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|Year : 2001
: 3 | Issue : 11 | Page
|Absence of hair cell protection by exogenous FGF-1 and FGF-2 delivered to guinea pig cochlea in vivo
Tatsuya Yamasoba1, Richard A Altshuler2, Yehoash Raphael2, Amy L Miller2, Fumi Shoji3, Josef M Miller2
1 Kresge Hearing Research Institute, The University of Michigan, Ann Arbor, Michigan USA; Department of Otolaryngology, University of Tokyo, Tokyo, Japan
2 Kresge Hearing Research Institute, The University of Michigan, Ann Arbor, Michigan USA
3 Kresge Hearing Research Institute, The University of Michigan, Ann Arbor, Michigan USA; Department of Otolaryngology, University of Tohoku, Sendai, Japan
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Recent findings that glial cell line-derived neurotrophic factor (GDNF), neurotrophin-3 (NT3), and transforming growth factor α can protect the auditory hair cells from acoustic trauma or aminoglycoside ototoxicity in vivo raise the question of whether other neurotrophic factors can also protect the hair cells in vivo. Fibroblast growth factor-2 (FGF-2) can protect hair cells from neomycin ototoxicity in vitro, and in vivo study has shown upregulation of FGF receptor3 in the cochlea following noise exposure, suggesting that some FGF family members might play a role in protection or repair of the cochlea from damage. We therefore examined if FGF1 and FGF-2 chronically delivered to the cochlea prior to noise overstimulation can attenuate noise-induced hair cell damage in vivo under conditions in which GDNF and NT-3 were effective. Pigmented female guinea pigs underwent left scala tympani implantation of a microcannula attached to an osmotic pump filled with artificial perilymph only or containing FGFs (10 or 1 µµg/ml FGF-1 or 10 µµg/ml FGF-2). They were exposed to noise (4 kHz octave band, 115 dB SPL, 5 hr) 4 days after surgery. Threshold shifts 10 days postexposure were essentially equivalent at all frequencies tested across different treatment groups. No significant difference in threshold shifts was observed between the treated and untreated ears in any of the groups. The extent of hair cell damage was also comparable among the different treatment groups. These findings indicate that exogenous FGF-1 or FGF-2 does not influence noiseinduced hair cell damage under the experimental conditions used in this study, suggesting that these FGFs are not good candidates as auditory hair cell protectors in vivo.
Keywords: acoustic trauma, noise-induced hearing loss, neurotrophic factor, acidic fibroblast growth factor, basic fibroblast growth factor
|How to cite this article:|
Yamasoba T, Altshuler RA, Raphael Y, Miller AL, Shoji F, Miller JM. Absence of hair cell protection by exogenous FGF-1 and FGF-2 delivered to guinea pig cochlea in vivo. Noise Health 2001;3:65-78
|How to cite this URL:|
Yamasoba T, Altshuler RA, Raphael Y, Miller AL, Shoji F, Miller JM. Absence of hair cell protection by exogenous FGF-1 and FGF-2 delivered to guinea pig cochlea in vivo. Noise Health [serial online] 2001 [cited 2020 Jun 4];3:65-78. Available from: http://www.noiseandhealth.org/text.asp?2001/3/11/65/31764
| Introduction|| |
Fibroblast growth factors (FGFs) form a family of at least 17 structurally-related proteins and display a myriad of biological activities, including embryonic development, angiogenesis, mitogenesis, wound healing, tumorigenesis, and neurotrophic properties (Basilico and Moscatelli, 1992; Johnson and Williams, 1993; Muenke and Schell, 1995; Martin, 1998, Xu et al., 1999). Three classes of receptors are known to bind members of the FGF family. These include a family of four transmembrane tyrosine kinase receptors (FGF receptors 1-4 [FGFRs 1-4])(reviewed by Basilico and Moscatelli, 1992; Johnson and Williams, 1993; Ornitz et al., 1996, Xu et al., 1999), a cysteine-rich transmembrane protein (Zuber et al., 1997), and heparin or heparin sulphate proteoglycans (Moscatelli, 1987). Cellular responses to FGF are mediated mainly through transmembrane tyrosine kinase receptors. Only the FGFRs can transmit an intracellular signal upon binding FGF, although heparin or heparin sulphate proteoglycans serve as obligatory co-factors, which are required for FGFs to bind to and activate the tyrosine kinase receptors (Yayon et al., 1991). The function of cysteine-rich receptors is not well known, but a recent study (Zuber et al., 1997) suggests an involvement of this receptor in intracellular FGF trafficking and the regulation of cellular responses to FGFs. Targeted gene disruption of FGFs in mice results in phenotypes ranging from excessive hair growth to early embryonic lethality (Hebert et al., 1994). A study using embryos carrying different combinations of alleles of FGF-8 shows that FGF-8 gene function is required during gastrulation and cardiac, craniofacial, forebrain, midbrain, and cerebellar development (Meyers et al., 1998). Targeted disruption of FGFR1 or FGFR2 results in early embryonic lethality (reviewed by Muenke and Schell, 1995). Interestingly, targeted disruption of FGFR3 results in skeletal and inner ear defects with an apparent arrest in the development of the cochlea, specifically the morphologically distinct pillar cells (Colvin et al., 1996).
