisition and Meta-Morph Offline™ software for data analysis. The outer edge of each region of Rosenthal’s canal was traced using MetaMorph Offline™ software and the two-dimensional area was calculated by the software. The two-dimensional somatic area of SGCs with clearly defined nuclei was assessed in the same way, and the number of these SGCs per Rosenthal’s canal region was counted.
2.9. Data Analysis
Log transformations on all psychophysical and electrophysiological data points were completed to normalize psychophysical and electrophysiological thresholds. For individual animal data, there were no differences seen in either change over time (slope) or range (maximum – minimum values) in psychophysical and electrophysiological thresholds. Therefore, all data points throughout the 80-day test period were averaged to determine a threshold value for each animal for each electrode configuration (six values for each animal) ( and ). Each of the electrode configurations tested represents an independent variable, and Student’s t-test was used to determine differences between experimental and control groups for each configuration.
Psychophysical detection thresholds averaged over time
EABR thresholds averaged over time
The data for one configuration (A–B) was separated by date post-implantation, and a two-way repeated measures ANOVA test was performed on both psychophysical and EABR data ( and ). Student’s multiple comparison post-hoc tests determined specific differences between groups and over time. EABRs were collected every two weeks and an average threshold and standard error were determined for each of these dates for control and treated groups. Psychophysical detection thresholds were collected on a daily basis, but not every configuration was run each day. In order to compare to electrophysiological data, psychophysical threshold levels for each animal within a two-week timeframe (i.e. data collected between 7 and 21 days post-implantation) were averaged, and these values were used to determine a psychophysical detection threshold average and standard error.
Psychophysical detection thresholds over time, A–B configuration
EABR detection thresholds over time, A–B configuration
To determine differences in morphology, SGC density and cross-sectional somatic area were compared between groups. The data from the three slides of each individual were averaged to determine a mean cross-sectional somatic area for each turn of the cochlea (lower basal, upper basal, middle, and apical) (). Density was calculated for each animal by dividing the average number of SGCs within each region of Rosenthal’s canal by the average cross-sectional area of that region. To eliminate the possibility of individual animal differences in the area of Rosenthal’s canal, we compared the areas of this region for each turn of the cochlea (). One-way ANOVAs and Student’s t-tests were used to evaluate differences between the Ad.BDNF, Ad.Empty, and non-deafened groups for each turn of the cochlea. Our non-deafened control group was composed of the right, non-deafened, non-implanted, and non-inoculated cochleae of both Ad.BDNF and Ad.Empty inoculated groups. Linear regressions were used to determine correlations between psychophysical detection thresholds and SGC density as well as between EABR detection thresholds and SGC density. All analyses were completed using SigmaStat™ (Jandel Scientific).
Two-dimensional ganglion cell and Rosenthal’s canal characteristics.
2.10. Ad.BDNF activity in vivo
In a separate experiment, six guinea pigs were inoculated with Ad.BDNF (same concentration as above) in the left cochlea through the round window, using the same surgical procedure as detailed above. One or two weeks post-inoculation (n = 3 for each timepoint), animals were sacrificed, decapitated, and both left and right temporal bones were removed. The basal turn of the cochleae from each ear was carefully thinned using fine-tipped forceps until a hole was made in the basal turn outer wall. A micro-capillary tube was gently inserted into this hole, and 1–2 µl of cochlear fluids were extracted from each cochlea. Samples were diluted 100 fold using the sample diluent from a ChemiKine™ BDNF Sandwich ELISA Kit. This kit was used to determine BDNF levels in the cochlear fluids, following the instructions in the kit. The non-inoculated right ears of both groups (n = 6) were used as controls. A one-way ANOVA was performed to determine differences between the one week inoculated ears, the two week inoculated ears, and the right, non-inoculated control ears.
2.11. Animal Care
This study was performed in accordance with National Institutes of Health Guidelines (Guide for the Care and Use of Laboratory Animals, 1996). The University Committee on the Use and Care of Animals at the University of Michigan approved the experimental protocols. Veterinary care and animal husbandry were provided by the Unit for Laboratory Animal Medicine, in facilities certified by the Association for Assessment and Accreditation of Laboratory Animal Care, International (AAALAC, Intl.)