Eur Arch Otorhinolaryngol DOI 10.1007/s00405-015-3760-0
OTOLOGY
In vitro and in vivo pharmacokinetic study of a dexamethasone-releasing silicone for cochlear implants Ya Liu1 • Claude Jolly2 • Susanne Braun3 • Thomas Stark4 • Elias Scherer4 Stefan K. Plontke5 • Jan Kiefer6
•
Received: 16 December 2014 / Accepted: 21 August 2015 Ó Springer-Verlag Berlin Heidelberg 2015
Abstract Cochlear implants have been widely used for patients with profound hearing loss and partial deafness. Residual low-frequency hearing, however, may deteriorate due to insertion trauma and tissue response around the electrode array. The present study investigated in vitro and in vivo release of dexamethasone from silicone used for cochlear implant electrode carriers. The in vitro experiment involved an apparatus simulating the inner ear fluid environment in humans. Release from two sizes of silicone films (200 lm 9 1 mm 9 10 mm and 500 lm 9 1 mm 9 10 mm), each loaded with 2 % dexamethasone, and was measured for 24 weeks. In the in vivo experiment, silicone rods loaded with 2 or 10 % dexamethasone, respectively, were implanted into the scala tympani of guinea pigs. Perilymph concentrations were measured during the first week after
& Jan Kiefer [email protected] Ya Liu [email protected] 1
Department of Otolaryngology, Head and Neck Surgery, Beijing Naval General Hospital, Beijing 100048, People’s Republic of China
2
Electrode Research Section, MED-EL Medical Electronics, Innsbruck, Austria
3
MED-EL Deutschland GmbH, Starnberg, Germany
4
Clinic for Otorhinolaryngology, Head- and Neck Surgery, Klinikum rechts der Isar, Technical University of Munich, Munich, Germany
5
Department of Otorhinolaryngology, Head and Neck Surgery, Martin Luther University Halle-Wittenberg, Halle (Saale), Germany
6
HNO-Zentrum, Neupfarrplatz 12/II, 93047 Regensburg, Germany
implantation. The results showed that dexamethasone was released from the silicone in a sustained manner. After a burst release, perilymph concentration was similar for silicone incorporated with 2 and 10 % dexamethasone, respectively. The similar pharmacokinetic profile was found in the in vitro experiment. The period of sustained drug delivery was maintained for 20 weeks in vitro and for 1 week in vivo. The results of the present study suggest that drugs like dexamethasone are released in a controlled manner from silicon electrode carriers of cochlear implants. Further studies will identify optimal release profiles for the use with cochlear implants to improve their safety and longterm performance. Keywords Cochlear implant Dexamethasone Drug delivery Inner ear Pharmacokinetics
Introduction Since the Food and Drug Administration (FDA) approved the House 3M cochlear implant system in 1984, cochlear implants have been widely used for patients with profound hearing loss. With the strategy of combined electric and acoustic stimulation (EAS) or hybrid stimulation in partial deafness, nerve fibres coding for high and mid frequencies can be electrically stimulated by the cochlear implant, while the residual low-frequency hearing can be aided by acoustic amplification [1, 2]. However, the insertion of a cochlear implant electrode may cause direct trauma or indirect early or delayed damage to the cochlea, which may lead to deterioration or loss of residual hearing [3–5]. The formation of scar tissue around the electrode after cochlear implantation is suggested to be one of the most important causes of increase in electrode impedance and loss of
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residual hearing after surgery [5, 6]. Physical disruption of the organ of Corti and molecular mechanisms of cell death induced by the electrode insertion trauma are other contributing factors [7]. Therefore, the minimization of electrode insertion trauma and reduction of scar tissue around the electrode are important for preserving residual hearing, which is necessary for successful electric-acoustic stimulation [8–10]. Methods should include the use of less traumatic implant electrodes, modification of surgical techniques [11], and pharmaceutical treatments with additional drug delivery or by the means of drug device combinations, e.g. through drug release from cochlear implant electrode carriers. The present study focused on a pharmaceutical strategy to mitigate electrode insertion trauma. The most frequently studied drugs are glucocorticoids because of their immune suppression and anti-inflammatory functions, such as inhibition of the expression of tumour necrosis factor alpha (TNF-alpha), nuclear transcription factor-jB (NF-jB), and some of the cell death signal cascades (JNK) [12, 13]. Dexamethasone is a long-acting glucocorticoid (biological half-life 36–72 h) that possesses a high anti-inflammatory potency and is stable in vitro. There are three methods for local glucocorticoid delivery in cochlear implant surgery: 1.
Single drug application to the round window membrane or cochleostomy.
Several animal [10, 14, 15] and human studies [16, 17] have examined the effects of a single glucocorticoid application in cochlear implant surgery, and have shown benefits in hearing, histology, and postoperative electrode impedance. 2.
Direct drug irrigation into the cochlea with a modified electrode array.
Several investigators modified the electrode array for sustained drug delivery into cochlea [18–21]. In most studies, the drug was delivered from a mini pump into the cochlea through or cannula or through the electrode array. This application allows variation of drug volume and delivery rate after implantation. However, the safety of the drug delivery system needs to be further investigated. 3.
Drug release from cochlear implant electrode carriers.
A few studies reported the use of an electrode array made of a biocompatible silicone in which a drug is incorporated to provide a long-term sustained release. Bohl et al. [22] demonstrated an in vitro biocompatibility study that sustained-release dexamethasone was non-toxic to cultured spiral ganglion cells. In an own study electrode dummies with 10 % dexamethasone were implanted into the inner era of guinea pigs through a cochleostomy. The histological result demonstrated the location of the
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electrode array in scala tympani, and found no increased risk of postoperative pneumococcal otogenic meningitis [23]. Stathopoulos et al. [24] performed a similar study with electrode dummies with 40 % dexamethasone in the rat and results consistent to ours. Nguyen et al. [25] found that compared with a drug-free electrode array, the insertion forces were not increased for a 0.4 mm diameter dexamethasone-eluting array for all insertion lengths. Further studies of dexamethasone-releasing electrodes addressed drug release profiles and material fabrication [26, 27]. The present study investigated dexamethasone-releasing silicone with in vitro and in vivo pharmacokinetic experiments. An in vitro apparatus was designed to simulate the inner ear fluid environment in human, and the in vivo experiments examined drug concentration in perilymph during one week after implantation.
Materials and methods Study design The aim of the present study was to investigate the pharmacokinetic aspects of a dexamethasone-releasing silicone for cochlear implants. The pharmacokinetic characterization of the device such as drug release and diffusion was investigated in an in vitro study using an apparatus to simulate the inner ear fluid environment in human. The in vitro study was followed by measurements of drug concentration in scala tympani perilymph of the guinea pig in vivo, comparing concentration time courses after implantation of silicone rods loaded with either 2 or 10 % dexamethasone. Dexamethasone-releasing silicone The dexamethasone-releasing silicone was manufactured as part of a research project with MED-EL GmbH (Innsbruck, Austria). The detailed preparation has been described previously by Farahmand Ghavi et al. [26]. Two or ten percent (weight of drug per weight of the final polymer electrode) pharmaceutical grade micronized dexamethasone (Aventis, Romainville, France) was homogeneously mixed with medical grade silicone, which is used in human cochlear implant electrode arrays. The in vitro study used two sizes of silicone films [200 lm 9 1 mm 9 10 mm (group A) and 500 lm 9 1 mm 9 10 mm (group B)] incorporated with 2 % dexamethasone. The in vivo experiment used silicone rods as an electrode analogue that contained no wires, and were similar to the electrode dummy described by Niedermeier et al. [23] and Takumi et al. [28]. The rods incorporating 2 or 10 %
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Fig. 1 Cross-section of the silicone rod used in the in vivo experiment. It was 6 mm long and 0.5 mm diameter. One end tapered to 0.5 mm. A length of 3 mm was inserted into the scala tympani of the guinea pigs
dexamethasone were adapted to a guinea pig’s anatomical dimensions, and were 6 mm long and 0.5 mm diameter. The rod was inserted into the scala tympani of the guinea pigs half its length, (i.e. for 3 mm). The intracochlear end of the rod was bevelled over a length of 0.5 mm (Fig. 1). In vitro release experiments Based on the data of the previous studies, the volume of human perilymph is 158.5–166.4 lL [29, 30]. Therefore, the capillary with a volume of 160 lL was used in the in vitro experiments, representing the human perilymph cavity. There are no studies on the perilymph flow rate in human, and the estimation has been made in the present in vitro experiment, based on the data in guinea pigs. The perilymph volume in the guinea pig is 16 lL and the flow of cerebrospinal fluid (CSF) entering perilymph is 2.304 lL per day [31, 32]. Therefore, the flow rate of human perilymph was taken as 24 lL per day, and 24 lL artificial perilymph was sampled in the present in vitro experiments. A vertical sectional view of the in vitro release apparatus is demonstrated in Fig. 2. The silicone film with 2 % dexamethasone (shadowed area in the figure) was kept in a glass capillary (outer diameter: 6.0 ± 0.2 mm; inner diameter: 2.7 ± 0.1 mm; length: 28.000 ± 0.5 m; Duran Group, Germany). The volume of the capillary was 160 lL, and was filled with artificial perilymph (an aqueous solution of 1.3 mmol/L calcium chloride, 1.8 mmol/L magnesium chloride, 5.4 mmol/L potassium chloride, 137 mmol/L sodium chloride, 5 mmol/L glucose, and 5 mmol/L HEPES). At the bottom of the capillary there was a sample needle, shortened from a HamiltonÒ 7770-01 (Germany) needle, that could be connected to a microsyringe (Hamilton 705 RN 50 lL, Germany). At the top of the capillary there was a small tube connected to a chamber, which was also filled with artificial perilymph. All components of the in vitro release apparatus were sealed with Kwik-CastTM (WPI, Los Angeles, USA) and TurboKITT (Germany). The in vitro apparatus were kept
Fig. 2 Drawing of the in vitro release experimental set-up (cross section). The silicone film (shadowed area) incorporated with 2 % dexamethasone was maintained vertically in a glass capillary containing a volume of 160 lL artificial perilymph. At the bottom of the capillary there is a sample needle. At the top of the capillary there is a small tube connected to a chamber filled with artificial perilymph
vertical. The level of the fluid in the chamber was kept higher than the top of the capillary. Therefore, when sampling fluid from the bottom of the capillary the same volume of fluid would flow from the chamber to the capillary. Five replicates of each group (n = 5) were studied at the same time. During the experiment the apparatus was kept in a 37 °C water bath, except during sampling from the capillary. Sampling was performed every day at the same time. The sample volume was 24 lL, and it was continued for 168 days. Animals for in vivo experiment The in vivo experiment consisted of 30 healthy pigmented guinea pigs (Charles River). Silicone rods containing 10 % dexamethasone (10 % DEXA group, 21 ears) or 2 % dexamethasone (2 % DEXA group, 21 ears) were implanted into the cochlea. Two ears were implanted with silicone rods containing no drug and were used as blank controls. For the reduction of animal experiments, some animals were implanted bilaterally using silicon rods with the same dexamethasone concentration.