Several FGF members have been reported to be present in the peripheral auditory system. FGF1(previously called as acidic FGF) mRNA has been shown to be expressed in the developing and adult rat cochlea (Luo et al., 1993). In the adult rat cochlea, FGF-1 mRNA is expressed in the neuronal somas in the spiral ganglion, the region of nerve terminals beneath the inner hair cells (IHCs), the efferent nerve terminals beneath the outer hair cells (OHCs), and the nonsensory epithelium in the stria vascularis, but it is not detected in the IHCs or OHCs (Pirvola et al., 1995). Immunocytochemical study has shown the presence of FGF-2 (previously called as basic FGF) in the supporting cells, but not in the hair cells or the hyaline cells in the basilar papillae of 5-day-old chicken (Lee and Cotanche, 1996). FGF-2 mRNA has not been detected in the rat cochlea at any age (Luo et al., 1993). FGF-3 (previously called as protooncogene int-2) has been reported to be expressed in both the sensory hair cells and supporting cells in the cochlea, the vestibule, and the semicircular canal in the developing mouse (Wilkinson et al., 1989). FGF-8 mRNA has recently been shown to be expressed in the IHCs in the developing and adult rat (Pirvola et al., 1998).
Several FGFRs have been observed in both the avian and mammalian inner ear. In the chick basilar papilla, it has been shown that FGFRs are concentrated at the bases of the streocilia on the hair cells with faint localization of the receptor on the apical surfaces of the supporting cells and that they are expressed on the expanded apical surfaces of the supporting cells following a 48 h noise exposure (Lee and Cotanche, 1996). Pickles and van Heumen (1997) have shown that FGFRs 1-3 are expressed in the otocyst, whole cochlea, cochlear nerve ganglion, and cochlear and vestibular sensory epithelium. A recent study (Bermingham-McDonogh et al., 1998) has demonstrated that, in developing chick cochlea, FGFR3 is initially expressed throughout the developing basilar papilla and later is confined to the supporting cell layer, while in the mature basilar papilla, FGFR3 is expressed in most supporting cells, but not in the hair cells. In mammals, FGFR1 has been detected in rat utricular epithelial cells (Saffer et al., 1996), while only FGFR3 has so far been detected in the cochlea. Peters et al. (1993) have shown FGFR3 expression in the differentiating IHCs and OHCs, but not in the vestibule, during mouse development. In adult rats, FGFR3 mRNA is expressed in the Deiters and pillar cells and, at low levels, in the limbus and lateral wall of the cochlea; FGFR3 transcripts are present at an extremely low level in the OHCs and absent in the IHCs (Pirvola et al., 1995). FGFR3 is known to be preferentially activated by FGF1, FGF-2, FGF-4, FGF-8, and FGF-9 (Ornitz and Leder, 1992; Hecht et al., 1995; Macarthur et al., 1995; Ornitz et al., 1996).
Loss of the auditory hair cell eventually leads to degeneration of the auditory neurons. Certain types of neurotrophic factors, such as brainderived neurotrophic factor (BDNF), neurotrophin-3 (NT-3), NT-4/5, and glial cell line-derived neurotrophic factor (GDNF), can significantly enhance survival of spiral ganglion cells both in vitro and in vivo after hair cell damage (Lefebvre et al., 1994; Zheng et al., 1995; Ernfors et al., 1996; Staecker et al., 1996; Zheng and Gao, 1996; Miller et al., 1997; Gabaizadeh et al., 1997; Ylikoski et al., 1998). Although a certain degree of hair cell regeneration or repair has been reported in mammalian inner ear structures, specifically in the vestibule (Li et al., 1995; Rubel et al., 1995), hearing loss resulting from hair cell death is virtually irreversible in mammals. Therefore, it is of value to identify neurotrophic or growth factors that may protect the hair cells from environmental stress factors that cause cell death, such as ototoxic drugs and noise. It has been shown that a combination of transforming growth factor a (TGF a) with and without retinoic acid can protect the hair cells both in vitro and in vivo from neomycin ototoxicity (Malgrange et al., 1994; Staecker et al., 1997). Also, recent in vivo studies have demonstrated that GDNF can attenuate hearing loss and hair cell damage induced by gentamicin ototoxicity (Park et al., 1998) or acoustic trauma (Shoji et al., 2000a) and that NT-3 but not BDNF can attenuate noise-induced hearing loss and hair cell damage (Shoji et al., 2000b).
FGF family members are considered another possible candidate for hair cell protectors (Schimmang and Represa, 1997) based upon the following findings: 1) FGF-2 can protect the neonatal OHCs from neomycin ototoxicity in vitro (Low et al., 1996), 2) noise exposure leads to upregulation of FGFR3 in the cochlea (Pirvola et al., 1995), and, 3) mice lacking FGFR3 are deaf due to incomplete differentiation of the sensory epithelia (Colvin et al., 1996). Therefore, in this study, we examined if FGF-1 and FGF-2 can protect the hair cells from acoustic trauma in vivo. We exposed animals to 115 dB SPL 4 kHz octave band noise for 5 hours. At this exposure level, pre-treatments with GDNF, NT-3, an iron chelator (deferoxamine mesylate), and a free radical scavenger (mannitol) were all able to significantly attenuate hair cell damage from acoustic trauma (Shoji et al., 2000a, 2000b; Yamasoba et al., 1999).
| Materials and Methods|| |
Twenty-four pigmented female guinea pigs with normal Preyer's reflex, weighing between 250 and 300 g, were used in this study. Only female guinea pigs were used, since reactive oxygen species (ROS) generation is thought to be involved in noise-induced cochlear damage (Hu et al., 1997; Yamasoba et al., 1998a,1998b) and sex differences are associated with differing abilities to detoxify ROS (Julicher et al., 1984).
Animals were randomly assigned to one of four groups (n=6 each). All animals underwent implantation of a microcannula attached to an osmotic pump filled with artificial perilymph (AP) containing 1 mg/ml guinea pig serum albumin (GSA) and 0.1% heparin, with or without FGF, in the left ear and were exposed to intense sound (115 dB 4 kHz octave band noise, 5 h) 4 days following surgery. Group I (control) received AP alone. Groups II, III, and IV received FGF-2 (Human recombinant, Sigma, 10 µg/ml), FGF-1 (Human recombinant, Sigma, 10 µg/ml), and FGF-1 (1 µg/ml), respectively, each dissolved in AP. Two animals, one in group II and the other in group IV, developed middle ear effusions after surgery, and were excluded from the study before noise exposure. Thus, total of 22 animals were included for data analysis.
The experimental protocol was approved by the University Committee for the Use and Care of Animals at the University of Michigan and conforms to the NIH Guidelines for the Care and Use of Laboratory Animals.