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Surgery and anaesthesia Silicone rod implantation was performed under general anaesthesia. Guinea pig body temperature was maintained at about 38.5 °C using a Medax heater (Medax GmbH und Co. KG, Neumu¨nster, Germany). Animals were anesthetised using a mixture of medetomidine (200 lg/kg), fentanyl (25 lg/kg) and midazolam (1 mg/kg). The level of anaesthesia was maintained by regular intramuscular injections of the same mixture, if necessary [33]. The animals received subcutaneous injections of 1 % mepivacaine hydrochloride (Curasan AG, Kleinostheim, Germany) before their skin and muscle tissues were cut. A retroauricular incision was made, and after location of the bulla a small hole was drilled and the opening was enlarged with tweezers to expose the round window membrane niche and the promontory. Using a 0.8 mm diamond drill, a cochleostomy was performed in the basal turn of the cochlea about 1 mm above the ridge of the round window membrane niche. The diameter of the cochleostomy was chosen as 0.8 mm to allow optimal placement of the silicone rod. Since it was not a functional study, the issue of hearing preservation had not been taken into account. After drilling the cochleostomy a silicone rod was inserted three mm into scala tympani. Immediately after implantation, a soft tissue was placed around the rod to seal the cochleostomy. The bulla was sealed with bone wax. All operations were performed by the same surgeon (YK).
(100 9 46 mm) columns. The detection wavelength was 254 nm. Before measurement all samples were centrifuged at 1000 rpm for 10 min. The supernatant was carefully removed and mixed with the same volume of internal standard for sample injection. Compliance and ethics The study was performed in the lab of Clinic for Otorhinolaryngology, Head- and Neck Surgery, Klinikum rechts der Isar, Technical University of Munich, Munich, Germany. It was in compliance with the PHS Policy on Humane Care and Use of Laboratory Animals, the NIH Guide for the Care and Use of Laboratory Animals, and the Animal Welfare Act. The protocol (no. 6-075/06) permitting the use of animals was approved by the Government of Oberbayern (Munich, Germany). Statistical analyses Perilymph concentrations were compared using group as the test variable (in vitro experiment: group A and group B; in vivo experiment: group 2 % DEXA and group 10 % DEXA). Statistical analyses were performed with SPSS 18.0 (IBM SPSS Statistics). Differences between groups were compared using the nonparametric Mann–Whitney U test for two independent samples (1-tailed). A value of p B 0.05 was considered to indicate statistical significance.
Perilymph sampling from the cochlear apex
Results
Perilymph was sampled according to the apical sampling procedure described by Salt et al. [34]. Eight microliters of perilymph were sampled from the cochlear apex, consisting of approximately 4.5–5 lL of scala tympani perilymph and 3 lL of cerebrospinal fluid. Samples were obtained at 10 min, 30 min, 1.5 h, 3 h, 7 h, 1 day, and 1 week after surgery (each n = 3). Each implanted cochlea was sampled only once. The samples were diluted into 17 lL of artificial perilymph and stored at -20 °C. Perilymph from the cochleae implanted with a blank silicone rod was sampled 30 min after surgery. After sampling, animals were sacrificed with an intracardiac injection of pentobarbital (40 mg/kg, Meral GmbH, Hallbergmoos, Germany).
In vitro release
High performance liquid chromatography (HPLC) Concentrations of dexamethasone were determined by HPLC (Shimadzu LC-20AT, Japan) using prednisolone (1 lg/mL) as the internal standard. The mobile phase was methanol–water (55:45, v/v) at a flow rate of 1.5 mL/min, running through 2 serial Onyx Monolithic C18
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When standard dexamethasone solutions ranged from 100 ng/mL to 20 lg/mL, the standard HPLC curve was f(x) = 830.9x (R2 = 0.9988). A summary of the real-time concentrations of group A and group B is shown in Fig. 3 (data are shown in Table 1). At the beginning of the experiment, the concentration in the capillary was very high, then decreased rapidly in the first 2 weeks, and subsequently decreased slowly in the 3rd week. From the 4th week onward, the concentration in both groups remained relatively stable. The concentration in group B was consistently higher than group A for 15 weeks (p \ 0.01), then the difference between the two groups became smaller with time. After the 16th week here was no difference in concentration between the groups (p [ 0.05). The cumulative amount released was determined with the equation: Q n ¼ C n Vt þ Vs
i¼n1 X i¼1
Cn1 :
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Fig. 3 Real-time concentrations of group A and group B. The concentration in group B was consistently higher than in group A for 15 weeks (p \ 0.01), though the difference between the two groups became smaller with time. After the 16th week there was no difference in concentration between the two groups (p [ 0.05)
Vt was the volume of the capillary, 0.16 mL in the present experiment, and Vs was the volume of the sample, 0.024 mL in the present experiment. The amount released in group B was greater than that of group A (Fig. 4). A summary of the cumulative percentage data is shown in Fig. 5 (data presented in Table 1). In contrast with the cumulative released amount, the percentage released in group A was greater than that in group B; e.g. until 24 weeks the amount of dexamethasone released in group A was 30.7 % and in group B was 10.3 %. In vivo experiment When standard dexamethasone solutions ranged from 50 ng/mL to 800 ng/mL, the standard HPLC curve was f(x) = 0.000838181x ? 0.00109879 (R2 = 0.9999678).
Table 1 Summary of the realtime concentration, cumulative amount released, and cumulative percentage released in the in vitro experiment
Time
Fig. 4 Cumulative amount released in group A and group B. The amount released in group B was higher than that of group A from the beginning to the end of the experiment (p \ 0.01)
Since 8 lL perilymph samples were diluted with 17 lL artificial perilymph, the concentration in perilymph was extrapolated from the concentration detected with HPLC (Cperilymph = Cn/8 9 25). The perilymph concentrations of dexamethasone are shown in Table 2 and Fig. 6. There was a burst release in both the 2 % DEXA group and the 10 % DEXA group. The peak concentration occurred 30 min after surgery in both groups, and then decreased rapidly until 3 h after surgery. From 3 h to 1 day the concentration in the 10 % DEXA group was significantly higher than in the 2 % DEXA group (p \ 0.01), after which the difference became non-significant. One week after surgery the concentration was 101.21 ± 34.04 ng/mL in the 2 % DEXA group, and 159 ± 74.64 ng/mL in the 10 % DEXA group (p [ 0.05). The elimination half-time of dexamethasone in perilymph was at an average of 34.3 min for the 2 % DEXA group and 39.0 min for the 10 % DEXA
Concentration (lg/mL)
Cumulative amount (lg)
Cumulative percentage (%)
Group A
Group A
Group A
Group B
Group B
Group B
1 day
4.05 ± 0.76
14.67 ± 3.90
0.65 ± 0.12
2.35 ± 0.62
2.71 ± 0.50
1.96 ± 0.52
2 days
4.61 ± 1.33
13.07 ± 4.09
0.84 ± 0.23
2.44 ± 0.74
3.50 ± 0.96
2.03 ± 0.62
3 days
4.66 ± 1.63
11.23 ± 3.20
0.95 ± 0.30
2.46 ± 0.66
3.96 ± 1.25
2.05 ± 0.55
1 week
3.75 ± 1.59
7.45 ± 1.73
1.22 ± 0.44
2.73 ± 0.58
5.08 ± 1.83
2.28 ± 0.48
4 weeks
1.78 ± 0.67
3.04 ± 0.72
2.25 ± 0.64
4.24 ± 0.45
9.38 ± 2.67
3.53 ± 0.38
8 weeks
1.74 ± 0.46
3.73 ± 0.31
3.42 ± 0.91
6.71 ± 0.50
14.25 ± 3.79
5.59 ± 0.42
12 weeks
1.36 ± 0.13
3.20 ± 0.88
4.52 ± 0.83
8.16 ± 0.70
18.83 ± 3.46
6.80 ± 0.58
16 weeks
2.22 ± 1.12
2.47 ± 0.57
5.48 ± 1.28
10.01 ± 0.96
22.83 ± 5.33
8.34 ± 0.80
20 weeks
1.15 ± 0.53
1.60 ± 0.83
6.24 ± 1.50
11.15 ± 1.10
26.00 ± 6.25
9.29 ± 0.92
24 weeks
1.43 ± 0.27
1.58 ± 0.85
7.37 ± 1.56
12.40 ± 1.12
30.71 ± 6.50
10.33 ± 0.93
Data are reported as mean ± standard deviation
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Fig. 5 Cumulative percentage released in group A and group B. The percentage released in group A was higher than that in group B (p \ 0.01)
Table 2 Real-time concentrations (ng/mL) in perilymph in vivo Time
2 % DEXA Group
10 % DEXA Group
10 min
395.46 ± 83.09
1583.17 ± 280.68
30 min
1373.37 ± 338.17
2444.43 ± 275.06
1.5 h
534.56 ± 22.02
1461.93 ± 49.98
3h
323.03 ± 24.71
7h
241.4 ± 107.89
773.43 ± 144.89 719.01 ± 238.89
1 day
104.25 ± 22.14
364.3 ± 145.19
1 week
101.21 ± 34.04
159 ± 74.64
Data are reported as mean ± standard deviation
group. No dexamethasone was detected in the two ears implanted with silicone rods containing no drug.
Discussion In vitro experimental model The recently reported study on guinea pigs demonstrates a faster flowing of CSF into perilymph than the previous references have reported [31, 32, 35]. However, no studies report on the real perilymph flow rate in human. The flow rate of 24 lL per day in human may be a lower estimation, whereas it would not affect the comparison of these dexamethasone-releasing silicones in vitro. The possible uptake of dexamethasone receptors in the wall of the perilymph was not taken into account, nor was transverse fluid diffusion (scala vestibuli–scala media–scala tympany) [36]. Farahmand Ghavi et al. [26] studied silicone impregnated with 0.25–2 % dexamethasone in vitro, in which the release medium was 1 mL and the whole medium was
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Fig. 6 Dexamethasone concentration–time curves in perilymph. The image in the top right is the detailed display of the curves within 7 h after implantation. The peak concentration occurred 30 min after surgery in both groups, and then decreased rapidly until 3 h after surgery. From 3 h to 1 day the perilymph concentration in the 10 % DEXA group was significantly higher than in the 2 % DEXA group (p \ 0.01), after which the difference became non-significant (p [ 0.05)
refreshed at each sampling. They found that the more dexamethasone that was incorporated, the more total amount of drug was released. The authors also reported that the cumulative percentage of drug released was the opposite: the cumulative percentage drug released in silicone impregnated with 0.25 % dexamethasone was higher than silicone with 2 % dexamethasone. Krenzlin et al. [27] found similar dexamethasone release kinetics with drugincorporated cochlear implants in vitro, in which the release medium was 10 and 1 mL was withdrawn for each sampling. The present experiment used silicone manufactured in the same way as by Farahmand Ghavi et al. [26], but the release medium was 160 lL, and only 24 lL medium were withdrawn and refreshed at each sampling. The results of our study are consistent with those of previous studies. In group A there was less total dexamethasone than in group B, and the total released dexamethasone in group A was less than in group B (Fig. 4), whereas the cumulative percentage of drug released was higher than in group B (Fig. 5). The derived drug diffusion equation can explain this phenomenon (monolithic solutions exhibiting the geometries of slabs) [27, 37, 38]: ! 1 Mt 8X 1 Dð2n þ 1Þ2 p2 t ¼1 2 exp ; p n¼0 ð2n þ 1Þ2 L2 M1 where Mt and M1 denote the absolute cumulative amounts of drug released at time t and infinity, respectively, and Mt/ M1 is the release percentage. In group A and group B the parameters of n (a dummy variable) and D (the ‘‘apparent’’
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diffusion coefficient of the drug within the polymeric system) are similar, whereas L (the thickness of the film) is different. Therefore, the release percentage of group A (200 lm in thickness) was higher than in group B (500 lm in thickness). Moreover, we found that the concentration of group B was higher than that of group A initially, and the difference decreased to the 16th week where no difference was noted. This indicated that the release volume of 160 lL restricted drug release from group B more than in group A. Therefore, concentrations in the two groups could be kept at a relative similar and stable level. While the remaining amount of group B was more than group A at the end of the in vitro experiment, it could be predicted that the duration of drug release in group B would be longer than in group A. Drug release from cochlear implant electrode arrays in vivo A number of studies have shown that samples taken from the basal turn of scala tympani are highly contaminated with CSF through the cochlear aqueduct [39, 40]. Salt et al. [34] developed the method of sampling perilymph from the cochlear apex, and found that the purity of perilymph samples can be increased using this method. Although the contamination of CSF from the cochlear aqueduct could not be absolutely avoided even if samples are taken from cochlear apex, the parallel samples at different time intervals would reduce this influence on result analysis. Therefore, the method of sampling from the cochlear apex was used in the present study, with three replicates of each group (n = 3) at the same time. Several studies have examined drug delivery into the cochlea via a cannula. Paasche et al. [19] opened the stylet channel at the tip of a Nucleus 24 Contour electrode and connected it to a micro pump at its proximal, thus using it as a drug delivery channel. The modified electrode was inserted into the scala tympani of human temporal bones, and the of contrast dye outside the electrode array was observed in the basal turn of the cochlea. A follow-up study by the same group showed concentrations comparable to those predicted by a computer model (http://www.oto. wustl.edu/cochlea) [20]. While temporal bone studies allowed investigation of distribution in vitro (ex vivo), important pharmacokinetic factors such as clearance and metabolism could not be taken into account. Most of the in vivo inner ear pharmacokinetic studies used extracochlear drug application to the round window membrane [41–43]. The longest observed duration among the inner ear pharmacokinetic studies was 28 days [41], using microimplant with beclomethasone. Most studies, however, investigated drug concentration in the cochlea for no longer than 6 h, because of the fast drug clearance from