For cochlear infusion, a fine-tipped cannula was prepared as previously described (Yamasoba and Dolan, 1997, 1998). Before implantation, each cannula was flushed with AP containing 1 mg/ml GSA and 0.1% heparin. Each cannula was then filled with AP with or without FGF. After receiving, FGFs were kept in freezer until needed for surgery; FGFs were dissolved in AP immediately before filling into an osmotic pump. Heparin (0.1%) was added to AP because FGF-1 and FGF-2 are considered more stable in a complex with heparin, and heparin participates directly in the binding of FGFs to their receptors (see Basilico and Moscatelli, 1992). Biological activity of FGFs is greatly potentiated by the addition of heparin at a concentration ranging from 0.01 to 1 % (Mueller et al., 1989). Moreover, Wen et al. (1995) have reported that FGF-2 co-infused via an osmotic pump with 0.1 % heparin, but not FGF-2 alone, rescues ischemic hippocampal neurons, in vivo. Heparin was added to AP not only with FGFs but also without FGFs, since the effect of 0.1 % heparin on noise-induced cochlear damage is not known. GSA was added in AP since it is recommended by the producer (see a catalogue from Promega Corporation, Madison, Wisconsin, USA) for reducing loss of FGFs in dilute solution.
Animals were anaesthetized with xylazine (10 mg/kg, i.m.) and ketamine (40 mg/kg, i.m.); local infusion of 1 % lidocaine was used as necessary. Chloramphenicol sodium succinate (50 mg/kg, i.m.) was injected prophylacticly. Under aseptic conditions, the left bulla was exposed from the occipitolateral direction so that the basal turn and the round window were clearly visible. A small hole was opened in the bony wall of the basal turn lateral to the round window, allowing access to the scala tympani. The tip of the cannula, pre-filled with AP with or without FGF, was inserted into the hole. The cannula was secured to the bulla with tissue adhesive and dental acrylic cement, then looped around a stainless steel screw placed in the skull and secured to it with methyl methacrylate. The osmotic pump (model 2002, 200 µl, 0.5µl/h for 14 days; Alza Corp.) filled with AP with or without FGF was attached to the distal end of the cannula. A subcutaneous pocket was formed between the scapulae to accommodate the pump, which had been placed in a 38°C, 0.9% saline bath for 4 h, allowing it to be operable immediately upon implantation.
Intense sound (115 dB SPL 4 kHz octave band noise, 5 hours) was generated within a lighted, ventilated sound exposure chamber and two separately caged animals were noise-exposed at a time. Animal had access to food and water throughout the exposure. The sound chamber was fitted with speakers (JBL Inc., Northridge, CA., Model 2385A) driven by a noise generator (RANE Corp., Mukilteo, WA., GE 60 graphic equalizer) and power amplifier. The sound levels were calibrated and measured (Briiel & Kjaer microphone, Bruel & Kjaer modular precision sound level meter Type 2231) at multiple locations within the sound chamber to ensure stimulus uniformity, using a fast Fourier transform network analyser with a linear scale. The power analysis of the noise used has been shown elsewhere (Yamasoba et al. 1999).
Auditory brainstem measurement
Auditory evoked brainstem responses (ABR) were measured for each ear of each animal before and 3 days after surgery (1 day before noise exposure) and 10 days after noise exposure. Animals were anaesthetised with a mixture of xylazine (10 mg/kg) and ketamine (40 mg/kg) given intramuscularly prior to ABR measurement. A reference electrode was placed subcutaneously below the test ear and a needle electrode (before surgery) or a screw implanted at the vertex (after surgery) was used as an active electrode. A ground electrode was positioned below the contralateral ear. The sound stimulus consisted of a 15 ms tone burst, with a rise-fall time of 1 ms at frequencies of 2, 4, 8, 12, 16, and 20 kHz. The sound intensity was decreased in 5 dB intervals near threshold. One thousand twenty-four tone presentations given at the rate of 12.5/s were averaged using a microcomputer and custom software to obtain a waveform. Hearing threshold was defined as the lowest stimulus intensity that produced a reliable peak 3 or 4 in ABR waveforms. ABR thresholds obtained 1 day before noise exposure were used as baseline thresholds for estimating the noiseinduced threshold shifts.