the cochlea after a single application [41–43]. Parnes et al. [41] investigated dexamethasone concentration in scala tympani, scala vestibuli and endolymph after a single intratympanic application. The rapid decline of concentration in this study, however, could be explained by CSF contamination through multiple samples (4 9 10 lL) and leakage at the cochleostomy site [36]. The present study was the first to examine inner ear pharmacokinetics with a drug releasing cochlear implant for one week in animals. The silicone rod was inserted three mm into the basal scala tympani. The peak concentration occurred 30 min after implantation, which indicated the initial burst release during that period. The concentration then dropped rapidly from 30 min to 3 h after implantation, indicating a fast elimination of drug from the cochlear perilymph, e.g. through pharmacokinetic process like uptake of dexamethasone in cells, metabolism and the clearance to blood [36]. From 3 h to 1 week concentrations in the 2 % DEXA group and in the 10 % DEXA group decreased very slowly, indicating a balance of release and clearance for this period. The calculated elimination halftimes was at an average of 34.3 min for the 2 % DEXA group and 39.0 min for the 10 % DEXA group, respectively, which was somewhat longer than measured after intralabyrinthine application of dexamethasone solution (22.5 min) [44]. Although at the beginning the concentration in the 10 % DEXA group was higher than in the 2 % DEXA group, the difference ceased to exist by 1 week after implantation. Because the in vitro experiment in the present study indicated that silicone containing more dexamethasone will show a longer release period, it is suggested that the 10 % DEXA group would achieve a longer release period than the 2 % DEXA group in the in vivo experiment. Comparison of in vitro and in vivo experiments The present in vitro experiments studied two sizes of silicone films with the same drug loading (2 % dexamethasone), and the in vivo experiment studied the same shape of silicone rod loaded with 2 % dexamethasone or 10 % dexamethasone. In the in vitro study, drug metabolism and drug distribution routes (e.g. tissue binding) were not taken into account. Therefore, drug was released more slowly in the in vitro experiment, whereas, drug kinetics in both experiments were similar: (1) regardless of the drug amount the silicone contained, the concentration in the release medium became similar after a period of burst release; (2) balanced drug distribution could be achieved in vitro for 20 weeks and in vivo for 1 week at least; (3) silicone with a larger amount of drug predictably maintained a longer period of drug release. Although many studies have examined the effects of dexamethasone on the inner ear, no confirmed safe and
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effective drug concentration in the inner ear has been determined. In a previous study, we found that 2 % dexamethasone-loaded silicone electrodes could preserve hearing in guinea pigs after implantation [45]. Farhadi et al. [46] found that a 2 % dexamethasone-loaded silicone rod significantly reduced the infiltration with fibrocytes, macrophages, Foreign Body Giant Cells (FBGC) and lymphocytes infiltration post implantation. This result is consistent with the study of Takumi et al. [28], who showed that a 2 % dexamethasone-eluting electrode similar as that used in our study suppressed the expression of various genes associated with immune reaction and inflammation. Douchement’s et al. [47] performed a functional study in gerbils with electrode arrays loaded with 1 % dexamethasone and found long-term hearing preservation for one year at the 16 kHz region. Stathopoulos et al. [48] found a better spiral ganglion neuron survival three months after implantation with electrode arrays loaded with 20 % dexamethasone. The present study compared the pharmacokinetic properties of the 2 and 10 % dexamethasone-loaded cochlear implant silicone dummies. The drug contained in the silicone dummy loaded with 10 % dexamethasone is more than the previously investigated 1 and 2 % concentrations, and might prove even more beneficial with respect to safety and performance in the animal and the human. The results of this study demonstrated that electrode arrays loaded with 10 % dexamethasone achieve similar drug levels in the inner ear as electrode arrays with 2 % dexamethasone but release dexamethasone for a longer time period. An improved effect on the safety and performance of the cochlear implants will have to be investigated in pre-clinical studies using morphological and functional measures and finally in clinical studies. Acknowledgments We thank Dr. Heike Schneider from Institute of Clinical Chemistry and Phatobiochemistry, Klinikum rechts der Isar, for her kind assistance with the HPLC assay. We gratefully acknowledge Michael Todd for medical writing assistance following the preparation of a version of this manuscript. We thank Prof. Karsten Ma¨der (Halle/Saale) for comments on the manuscript. The authors express their sincere gratitude to the MED-EL Medical Electronics, Innsbruck, Austria for providing financial supports. Compliance with ethical standards Conflict of interest The study was supported by a research grant from MED-EL Medical Electronics, Innsbruck, Austria to the Technical University of Munich, Germany. The authors have no conflicts of interest to declare.
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the auditory system. New technology for severe hearing loss. ORL J Otorhinolaryngol Relat Spec 61:334–340 Kiefer J, Pok M, Adunka O, Stu¨rzebecher E, Baumgartner W, Schmidt M, Tillein J, Ye Q, Gstoettner W (2005) Combined electric and acoustic stimulation of the auditory system: results of a clinical study. Audiol Neurootol 10:134–144 Fayad J, Linthicum FH Jr, Otto SR, Galey FR, House WF (1991) Cochlear implants: histopathologic findings related to performance in 16 human temporal bones. Ann Otol Rhinol Laryngol 100:807–811 Barbara M, Mattioni A, Monini S, Chiappini I, Ronchetti F, Ballantyne D, Mancini P, Filipo R (2003) Delayed loss of residual hearing in Clarion cochlear implant users. J Laryngol Otol 117:850–853 Eshraghi AA, Yang N, Balkany TJ (2003) Comparative study of cochlear damage with three perimodiolar electrode designs. Laryngoscope 113:415–419 Shinomori Y, Spack DS, Jones DD, Kimura RS (2001) Volumetric and dimensional analysis of the guinea pig inner ear. Ann Otol Rhinol Laryngol 110:91–98 Eshraghi AA, Van de Water TR (2006) Cochlear implantation trauma and noise-induced hearing loss: apoptosis and therapeutic strategies. Anat Rec A Discov Mol Cell Evol Biol 288:473–481 Gstoettner WK, Helbig S, Maier N, Kiefer J, Radeloff A, Adunka OF (2006) Ipsilateral electric acoustic stimulation of the auditory system: results of long-term hearing preservation. Audiol Neurootol 11:49–56 Ye Q, Tillein J, Hartmann R, Gstoettner W, Kiefer J (2007) Application of a corticosteroid (Triamcinolon) protects inner ear function after surgical intervention. Ear Hear 28:361–369 Braun S, Ye Q, Radeloff A, Kiefer J, Gstoettner W, Tillein J (2011) Protection of inner ear function after cochlear implantation: compound action potential measurements after local application of glucocorticoids in the guinea pig cochlea. ORL 73:219–228 Sto¨ver T, Issing P, Graurock G, Erfurt P, ElBeltagy Y, Paasche G, Lenarz T (2005) Evaluation of the advance off-stylet insertion technique and the cochlear insertion tool in temporal bones. Otol Neurotol 26:1161–1170 Ma W, Gee K, Lim W, Chambers K, Angel JB, Kozlowski M, Kumar A (2004) Dexamethasone inhibits IL-12p40 production in lipopolysaccharide-stimulated human monocytic cells by downregulating the activity of c-Jun N-terminal kinase, the activation protein-1, and NF-kappa B transcription factors. J Immunol 172:318–330 Maeda K, Yoshida K, Ichimiya I, Suzuki M (2005) Dexamethasone inhibits tumor necrosis factor-alpha-induced cytokine secretion from spiral ligament fibrocytes. Hear Res 202:154–160 Eshraghi AA, Adil E, He J, Graves R, Balkany TJ, Van De Water TR (2007) Local dexamethasone therapy conserves in an animal model of electrode insertion trauma induced hearing loss. Otol Neurotol 28:842–849 ´ Leary SJ (2008) James DP, Eastwood H, Richardson RT, O Effects of dexamethasone on residual hearing in a guinea pig model of cochlear implantation. Audiol Neurotol 13:86–96 Kiefer J, Gstoettner W, Baumgartner W, Pok SM, Tillein J, Ye Q, von Ilberg C (2004) Conservation of low-frequency hearing in cochlear implantation. Acta Otolaryngol 124:272–280 De Ceulaer G, Johnson S, Yperman M, Daemers K, Offeciers FE, O’Donoghue GM, Govaerts PJ (2003) Long-term evaluation of the effect of intracochlear steroid deposition on electrode impedance in cochlear implant patients. Otol Neurotol 24:769–774 Shepherd RK, Xu J (2012) A multichannel scala tympani electrode array incorporating a drug delivery system for chronic intracochlear infusion. Hear Res 172:92–98