Animals were sacrificed 2 weeks after noise exposure. Before euthanasia, the placement of the catheter tip and the condition of middle ear spaces were examined by ventral approach under deep anaesthesia with a mixture of xylazine and ketamine. We also examined the connection between the pump and the catheter and the remaining amount of solution. Then, animals were decapitated, the temporal bones immediately removed, and the perilymphatic spaces perfused with 2% paraformaldehyde in phosphate buffer and kept in solution for 1 h. After permeabilization with 0.3 % Triton X-100 for 5 min, whole mounts of the organ of Corti were stained for actin with rhodamine phalloidin for 40 min and examined as surface preparations. The slide preparations were observed under fluorescence microscopy, and missing and present IHCs and OHCs in 0.19 mm segments of sensory epithelium were counted from apex to base. Missing hair cells were recognized as dark, empty sites (after careful focus through the tissue) and/or typical phalangeal scar formation of the supporting cells (Raphael and Altschuler, 1991). For IHCs and first, second, and third OHC rows, the percentage of hair cell loss was calculated at each observation point in each animal, and the percentage at each 0.19 mm segment was plotted to create individual and averaged cytocochleograms. As demonstrated in Results section, with the noise exposure we used, little hair cell damage was found in the apical and third cochlear turns, between 0 and 7.6 mm from the apex. Therefore, only the organ of Corti between 7.6 mm and 19.0 mm from the apex was analysed for noise-induced hair cell damage.
Sigma Stat TM statistical software was used for statistical analysis. The thresholds before noise exposure and threshold shifts following noise exposure at each frequency, as well as the percentages of missing OHCs between 7.6 mm and 19.0 mm from the apex, were compared between the treated and untreated ears in each group using Student's-t test. They were also compared in the treated and untreated ears among groups I through IV using one-way analysis of variance (ANOVA).
| Results|| |
Hearing thresholds before noise exposure were essentially equivalent in all ears [Figure - 1]. No significant threshold differences were found at any frequencies between the treated and untreated ears in each group or in either treated and untreated ears among groups. At euthanasia, no displacement of the catheter from the cochlea or the pump was observed and the pump was virtually empty in all animals, indicating successful delivery of solution by the pump and catheter. No middle ear abnormalities, such as mucosal thickening and granulation, were observed.
[Figure - 2] shows the threshold shifts 10 days following noise exposure, observed across the tested frequencies in each of the groups. Consistent with the spectrum of the exposure stimulus, threshold shifts were smallest at 2 kHz and greatest at 4 and 8 kHz, with gradual reduction toward higher frequencies, in all groups except for treated ears in group II, which showed equivalent threshold shifts at higher frequencies. No significant differences of threshold shifts were observed between the treated and untreated ears in any group or in either treated and untreated ears across groups. Although the treated ears in group II showed somewhat greater shifts at the highest three frequencies compared to the untreated ears in group II and treated ears in other groups, these differences were not statistically significant.
[Figure - 3] shows the average percent loss of rows 1, 2, and 3 OHCs in treated and untreated ears in each group. Damage to the OHCs was greatest in the basal half of the second turn and apical half of the basal turn in all groups. Few hair cell lesions were observed in the third or apical turns. In all groups, row 1 OHCs showed the most damage, followed by row 2 OHCs, while the IHCs were well preserved. [Figure - 4] shows the extent of mean loss of OHCs of rows 1, 2, and 3, between 7.6 mm and 19.0 mm from the apex, in treated and untreated ears. Although the extent of hair cell loss varied somewhat among animals in each group, the tendency was similar among groups. In any OHC row, no significant difference in the extent of hair cell loss was observed between the treated and untreated ears in any group or in the treated versus untreated ears across groups.
| Discussion|| |
This study demonstrates that threshold shifts and hair cell damage were identical between the ears treated with FGF-1or FGF-2 and those treated with AP, as well between the treated and untreated ears in all treatment groups. These results indicate that neither FGF-1 nor FGF-2 exogenously applied to the guinea pig cochlea at the concentration used attenuates threshold shifts or hair cell damage due to intense sound exposure.
The negative findings may reflect the particular noise exposure we used. It is possible that protective effects by these FGF members might have occurred with a different type, intensity, and/or duration of noise. However, using the same exposure paradigm in previous studies, we found that pre-treatments with GDNF, NT-3, an iron chelator (deferoxamine mesylate), and a hydroxyl radical scavenger (mannitol) provided statistically significant attenuation of hair cell damage from acoustic trauma (Yamasoba et al., 1999; Shoji et al., 2000a, 2000b). The negative effects in the present investigation could also reflect an inappropriate delivery or concentration of FGFs. However, the delivery system has been demonstrated effective with GDNF (Shoji et al., 2000a; Yamasoba et al., 1999), BDNF, and NT-3 (Ernfors et al., 1996; Miller et al., 1997; Shoji et al., 2000b) in protection of hair cells from noise or drugs, or rescue and regrowth of auditory nerve fibres following hair cell destruction. Importantly we have observed that FGF-1, with or without BDNF, induces biological activity in the auditory nerve (cell survival and dendritic growth), when delivered in an identical manner in a similar study (Cho et al, 1998). Moreover, it has been demonstrated that FGF-2 retains its biological activity at 37 °C for periods of up to 4 weeks when dispensed from a sequestered source (Edelman et al., 1991) and that FGF-2 delivered with an osmotic pump for 2 or 4 weeks can induce biological effects, such as myocrdial angiogenesis (Landau et al., 1995), rescue of neuronal loss (Wen et al., 1995), and increased collateral blood flow (Yang et al., 1996). Thus, there is substantial evidence that FGFs delivered with an osmotic pump can be biologically active at least for 2 weeks.
It is difficult to specify effective or optimal concentrations of FGFs to protect the hair cells in vivo. It has been shown that GDNF delivered to the cochlea with an osmotic pump can afford hair cell protection at 10 and 100 ng/ml, but not at 1 ng/ml, and at 1 µg/ml, it may increase damage (Shoji et al., 1998) and that GDNF delivered to the cochlea with an osmotic pump can enhance spiral ganglion cell survival after drug-induced hair cell death at 10 and 50 ng/ml (Altschuler et al., 1997). It has been shown that GDNF in vitro rescues rat superior cervical ganglion neurons in a dose-dependent manner, with half-maximal response at 4-5 ng/ml, while it provides maximal survival of dorsal and nodose ganglion neurons in chick embryo at 10 ng/ml (Trupp et al., 1995). Thus, the effective concentrations, in vitro and in vivo using an osmotic pump, are considered to be generally comparable. If there is a difference, it is that optimal concentration is somewhat higher in vivo than in vitro. Taking into account these in vivo and in vitro findings, it seems appropriate to use trophic/growth factors in vivo with an osmotic pump at the concentrations equal to and up to 10 times higher than, those showing an optimal effect in vitro. FGF-2 has been shown to effectively enhance survival of the auditory neurons, hair cells, and utricular epithelial cells in vitro at concentrations of 100-1000 ng/ml (Lefebvre et al., 1991), 500 ng/ml (Low et al., 1996), and 1-100 ng/ml (Zheng et al., 1997), respectively. Thus, the concentrations of FGFs used in this study (1 and 10 µg/ml delivered at a rate of 0.5 µl/h) probably were appropriate to efficiently elicit a biological response in the inner ear. Since the molecular weight of GDNF, FGF-1, and FGF-2 is 30 kiloDalton (kDa), 15.5 kDa, and 17.5 kDa, respectively, even when compared with molar concentration, the concentrations of FGF used are likely to be biologically appropriate. Therefore, based on our data, we consider it unlikely that FGF-1 and FGF-2 act as hair cell protectors in the mature cochlea in vivo. We do note that it is possible that other FGF members may provide hair cell protection. If this is the case, the most likely candidates are considered FGF-4, FGF-8, and FGF-9. These members, as well as FGF-1 and FGF-2, bind with high affinity to FGFR3 (Ornitz and Leder, 1992; Hecht et al., 1995; Macarthur et al., 1995; Ornitz et al., 1996), which has been identified in the organ of Corti (Peters et al., 1993; Pirvola et al., 1995).