Eur Arch Otorhinolaryngol 19. Paasche G, Gibson P, Averbeck T, Becker H, Lenarz T, Sto¨ver T (2003) Technical report-modification of a cochlear implant electrode for drug delivery to the inner ear. Otol Neurotol 24:222–227 20. Paasche G, Bo¨gel L, Leinung M, Lenarz T, Sto¨ver T (2006) Substance distribution in a cochlea model using different pump rates for cochlear implant drug delivery electrode prototypes. Hear Res 212:74–82 21. Sto¨ver T, Paasche G, Lenarz T, Ripken T, Breitenfeld P, Lubatschowski H, Fabian T (2007) Development of a drug delivery device: using the femtosecond laser to modify cochlear implant electrodes. Cochlear Implants Int 8:38–52 22. Bohl A, Rohm HW, Ceschi P, Paasche G, Hahn A, Barcikowski S, Lenarz T, Sto¨ver T, Pau HW, Schmitz KP, Sternberg K (2012) Development of a specially tailored local drug delivery system for the prevention of fibrosis after insertion of cochlear implants into the inner ear. J Mater Sci Mater Med 23:2151–2162 23. Niedermeier K, Braun S, Fauser C, Kiefer J, Straubinger RK, Stark T (2012) A safety evaluation of dexamethasone-releasing cochlear implants: comparative study on the risk of otogenic meningitis after implantation. Acta Otolaryngol 132:1252–1260 24. Stathopoulos D, Chambers S, Adams L, Robins-Browne R, Miller C, Enke YL, Wei BP, O’Leary S, Cowan R, Newbold C (2015) Meningitis and a safe dexamethasone-eluting intracochlear electrode array. Cochlear Implants Int 16:201–207 25. Nguyen Y, Bernardeschi D, Kazmitcheff G, Miroir M, Vauchel T, Ferrary E, Sterkers O (2014) Effect of embedded dexamethasone in cochlear implant array on insertion forces in an artificial model of scala tympani. Otol Neurotol 36:354–358 26. Farahmand Ghavi F, Mirzadeh H, Imani M, Jolly C, Farhadi M (2010) Corticosteroid-releasing cochlear implant: a novel hybrid of biomaterial and drug delivery system. J Biomed Mater Res B Appl Biomater 94:388–398 27. Krenzlin S, Vincent C, Munzke L, Gnansia D, Siepmann J, Siepmann F (2012) Predictability of drug release from cochlear implants. J Control Release 159:60–68 28. Takumi Y, Nishio SY, Mugridge K, Oguchi T, Hashimoto S, Suzuki N, Iwasaki S, Jolly C, Usami S (2014) Gene expression pattern after insertion of dexamethasone-eluting electrode into the guinea pig cochlea. PLoS One 9:e110238 29. Igarashi M, Ohashi K, Ishii M (1986) Morphometric comparison of endolymphatic and perilymphatic spaces in human temporal bones. Acta Otolaryngol 101:161–164 30. Buckingham RA, Valvassori GE (2001) Inner ear fluid volumes and the resolving power of magnetic resonance imaging: can it differentiate endolymphatic structures? Ann Otol Rhinol Laryngol 110:113–117 31. Ohyama K, Salt AN, Thalmann R (1988) Volume flow rate of perilymph in the guinea-pig cochlea. Hear Res 35:119–129 32. Salt AN, Thalmann R (1988) Cochlear fluid dynamics. In: Jahn AF, Santos-Sacchi J (eds) Physiology of the ear. Raven Press, New York, pp 341–357 33. Henke J, Erhardt W (2002) Notfa¨lle unter Ana¨sthesie bei Kleinsa¨ugern. In: Erhardt W, Henke J, Lendl C (eds) NarkoseNotfa¨lle. Enke, Stuttgart, pp 96–218
34. Salt AN, Hale SA, Plonkte SK (2006) Perilymph sampling from the cochlear apex: a reliable method to obtain higher purity perilymph samples from scala tympani. J Neurosci Methods 153:121–129 35. Salt AN, Gill RM, Hartsock JJ (2015) Perilymph kinetics of FITC-dextran reveals homeostasis dominated by the cochlear aqueduct and cerebrospinal fluid. J Assoc Res Otolaryngol 16:357–371 36. Salt AN, Plontke SK (2005) Local inner-ear drug delivery and pharmacokinetics. Drug Discov Today 10:1299–1306 37. Crank J (1979) The mathematics of diffusion, 2nd edn. Clarendon Press, Oxford, p 50 38. Siepmann J, Siepmann F (2012) Modeling of diffusion controlled drug delivery. J Controll Release 161:351–362 39. Hara A, Salt AN, Thalmann R (1989) Perilymph composition in scala tympani of the cochlea: influence of cerebrospinal fluid. Hear Res 42:265–271 40. Salt AN, Kellner C, Hale S (2003) Contamination of perilymph sampled from the basal cochlear turn with cerebrospinal fluid. Hear Res 182:24–33 41. Parnes LS, Sun AH, Freeman DJ (1999) Corticosteroid pharmacokinetics in the inner ear fluids: an animal study followed by clinical application. Laryngoscope 109:1–17 42. Arnold W, Senn P, Hennig M, Michaelis C, Deingruber K, Scheler R, Steinhoff HJ, Riphagen F, Lamm K (2005) Novel slow- and fast-type drug release round-window microimplants for local drug application to the cochlea: an experimental study in guinea pigs. Audiol Neurootol 10:53–63 43. Mynatt R, Hale SA, Gill RM, Plontke SK, Salt AN (2006) Demonstration of a longitudinal concentration gradient along scala tympani by sequential sampling of perilymph from the cochlear apex. J Assoc Res Otolaryngol 7:182–193 44. Salt AN, Hartsock JJ, Gill RM, Piu F, Plontke SK (2012) Perilymph pharmacokinetics of markers and dexamethasone applied and sampled at the lateral semi-circular canal. J Assoc Res Otolaryngol 13:771–783 45. Liu Y, Jolly C, Braun S, Janssen T, Scherer E, Steinhoff J, Ebenhoch H, Lohner A, Stark T, Kiefer J (2015) Effects of a dexamethasone-releasing implant on cochleae: a functional, morphological and pharmacokinetic study. Hear Res 327:89–101 46. Farhadi M, Jalessi M, Salehian P, Ghavi FF, Emamjomeh H, Mirzadeh H, Imani M, Jolly C (2013) Dexamethasone eluting cochlear implant: histological study in animal model. Cochlear Implants Int 14:45–50 47. Douchement D, Terranti A, Lamblin J, Salleron J, Siepman F, Siepmann J, Vincent C (2015) Dexamethasone eluting electrodes for cochlear implantation: effect on residual hearing. Cochlear Implants Int 16:195–200 48. Stathopoulos D, Chambers S, Enke YL, Timbol G, Risi F, Miller C, Cowan R, Newbold C (2014) Development of a safe dexamethasone-eluting electrode array for cochlear implantation. Cochlear Implants Int 15:254–263
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OTOLOGY
In vitro and in vivo pharmacokinetic study of a dexamethasone-releasing silicone for cochlear implants Ya Liu1 • Claude Jolly2 • Susanne Braun3 • Thomas Stark4 • Elias Scherer4 Stefan K. Plontke5 • Jan Kiefer6
•
Received: 16 December 2014 / Accepted: 21 August 2015 Ó Springer-Verlag Berlin Heidelberg 2015
Abstract Cochlear implants have been widely used for patients with profound hearing loss and partial deafness. Residual low-frequency hearing, however, may deteriorate due to insertion trauma and tissue response around the electrode array. The present study investigated in vitro and in vivo release of dexamethasone from silicone used for cochlear implant electrode carriers. The in vitro experiment involved an apparatus simulating the inner ear fluid environment in humans. Release from two sizes of silicone films (200 lm 9 1 mm 9 10 mm and 500 lm 9 1 mm 9 10 mm), each loaded with 2 % dexamethasone, and was measured for 24 weeks. In the in vivo experiment, silicone rods loaded with 2 or 10 % dexamethasone, respectively, were implanted into the scala tympani of guinea pigs. Perilymph concentrations were measured during the first week after
& Jan Kiefer [email protected] Ya Liu [email protected] 1
Department of Otolaryngology, Head and Neck Surgery, Beijing Naval General Hospital, Beijing 100048, People’s Republic of China
2
Electrode Research Section, MED-EL Medical Electronics, Innsbruck, Austria
3
MED-EL Deutschland GmbH, Starnberg, Germany
4
Clinic for Otorhinolaryngology, Head- and Neck Surgery, Klinikum rechts der Isar, Technical University of Munich, Munich, Germany
5
Department of Otorhinolaryngology, Head and Neck Surgery, Martin Luther University Halle-Wittenberg, Halle (Saale), Germany
6
HNO-Zentrum, Neupfarrplatz 12/II, 93047 Regensburg, Germany
implantation. The results showed that dexamethasone was released from the silicone in a sustained manner. After a burst release, perilymph concentration was similar for silicone incorporated with 2 and 10 % dexamethasone, respectively. The similar pharmacokinetic profile was found in the in vitro experiment. The period of sustained drug delivery was maintained for 20 weeks in vitro and for 1 week in vivo. The results of the present study suggest that drugs like dexamethasone are released in a controlled manner from silicon electrode carriers of cochlear implants. Further studies will identify optimal release profiles for the use with cochlear implants to improve their safety and longterm performance. Keywords Cochlear implant Dexamethasone Drug delivery Inner ear Pharmacokinetics
Introduction Since the Food and Drug Administration (FDA) approved the House 3M cochlear implant system in 1984, cochlear implants have been widely used for patients with profound hearing loss. With the strategy of combined electric and acoustic stimulation (EAS) or hybrid stimulation in partial deafness, nerve fibres coding for high and mid frequencies can be electrically stimulated by the cochlear implant, while the residual low-frequency hearing can be aided by acoustic amplification [1, 2]. However, the insertion of a cochlear implant electrode may cause direct trauma or indirect early or delayed damage to the cochlea, which may lead to deterioration or loss of residual hearing [3–5]. The formation of scar tissue around the electrode after cochlear implantation is suggested to be one of the most important causes of increase in electrode impedance and loss of
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residual hearing after surgery [5, 6]. Physical disruption of the organ of Corti and molecular mechanisms of cell death induced by the electrode insertion trauma are other contributing factors [7]. Therefore, the minimization of electrode insertion trauma and reduction of scar tissue around the electrode are important for preserving residual hearing, which is necessary for successful electric-acoustic stimulation [8–10]. Methods should include the use of less traumatic implant electrodes, modification of surgical techniques [11], and pharmaceutical treatments with additional drug delivery or by the means of drug device combinations, e.g. through drug release from cochlear implant electrode carriers. The present study focused on a pharmaceutical strategy to mitigate electrode insertion trauma. The most frequently studied drugs are glucocorticoids because of their immune suppression and anti-inflammatory functions, such as inhibition of the expression of tumour necrosis factor alpha (TNF-alpha), nuclear transcription factor-jB (NF-jB), and some of the cell death signal cascades (JNK) [12, 13]. Dexamethasone is a long-acting glucocorticoid (biological half-life 36–72 h) that possesses a high anti-inflammatory potency and is stable in vitro. There are three methods for local glucocorticoid delivery in cochlear implant surgery: 1.
Single drug application to the round window membrane or cochleostomy.
Several animal [10, 14, 15] and human studies [16, 17] have examined the effects of a single glucocorticoid application in cochlear implant surgery, and have shown benefits in hearing, histology, and postoperative electrode impedance. 2.
Direct drug irrigation into the cochlea with a modified electrode array.
Several investigators modified the electrode array for sustained drug delivery into cochlea [18–21]. In most studies, the drug was delivered from a mini pump into the cochlea through or cannula or through the electrode array. This application allows variation of drug volume and delivery rate after implantation. However, the safety of the drug delivery system needs to be further investigated. 3.
Drug release from cochlear implant electrode carriers.
A few studies reported the use of an electrode array made of a biocompatible silicone in which a drug is incorporated to provide a long-term sustained release. Bohl et al. [22] demonstrated an in vitro biocompatibility study that sustained-release dexamethasone was non-toxic to cultured spiral ganglion cells. In an own study electrode dummies with 10 % dexamethasone were implanted into the inner era of guinea pigs through a cochleostomy. The histological result demonstrated the location of the
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electrode array in scala tympani, and found no increased risk of postoperative pneumococcal otogenic meningitis [23]. Stathopoulos et al. [24] performed a similar study with electrode dummies with 40 % dexamethasone in the rat and results consistent to ours. Nguyen et al. [25] found that compared with a drug-free electrode array, the insertion forces were not increased for a 0.4 mm diameter dexamethasone-eluting array for all insertion lengths. Further studies of dexamethasone-releasing electrodes addressed drug release profiles and material fabrication [26, 27]. The present study investigated dexamethasone-releasing silicone with in vitro and in vivo pharmacokinetic experiments. An in vitro apparatus was designed to simulate the inner ear fluid environment in human, and the in vivo experiments examined drug concentration in perilymph during one week after implantation.
Materials and methods Study design The aim of the present study was to investigate the pharmacokinetic aspects of a dexamethasone-releasing silicone for cochlear implants. The pharmacokinetic characterization of the device such as drug release and diffusion was investigated in an in vitro study using an apparatus to simulate the inner ear fluid environment in human. The in vitro study was followed by measurements of drug concentration in scala tympani perilymph of the guinea pig in vivo, comparing concentration time courses after implantation of silicone rods loaded with either 2 or 10 % dexamethasone. Dexamethasone-releasing silicone The dexamethasone-releasing silicone was manufactured as part of a research project with MED-EL GmbH (Innsbruck, Austria). The detailed preparation has been described previously by Farahmand Ghavi et al. [26]. Two or ten percent (weight of drug per weight of the final polymer electrode) pharmaceutical grade micronized dexamethasone (Aventis, Romainville, France) was homogeneously mixed with medical grade silicone, which is used in human cochlear implant electrode arrays. The in vitro study used two sizes of silicone films [200 lm 9 1 mm 9 10 mm (group A) and 500 lm 9 1 mm 9 10 mm (group B)] incorporated with 2 % dexamethasone. The in vivo experiment used silicone rods as an electrode analogue that contained no wires, and were similar to the electrode dummy described by Niedermeier et al. [23] and Takumi et al. [28]. The rods incorporating 2 or 10 %
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Fig. 1 Cross-section of the silicone rod used in the in vivo experiment. It was 6 mm long and 0.5 mm diameter. One end tapered to 0.5 mm. A length of 3 mm was inserted into the scala tympani of the guinea pigs
dexamethasone were adapted to a guinea pig’s anatomical dimensions, and were 6 mm long and 0.5 mm diameter. The rod was inserted into the scala tympani of the guinea pigs half its length, (i.e. for 3 mm). The intracochlear end of the rod was bevelled over a length of 0.5 mm (Fig. 1). In vitro release experiments Based on the data of the previous studies, the volume of human perilymph is 158.5–166.4 lL [29, 30]. Therefore, the capillary with a volume of 160 lL was used in the in vitro experiments, representing the human perilymph cavity. There are no studies on the perilymph flow rate in human, and the estimation has been made in the present in vitro experiment, based on the data in guinea pigs. The perilymph volume in the guinea pig is 16 lL and the flow of cerebrospinal fluid (CSF) entering perilymph is 2.304 lL per day [31, 32]. Therefore, the flow rate of human perilymph was taken as 24 lL per day, and 24 lL artificial perilymph was sampled in the present in vitro experiments. A vertical sectional view of the in vitro release apparatus is demonstrated in Fig. 2. The silicone film with 2 % dexamethasone (shadowed area in the figure) was kept in a glass capillary (outer diameter: 6.0 ± 0.2 mm; inner diameter: 2.7 ± 0.1 mm; length: 28.000 ± 0.5 m; Duran Group, Germany). The volume of the capillary was 160 lL, and was filled with artificial perilymph (an aqueous solution of 1.3 mmol/L calcium chloride, 1.8 mmol/L magnesium chloride, 5.4 mmol/L potassium chloride, 137 mmol/L sodium chloride, 5 mmol/L glucose, and 5 mmol/L HEPES). At the bottom of the capillary there was a sample needle, shortened from a HamiltonÒ 7770-01 (Germany) needle, that could be connected to a microsyringe (Hamilton 705 RN 50 lL, Germany). At the top of the capillary there was a small tube connected to a chamber, which was also filled with artificial perilymph. All components of the in vitro release apparatus were sealed with Kwik-CastTM (WPI, Los Angeles, USA) and TurboKITT (Germany). The in vitro apparatus were kept
Fig. 2 Drawing of the in vitro release experimental set-up (cross section). The silicone film (shadowed area) incorporated with 2 % dexamethasone was maintained vertically in a glass capillary containing a volume of 160 lL artificial perilymph. At the bottom of the capillary there is a sample needle. At the top of the capillary there is a small tube connected to a chamber filled with artificial perilymph
vertical. The level of the fluid in the chamber was kept higher than the top of the capillary. Therefore, when sampling fluid from the bottom of the capillary the same volume of fluid would flow from the chamber to the capillary. Five replicates of each group (n = 5) were studied at the same time. During the experiment the apparatus was kept in a 37 °C water bath, except during sampling from the capillary. Sampling was performed every day at the same time. The sample volume was 24 lL, and it was continued for 168 days. Animals for in vivo experiment The in vivo experiment consisted of 30 healthy pigmented guinea pigs (Charles River). Silicone rods containing 10 % dexamethasone (10 % DEXA group, 21 ears) or 2 % dexamethasone (2 % DEXA group, 21 ears) were implanted into the cochlea. Two ears were implanted with silicone rods containing no drug and were used as blank controls. For the reduction of animal experiments, some animals were implanted bilaterally using silicon rods with the same dexamethasone concentration.
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Surgery and anaesthesia Silicone rod implantation was performed under general anaesthesia. Guinea pig body temperature was maintained at about 38.5 °C using a Medax heater (Medax GmbH und Co. KG, Neumu¨nster, Germany). Animals were anesthetised using a mixture of medetomidine (200 lg/kg), fentanyl (25 lg/kg) and midazolam (1 mg/kg). The level of anaesthesia was maintained by regular intramuscular injections of the same mixture, if necessary [33]. The animals received subcutaneous injections of 1 % mepivacaine hydrochloride (Curasan AG, Kleinostheim, Germany) before their skin and muscle tissues were cut. A retroauricular incision was made, and after location of the bulla a small hole was drilled and the opening was enlarged with tweezers to expose the round window membrane niche and the promontory. Using a 0.8 mm diamond drill, a cochleostomy was performed in the basal turn of the cochlea about 1 mm above the ridge of the round window membrane niche. The diameter of the cochleostomy was chosen as 0.8 mm to allow optimal placement of the silicone rod. Since it was not a functional study, the issue of hearing preservation had not been taken into account. After drilling the cochleostomy a silicone rod was inserted three mm into scala tympani. Immediately after implantation, a soft tissue was placed around the rod to seal the cochleostomy. The bulla was sealed with bone wax. All operations were performed by the same surgeon (YK).