Several factors may account for the lack of an important role of FGF-1 and FGF-2 in hair cell protection in the mature cochlea in vivo. First, it has been reported that exposure to noise does not change the level of FGF-1 in the rat cochlea (Pirvola et al., 1995), indicating the absence of significant noise-induced upregulation of endogenous FGF-1 synthesis in the cochlea. This suggests that increases in FGF-1 levels may not provide benefit to the damaged cochlea. We do not know, however, if synthesis of other FGF members is upregulated by noise exposure. Second, targeted disruption of FGFR3 in mice has been reported to result in apparently arrested development of morphologically distinct pillar cells, without affecting IHC and OHC development (Colvin et al., 1996). Since only FGFR3 has been detected in both the developing and the mature mammalian cochlea, FGF members might be important in the development and maintenance of supporting cells but may not be essential for growth or trophic support of the mammalian cochlear hair cells. Third, prevention of neuronal cells requires binding of certain growth factors to specific growth factor receptors for initiating signal processing. For example, BDNF, NT-3, and NT-4/5 can enhance survival of the spiral ganglion cells of postnatal rats (Zheng et al., 1995), where high affinity binding tyrosine kinase receptors, trk B and trk C, are present (Ylikoski et al., 1993). In contrast, FGF-2 does not enhance survival of the spiral ganglion cells (Zheng et al., 1995), where no FGFRs have been detected (Pirvola et al., 1995). Similarly, degeneration of the auditory neurons, induced by drugs such as cisplatin in organotypic cultures of postnatal rat cochlear explants, can be prevented by BDNF, NT-3, and NT-4/5, but not by FGF-2, FGF-5, or FGF-7 (Zheng and Gao, 1996), and hair cells can be protected from neomycin-induced injury by TGF-a (Staecker et al., 1997) whose receptor, epithelial growth factor receptor, is present in OHC stereociliary bundles (Lee and Cotanche, 1996). FGFR3 has been found in the hair cells in developing cochlea in both mammals and avians, but it is present only in supporting cells in the mature chicken cochlea (Bermingham-McDonogh et al., 1998) and nearly exclusively in the supporting cells in the mature rat cochlea (Pirvola et al., 1995). In addition, an in vitro study using cultured cochlea tissues from 3-5 day old rats, which are still developing, showed a protective effect of FGF-2 against neomycin ototoxicity (Low et al., 1996).
It has been shown that damaging the chicken basilar papilla by noise exposure results in upregulation of FGF receptors only in supporting cells (Lee and Cotanche, 1996). Zheng et al. (1997) have shown that FGF-4, FGF-6, FGF-7 and, most potently, FGF-2 stimulate cultured utricular supporting cells from postnatal day 4-5 rats and that proliferation is inhibited by neutralizing antibodies against FGF-2. Pirvola et al. (1995) have demonstrated that noise-induced upregulation of FGFR3 is seen in supporting cells in the mature rat cochlea. These results suggest that FGF members are candidate molecules for regulating proliferation of inner ear supporting cells. It is well known that the supporting cells play a role in scar formation during hair cell degeneration in the mature mammalian cochlea. It has been reported that, when guinea pigs are exposed to intense sound, two supporting cells form a scar for a given hair cell, by expanding and invading the sub-apical region of the dying hair cell (Raphael and Altschuler, 1991). Thus, it is likely that the binding of certain FGF members to FGFR3 initiates or stimulates expansion of supporting cells for scar formation following noise damage; FGFs may act to repair the organ of Corti following insults.
Given the findings that so many factors offer varying degrees of protection and rescue of inner ear sensory and neural elements from trauma, it is important to observe that one specific factor, FGF, with its receptors virtually restricted to non-sensorineural elements, does not provide hair cell protection.
| Acknowledgments|| |
The authors thank Ms. Diane M. Prieskorn and Ms. Alice Mitchell for valuable discussion and technical help, and Ms. J. Nadine Brown for editorial help. This work is supported by NIH research grant DC00105 and DC 00124.
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Josef M Miller
Kresge Hearing Research Institute, 1301 East Ann Street, Ann Arbor, Michigan 48109-0506, USA
Source of Support: None, Conflict of Interest: None
[Figure - 1], [Figure - 2], [Figure - 3], [Figure - 4]