(100 9 46 mm) columns. The detection wavelength was 254 nm. Before measurement all samples were centrifuged at 1000 rpm for 10 min. The supernatant was carefully removed and mixed with the same volume of internal standard for sample injection. Compliance and ethics The study was performed in the lab of Clinic for Otorhinolaryngology, Head- and Neck Surgery, Klinikum rechts der Isar, Technical University of Munich, Munich, Germany. It was in compliance with the PHS Policy on Humane Care and Use of Laboratory Animals, the NIH Guide for the Care and Use of Laboratory Animals, and the Animal Welfare Act. The protocol (no. 6-075/06) permitting the use of animals was approved by the Government of Oberbayern (Munich, Germany). Statistical analyses Perilymph concentrations were compared using group as the test variable (in vitro experiment: group A and group B; in vivo experiment: group 2 % DEXA and group 10 % DEXA). Statistical analyses were performed with SPSS 18.0 (IBM SPSS Statistics). Differences between groups were compared using the nonparametric Mann–Whitney U test for two independent samples (1-tailed). A value of p B 0.05 was considered to indicate statistical significance.
Perilymph sampling from the cochlear apex
Results
Perilymph was sampled according to the apical sampling procedure described by Salt et al. [34]. Eight microliters of perilymph were sampled from the cochlear apex, consisting of approximately 4.5–5 lL of scala tympani perilymph and 3 lL of cerebrospinal fluid. Samples were obtained at 10 min, 30 min, 1.5 h, 3 h, 7 h, 1 day, and 1 week after surgery (each n = 3). Each implanted cochlea was sampled only once. The samples were diluted into 17 lL of artificial perilymph and stored at -20 °C. Perilymph from the cochleae implanted with a blank silicone rod was sampled 30 min after surgery. After sampling, animals were sacrificed with an intracardiac injection of pentobarbital (40 mg/kg, Meral GmbH, Hallbergmoos, Germany).
In vitro release
High performance liquid chromatography (HPLC) Concentrations of dexamethasone were determined by HPLC (Shimadzu LC-20AT, Japan) using prednisolone (1 lg/mL) as the internal standard. The mobile phase was methanol–water (55:45, v/v) at a flow rate of 1.5 mL/min, running through 2 serial Onyx Monolithic C18
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When standard dexamethasone solutions ranged from 100 ng/mL to 20 lg/mL, the standard HPLC curve was f(x) = 830.9x (R2 = 0.9988). A summary of the real-time concentrations of group A and group B is shown in Fig. 3 (data are shown in Table 1). At the beginning of the experiment, the concentration in the capillary was very high, then decreased rapidly in the first 2 weeks, and subsequently decreased slowly in the 3rd week. From the 4th week onward, the concentration in both groups remained relatively stable. The concentration in group B was consistently higher than group A for 15 weeks (p \ 0.01), then the difference between the two groups became smaller with time. After the 16th week here was no difference in concentration between the groups (p [ 0.05). The cumulative amount released was determined with the equation: Q n ¼ C n Vt þ Vs
i¼n1 X i¼1
Cn1 :
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Fig. 3 Real-time concentrations of group A and group B. The concentration in group B was consistently higher than in group A for 15 weeks (p \ 0.01), though the difference between the two groups became smaller with time. After the 16th week there was no difference in concentration between the two groups (p [ 0.05)
Vt was the volume of the capillary, 0.16 mL in the present experiment, and Vs was the volume of the sample, 0.024 mL in the present experiment. The amount released in group B was greater than that of group A (Fig. 4). A summary of the cumulative percentage data is shown in Fig. 5 (data presented in Table 1). In contrast with the cumulative released amount, the percentage released in group A was greater than that in group B; e.g. until 24 weeks the amount of dexamethasone released in group A was 30.7 % and in group B was 10.3 %. In vivo experiment When standard dexamethasone solutions ranged from 50 ng/mL to 800 ng/mL, the standard HPLC curve was f(x) = 0.000838181x ? 0.00109879 (R2 = 0.9999678).
Table 1 Summary of the realtime concentration, cumulative amount released, and cumulative percentage released in the in vitro experiment
Time
Fig. 4 Cumulative amount released in group A and group B. The amount released in group B was higher than that of group A from the beginning to the end of the experiment (p \ 0.01)
Since 8 lL perilymph samples were diluted with 17 lL artificial perilymph, the concentration in perilymph was extrapolated from the concentration detected with HPLC (Cperilymph = Cn/8 9 25). The perilymph concentrations of dexamethasone are shown in Table 2 and Fig. 6. There was a burst release in both the 2 % DEXA group and the 10 % DEXA group. The peak concentration occurred 30 min after surgery in both groups, and then decreased rapidly until 3 h after surgery. From 3 h to 1 day the concentration in the 10 % DEXA group was significantly higher than in the 2 % DEXA group (p \ 0.01), after which the difference became non-significant. One week after surgery the concentration was 101.21 ± 34.04 ng/mL in the 2 % DEXA group, and 159 ± 74.64 ng/mL in the 10 % DEXA group (p [ 0.05). The elimination half-time of dexamethasone in perilymph was at an average of 34.3 min for the 2 % DEXA group and 39.0 min for the 10 % DEXA
Concentration (lg/mL)
Cumulative amount (lg)
Cumulative percentage (%)
Group A
Group A
Group A
Group B
Group B
Group B
1 day
4.05 ± 0.76
14.67 ± 3.90
0.65 ± 0.12
2.35 ± 0.62
2.71 ± 0.50
1.96 ± 0.52
2 days
4.61 ± 1.33
13.07 ± 4.09
0.84 ± 0.23
2.44 ± 0.74
3.50 ± 0.96
2.03 ± 0.62
3 days
4.66 ± 1.63
11.23 ± 3.20
0.95 ± 0.30
2.46 ± 0.66
3.96 ± 1.25
2.05 ± 0.55
1 week
3.75 ± 1.59
7.45 ± 1.73
1.22 ± 0.44
2.73 ± 0.58
5.08 ± 1.83
2.28 ± 0.48
4 weeks
1.78 ± 0.67
3.04 ± 0.72
2.25 ± 0.64
4.24 ± 0.45
9.38 ± 2.67
3.53 ± 0.38
8 weeks
1.74 ± 0.46
3.73 ± 0.31
3.42 ± 0.91
6.71 ± 0.50
14.25 ± 3.79
5.59 ± 0.42
12 weeks
1.36 ± 0.13
3.20 ± 0.88
4.52 ± 0.83
8.16 ± 0.70
18.83 ± 3.46
6.80 ± 0.58
16 weeks
2.22 ± 1.12
2.47 ± 0.57
5.48 ± 1.28
10.01 ± 0.96
22.83 ± 5.33
8.34 ± 0.80
20 weeks
1.15 ± 0.53
1.60 ± 0.83
6.24 ± 1.50
11.15 ± 1.10
26.00 ± 6.25
9.29 ± 0.92
24 weeks
1.43 ± 0.27
1.58 ± 0.85
7.37 ± 1.56
12.40 ± 1.12
30.71 ± 6.50
10.33 ± 0.93
Data are reported as mean ± standard deviation
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Fig. 5 Cumulative percentage released in group A and group B. The percentage released in group A was higher than that in group B (p \ 0.01)
Table 2 Real-time concentrations (ng/mL) in perilymph in vivo Time
2 % DEXA Group
10 % DEXA Group
10 min
395.46 ± 83.09
1583.17 ± 280.68
30 min
1373.37 ± 338.17
2444.43 ± 275.06
1.5 h
534.56 ± 22.02
1461.93 ± 49.98
3h
323.03 ± 24.71
7h
241.4 ± 107.89
773.43 ± 144.89 719.01 ± 238.89
1 day
104.25 ± 22.14
364.3 ± 145.19
1 week
101.21 ± 34.04
159 ± 74.64
Data are reported as mean ± standard deviation
group. No dexamethasone was detected in the two ears implanted with silicone rods containing no drug.
Discussion In vitro experimental model The recently reported study on guinea pigs demonstrates a faster flowing of CSF into perilymph than the previous references have reported [31, 32, 35]. However, no studies report on the real perilymph flow rate in human. The flow rate of 24 lL per day in human may be a lower estimation, whereas it would not affect the comparison of these dexamethasone-releasing silicones in vitro. The possible uptake of dexamethasone receptors in the wall of the perilymph was not taken into account, nor was transverse fluid diffusion (scala vestibuli–scala media–scala tympany) [36]. Farahmand Ghavi et al. [26] studied silicone impregnated with 0.25–2 % dexamethasone in vitro, in which the release medium was 1 mL and the whole medium was
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Fig. 6 Dexamethasone concentration–time curves in perilymph. The image in the top right is the detailed display of the curves within 7 h after implantation. The peak concentration occurred 30 min after surgery in both groups, and then decreased rapidly until 3 h after surgery. From 3 h to 1 day the perilymph concentration in the 10 % DEXA group was significantly higher than in the 2 % DEXA group (p \ 0.01), after which the difference became non-significant (p [ 0.05)
refreshed at each sampling. They found that the more dexamethasone that was incorporated, the more total amount of drug was released. The authors also reported that the cumulative percentage of drug released was the opposite: the cumulative percentage drug released in silicone impregnated with 0.25 % dexamethasone was higher than silicone with 2 % dexamethasone. Krenzlin et al. [27] found similar dexamethasone release kinetics with drugincorporated cochlear implants in vitro, in which the release medium was 10 and 1 mL was withdrawn for each sampling. The present experiment used silicone manufactured in the same way as by Farahmand Ghavi et al. [26], but the release medium was 160 lL, and only 24 lL medium were withdrawn and refreshed at each sampling. The results of our study are consistent with those of previous studies. In group A there was less total dexamethasone than in group B, and the total released dexamethasone in group A was less than in group B (Fig. 4), whereas the cumulative percentage of drug released was higher than in group B (Fig. 5). The derived drug diffusion equation can explain this phenomenon (monolithic solutions exhibiting the geometries of slabs) [27, 37, 38]: ! 1 Mt 8X 1 Dð2n þ 1Þ2 p2 t ¼1 2 exp ; p n¼0 ð2n þ 1Þ2 L2 M1 where Mt and M1 denote the absolute cumulative amounts of drug released at time t and infinity, respectively, and Mt/ M1 is the release percentage. In group A and group B the parameters of n (a dummy variable) and D (the ‘‘apparent’’
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diffusion coefficient of the drug within the polymeric system) are similar, whereas L (the thickness of the film) is different. Therefore, the release percentage of group A (200 lm in thickness) was higher than in group B (500 lm in thickness). Moreover, we found that the concentration of group B was higher than that of group A initially, and the difference decreased to the 16th week where no difference was noted. This indicated that the release volume of 160 lL restricted drug release from group B more than in group A. Therefore, concentrations in the two groups could be kept at a relative similar and stable level. While the remaining amount of group B was more than group A at the end of the in vitro experiment, it could be predicted that the duration of drug release in group B would be longer than in group A. Drug release from cochlear implant electrode arrays in vivo A number of studies have shown that samples taken from the basal turn of scala tympani are highly contaminated with CSF through the cochlear aqueduct [39, 40]. Salt et al. [34] developed the method of sampling perilymph from the cochlear apex, and found that the purity of perilymph samples can be increased using this method. Although the contamination of CSF from the cochlear aqueduct could not be absolutely avoided even if samples are taken from cochlear apex, the parallel samples at different time intervals would reduce this influence on result analysis. Therefore, the method of sampling from the cochlear apex was used in the present study, with three replicates of each group (n = 3) at the same time. Several studies have examined drug delivery into the cochlea via a cannula. Paasche et al. [19] opened the stylet channel at the tip of a Nucleus 24 Contour electrode and connected it to a micro pump at its proximal, thus using it as a drug delivery channel. The modified electrode was inserted into the scala tympani of human temporal bones, and the of contrast dye outside the electrode array was observed in the basal turn of the cochlea. A follow-up study by the same group showed concentrations comparable to those predicted by a computer model (http://www.oto. wustl.edu/cochlea) [20]. While temporal bone studies allowed investigation of distribution in vitro (ex vivo), important pharmacokinetic factors such as clearance and metabolism could not be taken into account. Most of the in vivo inner ear pharmacokinetic studies used extracochlear drug application to the round window membrane [41–43]. The longest observed duration among the inner ear pharmacokinetic studies was 28 days [41], using microimplant with beclomethasone. Most studies, however, investigated drug concentration in the cochlea for no longer than 6 h, because of the fast drug clearance from
the cochlea after a single application [41–43]. Parnes et al. [41] investigated dexamethasone concentration in scala tympani, scala vestibuli and endolymph after a single intratympanic application. The rapid decline of concentration in this study, however, could be explained by CSF contamination through multiple samples (4 9 10 lL) and leakage at the cochleostomy site [36]. The present study was the first to examine inner ear pharmacokinetics with a drug releasing cochlear implant for one week in animals. The silicone rod was inserted three mm into the basal scala tympani. The peak concentration occurred 30 min after implantation, which indicated the initial burst release during that period. The concentration then dropped rapidly from 30 min to 3 h after implantation, indicating a fast elimination of drug from the cochlear perilymph, e.g. through pharmacokinetic process like uptake of dexamethasone in cells, metabolism and the clearance to blood [36]. From 3 h to 1 week concentrations in the 2 % DEXA group and in the 10 % DEXA group decreased very slowly, indicating a balance of release and clearance for this period. The calculated elimination halftimes was at an average of 34.3 min for the 2 % DEXA group and 39.0 min for the 10 % DEXA group, respectively, which was somewhat longer than measured after intralabyrinthine application of dexamethasone solution (22.5 min) [44]. Although at the beginning the concentration in the 10 % DEXA group was higher than in the 2 % DEXA group, the difference ceased to exist by 1 week after implantation. Because the in vitro experiment in the present study indicated that silicone containing more dexamethasone will show a longer release period, it is suggested that the 10 % DEXA group would achieve a longer release period than the 2 % DEXA group in the in vivo experiment. Comparison of in vitro and in vivo experiments The present in vitro experiments studied two sizes of silicone films with the same drug loading (2 % dexamethasone), and the in vivo experiment studied the same shape of silicone rod loaded with 2 % dexamethasone or 10 % dexamethasone. In the in vitro study, drug metabolism and drug distribution routes (e.g. tissue binding) were not taken into account. Therefore, drug was released more slowly in the in vitro experiment, whereas, drug kinetics in both experiments were similar: (1) regardless of the drug amount the silicone contained, the concentration in the release medium became similar after a period of burst release; (2) balanced drug distribution could be achieved in vitro for 20 weeks and in vivo for 1 week at least; (3) silicone with a larger amount of drug predictably maintained a longer period of drug release. Although many studies have examined the effects of dexamethasone on the inner ear, no confirmed safe and
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effective drug concentration in the inner ear has been determined. In a previous study, we found that 2 % dexamethasone-loaded silicone electrodes could preserve hearing in guinea pigs after implantation [45]. Farhadi et al. [46] found that a 2 % dexamethasone-loaded silicone rod significantly reduced the infiltration with fibrocytes, macrophages, Foreign Body Giant Cells (FBGC) and lymphocytes infiltration post implantation. This result is consistent with the study of Takumi et al. [28], who showed that a 2 % dexamethasone-eluting electrode similar as that used in our study suppressed the expression of various genes associated with immune reaction and inflammation. Douchement’s et al. [47] performed a functional study in gerbils with electrode arrays loaded with 1 % dexamethasone and found long-term hearing preservation for one year at the 16 kHz region. Stathopoulos et al. [48] found a better spiral ganglion neuron survival three months after implantation with electrode arrays loaded with 20 % dexamethasone. The present study compared the pharmacokinetic properties of the 2 and 10 % dexamethasone-loaded cochlear implant silicone dummies. The drug contained in the silicone dummy loaded with 10 % dexamethasone is more than the previously investigated 1 and 2 % concentrations, and might prove even more beneficial with respect to safety and performance in the animal and the human. The results of this study demonstrated that electrode arrays loaded with 10 % dexamethasone achieve similar drug levels in the inner ear as electrode arrays with 2 % dexamethasone but release dexamethasone for a longer time period. An improved effect on the safety and performance of the cochlear implants will have to be investigated in pre-clinical studies using morphological and functional measures and finally in clinical studies. Acknowledgments We thank Dr. Heike Schneider from Institute of Clinical Chemistry and Phatobiochemistry, Klinikum rechts der Isar, for her kind assistance with the HPLC assay. We gratefully acknowledge Michael Todd for medical writing assistance following the preparation of a version of this manuscript. We thank Prof. Karsten Ma¨der (Halle/Saale) for comments on the manuscript. The authors express their sincere gratitude to the MED-EL Medical Electronics, Innsbruck, Austria for providing financial supports. Compliance with ethical standards Conflict of interest The study was supported by a research grant from MED-EL Medical Electronics, Innsbruck, Austria to the Technical University of Munich, Germany. The authors have no conflicts of interest to declare.
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